The Enzymes: Nucleic Acids, Vol. 15, Pt. B

The Enzymes VOLUME XV NUCLEIC ACIDS Part B Third Edition CONTRIBUTORS SIDNEY ALTMAN L. ANDREW BALL EDMUND W. BENZ, J...

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The Enzymes VOLUME XV

NUCLEIC ACIDS Part B Third Edition

CONTRIBUTORS SIDNEY ALTMAN L. ANDREW BALL EDMUND W. BENZ, JR. PETER BLACKBURN THOMAS BLUMENTHAL RICHARD R. BURGESS MICHAEL J. CHAMBERLIN MURRAY P. DEUTSCHER JOHN J. DUNN MARY EDMONDS MICHAEL J. ENGLER RICHARD I. GUMPORT JERARD HURWITZ LARRY K. KLINE

RYSZARD KOLE MARTIN K. LEWIS U. Z. LITTAUER STANFORD MOORE DANNY REINBERG CHARLES C. RICHARDSON T. RYAN D. SCHLESSINGER V. SHEN STEWART SHUMAN DIETER SOLL H. SOREQ KENJI TAKAHASHI OLKE C. UHLENBECK

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

THE ENZYMES Edited by PAUL D. BOYER Depurtment of Chemistry anti Molecular Bio1og.v Institute Uiiiversiry of Cdiforaia Los Angeles, Californiii

Volume XV NUCLEIC ACIDS Part B

THIRD EDITION

1982

ACADEMIC PRESS A Subsidiury of’ Harcourt Bruce Jovrinovicl~,Publishers

New York London

Paris San Diego San Francisco Sko Paul0 Sydney Tokyo Toronto

COPYRIGHT @ 1982, BY ACADEMIC PRESS, INC. ALL RIGHTS RESERVED. N O 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 I N WRlTING FROM T H E PUBLISHER.

ACADEMIC PRESS, INC. 111 Fifth Avenue, New

York,New York 10003

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

Library o f Congress Cafaloging i n Publication Data Main entry under t i t l e :

The Enzymes. Includes bibliographical references.

CDNTWTS: v. 1. Structure and control.--v.

2. Kine-

t i c s and mechanism.--v. 3. Hydrolysis: peptide bonds.

--Letc.1--v. 15 W l e i c acids, pt. 8. 1. Enzymes. I. Boyer, Paul D., ed.

mes.

MIU5 B891el ~~-

QP601. €523

~

574.19'25

ISW 0-12-122715-4 (v.15)

[DNLM:

75-117107

AACRl

PRINTED I N THE UNITED STATES O F AMERICA

82 83 84 85

9 8 7 6 5 4 3 2 1

1. Enzy-

Contents List of Contributors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Prrfoce . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

xi xiii

Section 1. Ligases

1 . DNA Ligases

MICHAEL J . ENGLER A N D CHARLES C . RICHARDSON I . Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . I1 . Isolation and Physical Properties . . . . . . . . . . . . . . . . . . . . 111. Catalytic Properties . . . . . . . . . . . . . . . . . . . . . . . . . . IV. Role of DNA Ligases it1 Viva . . . . . . . . . . . . . . . . . . . . . . V. Research Applications . . . . . . . . . . . . . . . . . . . . . . . . .

3 5 10 21 26

2 . 14 RNA Ligase

OLKEC . UHLENBECK A N D RICHARD I . GUMPORT I . Introduction . . . . . . I1 . Purification and Properties 111. Reactions Catalyzed . . . IV. Biological Role . . . . . V. Applications . . . . . .

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

31 33 35 47 52

Section I1 . RNA Polymerares and Related Enzymes

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3 Bacterial DNA-Dependent RNA Polymemses

MICHAEL J . CHAMBERLIN I . Background

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

I1 . Molecular Properties . . . . . . . . . . . . . . . . . . . . . . . . . 111. Catalytic Properties . . . . . . . . . . . . . . . . . . . . . . . . . .

61 64 82

CONTENTS

Vi

.

4 Bacteriophage DNA-Dependent RNA Polymerases

M . CHAMBERLIN AND T. RYAN I . Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 87 89 I1. T7-Like RNA Polymerases . . . . . . . . . . . . . . . . . . . . . . . 111 Other Bacteriophage RNA Polymerases . . . . . . . . . . . . . . . . . 105

.

.

5 Eukaryotic RNA Polymeraser

MARTINK . LEWISA N D RICHARDR . BURGESS I . Introduction

. . . . . . . . . . . 111. Subunit Structures . . IV. Subunit Functions . . . V. Stimulatory Factors . . 11. Purification

VI . VII . VIII . IX .

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DNA Binding and Catalytic Properties of Purified RNA Polymerases Inhibitors . . . . . . . . . . . . . . . . . . . . . . . . . . Eukaryotic Transcription Extract Systems . . . . . . . . . . . . Organelle- and Viral-Coded RNA Polymerases . . . . . . . . . .

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110

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111 117 128 137 138 145

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147 150

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6 Priming Enzymes

EDMUND w. BENZ.J R . , DANNYREINBERG. JERARD HURWITZ

AND

I . Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . I1. d17irG Gene Product from Eschrrichitr coli . . . . . . . . . . . . . . . 111. Multiple Pathways for t h G Priming in SS to RF DNA Replication . . . . IV. Studies on the Specificity of tlmrG-Template Interactions . . . . . . . . V. Priming by RNA Polymerases . . . . . . . . . . . . . . . . . . . . . VI . Priming on Double-Stranded DNA . . . . . . . . . . . . . . . . . . VII . Phage-Encoded Priming Enzymes . . . . . . . . . . . . . . . . . . . . VIII . Priming in Eukaryotic Systems . . . . . . . . . . . . . . . . . . . . .

155

. 156 . 160 . .

165 166 169 174 178

.

7 tRNA Nucleotidyltransferase

MURRAYP. DEUTSCHER I. I1 . 111. IV . V.

Introduction . . . . . . . . . . Purification and Structural Studies Catalytic Properties . . . . . . . Biological Role . . . . . . . . . Research Applications . . . . . .

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183

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213 215

.

8 Poly(A) Adding Enzymes

MARYEDMONDS I . Introduction and Perspective . . . . . . . . . . . . . . . . I1. Purification and Properties . . . . . . . . . . . . . . . . .

218 219

vii

CONTENTS 111. Multiple Poly(A) Polymerases . . . . . . . . . . . . IV. Properties of Poly(A) Polymerase Proteins . . . . . . V. Assay . . . . . . . . . . . . . . . . . . . . . . . VI . Stoichiometry . . . . . . . . . . . . . . . . . . . VII . Substrates . . . . . . . . . . . . . . . . . . . . . VIII . Primer Requirement . . . . . . . . . . . . . . . . . IX . Ion Requirements . . . . . . . . . . . . . . . . . . X . Inhibitors . . . . . . . . . . . . . . . . . . . . . XI . Kinetics and Reaction Mechanism . . . . . . . . . XI1. Biological Role . . . . . . . . . . . . . . . . . . . XI11 . Regulation of Poly(A) Polymerases . . . . . . . . . XIV. Research Applications . . . . . . . . . . . . . . . .

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223 225 226 227 227 . . . . . . . . 227 . . . . . . . . 229 . . . . . . . . 230 . . . . . . . . . 233 . . . . . . . . 235 . . . . . . . . . 239 . . . . . . . . 243

.

9 Capping Enzyme

STEWART SHUMAN A N D JERARD HURWITZ I . Introduction . . . . . . . . . . . . . . . . . . . . . It . Vaccinia Virus Capping Enzymes . . . . . . . . . . . . 111. HeLa Cell Capping Enzymes . . . . . . . . . . . . . . IV. Capping Enzyme from Rat Liver Nuclei . . . . . . . . V. Role of the Capping Enzyme System iu V i l v . . . . . . VI . Research Applications of Vaccinia Virus Capping Enzyme

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

246 249 256 . . . . . . . . . 257 . . . . . . . . . 258 . . . . . . . . . 264

. QP Replicase

10

THOMAS BLUMENTHAL I . Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11. Purification and Properties . . . . . . . . . . . . . . . . . . . . . . . 111. Catalytic Properties . . . . . . . . . . . . . . . . . . . . . . . . . .

267 269 273

.

1 1 2'.5 '.Oligoadenylate Synthetase

L . ANDREWBALL I . Introduction . . . . . . I1. Purification and Properties I11 . Reactions Catalyzed . . . IV. Biological Role . . . . .

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281 284 290 304

.

Section 111 RNA Nucleases and Related Enzymes

.

12 Pancreatic Ribonuclease

PETERBLACKBURN A N D STANFORD MOORE I . Introduction

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

11. Preparation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

317 318

...

CONTENTS

Vlll

I11. Chemical Properties . . . . . . . . . . . . . . . . . . . . . . . . . . IV. Physical Properties . . . . . . . . . . . . . . . . . . . . . . . . . . V. Species Variations . . . . . . . . . . . . . . . . . . . . . . . . . . . VI . Bovine Seminal Plasma RNase . . . . . . . . . . . . . . . . . . . . . VII . Cytoplasmic RNase Inhibitor . . . . . . . . . . . . . . . . . . . . . . VIII . Catalytic Properties . . . . . . . . . . . . . . . . . . . . . . . . . . IX . Research Applications . . . . . . . . . . . . . . . . . . . . . . . . .

320 364 397 411 416 424 433

.

13 Ribonuclease TI

KENJITAKAHASHI A N D STANFORD MOORE I . Introduction

. . . . . . . . . . . . . . . . . . . . . . 111. Reactions Catalyzed . . . . . . . . . . . . . . IV. Research Applications . . . . . . . . . . . . . V. Other Guanine-Specific RNases . . . . . . . . . I1. Purification and Chemical Properties

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435 . . . . . . . . . . . . . . . . . . . 436 447 . . . . . . . . . 463 . . . . . . . . . 465 . . . . . . . . .

.

14 tRNA Processing Enzymes from Ercherichia coli

RYSZARDKOLEA N D SIDNEY ALTMAN 1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 470 I1 . Ribonuclease P . . . . . . . . . . . . . . . . . . . . . . . . . . . . 471 111. Ribonuclease 111 . . . . . . . . . . . . . . . . . . . . . . . . . . . 476 IV. Ribonuclease Pt and Ribonuclease 0 . . . . . . . . . . . . . . . . . . 477 V. Ribonuclease D . . . . . . . . . . . . . . . . . . . . . . . . . . . . 479 VI . Other Nucleases . . . . . . . . . . . . . . . . . . . . . . . . . . . 482 VII . Concluding Remarks . . . . . . . . . . . . . . . . . . . . . . . . . 483

.

15 Ribonuclease 111

JOHN J . DUNN 1. Introduction . . . . . . . . . . I1 . Purification . . . . . . . . . . . 111. Structure . . . . . . . . . . . . IV. Cleavage of Double-Stranded R N A V. Cleavage of Single-Stranded RNA VI . Related Enzymes from Eukaryotes

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485 . . . 486 . . . 487 . . . . . . . 489

. . . . . . . . . . . . . . . . . . . 490 . . . . . . . . . . . . . . . . . . . 497

.

16 RNases I. II. and IV of Escherichia coli

V. SHENA N D D . SCHLESSINGER I . Introduction . . . . . . . . . . . . . . . . I1. RNase I of Esclierichicr coli . . . . . . . . . 111. RNase 11 of Eschrrichirr coli . . . . . . . . . IV. RNase IV of Esrhrricliin c.o/~. . . . . . . . .

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501 503 506 512

ix

C0N TE N TS

.

17 Polynucleotide Photphorylase

U . Z . LITTAUER A N D H . SOREQ I . Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

518

I1. Properties . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

I11. The Reactions Catalyzed . . . . . . . . . . . . . . . . . . . . . . . . IV. Attributed Physiological Functions . . . . . . . . . . . . . . . . . V . Research Applications . . . . . . . . . . . . . . . . . . . . . . . . .

. .

519 530 537 539

.

Section IV RNA Modification

18. RNA Methylation

DIETER SOLL A N D LARRY K . KLINE I . Introduction . . . . . . . . . . . . . . . . I1. Structures and General Assay Procedure . . . I11 . Specific tRNA Methyltransferase Enzymes . IV. Conclusion . . . . . . . . . . . . . . . . .

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. . . . . . . . . 557 . . . . . . . . . . 558 . . . . . . . . . . 559 . . . . . . . . . 566

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19 Nucleotide Modification in RNA

LARRY K . KLINEA N D DIETER SOLL . . . . . . . . . . . . . . . . . . . . . . . . . . . . . I . Introduction

I1 . Modification of Uridine . . . . . . . . . . . . . . . . . . . . . . . . I11. Modification of Cytidine . . . . . . . . . . . . . . . . . . . . . . . . IV . Modification of Adenosine . . . . . . . . . . . . . . . . . . . . . . . V. Modification of Guanosine . . . . . . . . . . . . . . . . . . . . . . . VI . Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Auihorltidex Subject Index

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567 568 574 575 578 582 583

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627

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647

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

SIDNEY ALTMAN (469), Department of Biology, Yale University, New Haven, Connecticut 065 10

L. ANDREW BALL (281), Biophysics Laboratory and Biochemistry Department, University of Wisconsin, Madison, Wisconsin 53706 EDMUND W. BENZ, JR. ( 1 5 3 , Department of Developmental Biology and Cancer, Albert Einstein College of Medicine of Yeshiva University, Bronx, New York 10461 PETER BLACKBURN (317), The Rockefeller University, New York, New York 10021 THOMAS BLUMENTHAL (267), Department of Biology, Indiana University, Bloomington, Indiana 47401 RICHARD R. BURGESS (109), McArdle Laboratory for Cancer Research, Madison, Wisconsin 53706 MICHAEL J. CHAMBERLIN (61, 87), Department of Biochemistry, University of California, Berkeley, California 94720 MURRAY P. DEUTSCHER (183), Department of Biochemistry, University of Connecticut Health Center, Farmington, Connecticut 06032 JOHN J . DUNN (485), Department of Biology, Brookhaven National Laboratory, Upton, New York 11973 MARY EDMONDS (217), Department of Biological Sciences, University of Pittsburgh , Pittsburgh, Pennsylvania 15260 MICHAEL J. ENGLER (3), Department of Biological Chemistry, Harvard Medical School, Boston, Massachusetts 021 15 RICHARD I. GUMPORT (31), Department of Biochemistry and School of Basic Medical Sciences, University of Illinois, Urbana, Illinois 61801 JERARD HURWITZ (155, 245), Department of Developmental Biology and Cancer, Albert Einstein College of Medicine of Yeshiva University, Bronx, New York 10461 xi

xii

LIST OF CONTRIBUTORS

LARRY K. KLINE (557, 567), Department of Biological Sciences, State University College, Brockport, New York 14420 RYSZARD KOLE" (469), Department of Biology, Yale University, New Haven, Connecticut 06510 MARTIN K. LEWIS (109), McArdle Laboratory for Cancer Research, Madison, Wisconsin 53706 U. Z. LITTAUER (517), Department of Neurobiology, The Weizmann Institute of Science, Rehovot,Israel 76100 STANFORD MOORE (317, New York, New York 10021

4 3 9 , The

Rockefeller

University,

DANNY REINBERG (155), Department of Developmental Biology and Cancer, Albert Einstein College of Medicine of Yeshiva University, Bronx, New York 10461 CHARLES C. RICHARDSON (3), Department of Biological Chemistry, Harvard Medical School, Boston, Massachusetts 021 15 T. RYAN (87), Department of Biochemistry, University of California, Berkeley, California 94720 D. SCHLESSINGER (501), Department of Microbiology and Immunology, Washington University School of Medicine, St. Louis, Missouri 631 10

V. SHEN (501), Department of Microbiology and Immunology, Washington University School of Medicine, St. Louis, Missouri 631 10 STEWART SHUMAN (245), Department of Developmental Biology and Cancer, Albert Einstein College of Medicine of Yeshiva University, Bronx, New York 10461 DIETER SOLL (557, 567), Department of Molecular Biophysics and Biochemistry, Yale University, New Haven, Connecticut 0651 1 H. SOREQ (5 17), Department of Neurobiology , The Weizmann Institute of Science, Rehovot JIsrael 76100 KENJI TAKAHASHI (439, Primate Research Institute, Kyoto University, Inuyama Aichi 484, Japan

OLKE C. UHLENBECK (3 l), Department of Biochemistry, University of Illinois, Urbana, Illinois 61801 * Present address: Department of Human Genetics, Yale University School of Medicine, New Haven, Connecticut 065 10.

Pvefa ce This is the second of two volumes that cover nucleic acid enzymology. The striking advances and crucial importance of this rapidly developing area made review at this time imperative, even though there is still much to be learned about the molecular enzymology involved. With the exception of the chapter on DNA ligases, this volume centers on enzymes involved in the formation, degradation, and modification of RNA. Present information is extensive, and readers will likely recognize an indebtedness to the excellent authors for their authoritative coverage. It is a distinct pleasure to record appreciation for the guidance provided by the Advisory Board members of this and the preceding volume. Their exceptional professional competence and breadth of knowledge made essential contributions to the excellence of the volumes. This volume records a milestone along the path of one of the most vital and revealing areas of biological research of all times. Paul D. Boyer

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Section I

Ligases

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DNA Ligases MICHAEL J. ENGLER

CHARLES C. RICHARDSON

. . A. Assays . . . . . . . . . . . B. Purification . . . . . . . . . C. Physical Properties . . . . . . 111. Catalytic Properties . . . . . . . I. Introduction

. . .

. . . . .

11. Isolation and Physical Properties

.

,

. .

,

,

,

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

. . . . . . . . . . . . . . . . . . A. Mechanism of Phosphodiester Formation . . B. Reversal of the Ligase Reaction . . . . . . C. Formation of Phosphodiesters at Nicks . . . D. Blunt-End Joining . . . . . . . . . . . . IV. Role of DNA Ligases in Vivo . . . . . . . . , A. Phage-Induced DNALigases . . , . . . . B. Escherichiu coli DNA Ligase . . . . . . .

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. . . . . . . . . . . . . C. Physiological Requirement for Bacterial and Phage DNA Ligases . D. Yeast DNA Ligase . . . , , , , , . . . . . . . . , , . . . . V. Research Applications . . . . . . . . . . . . . . . . . . . . . .

I.

. .

3 5 5 8 9 10 10 14

IS 19 21 21 23 25 26 26

Introduction

DNA ligases are enzymes that catalyze the formation of a phosphodiester linkage between DNA chains. Condensation of the 5’ -phosphoryl group with the adjacent 3’-hydroxyl group is coupled with the hydrolysis of a pyrophosphate moiety of NAD (bacterial enzyme) or ATP (phage or eukaryotic enzymes). Prior to the discovery of DNA ligase several experimental observations suggested the existence of an enzyme that could catalyze the covalent 3 THE ENZYMES. VOL.X V Copyright @ 1982 by Academic Press. Inc. AU rights of reproduction in any form reserved. ISBN 0-12-122715-4

4

MICHAEL J. ENGLER AND CHARLES C. RICHARDSON

joining of polynucleotides. Studies on bacteriophage had shown that genetic recombination involved the breaking and rejoining of polynucleotide strands ( I , t ) , and physical and genetic studies on the repair of ultraviolet-irradiated DNA suggested a terminal step requiring strand joining ( 3 ) . A more specific demonstration of a joining activity came from the observation that the linear DNA molecule of phage lambda was converted to a covalently closed duplex circle shortly after injection into its host, Escherichiu coli ( 4 , 5 ) . An added impetus to search for such an enzyme activity was the growing realization that a novel mechanism might exist for the overall 3‘-5’ directional growth of a DNA strand (lagging strand) during replication [see Ref. ( 6 ) ] .One postulated mechanism, the synthesis of small chromosomal units on both strands in an antiparallel manner (7), required a subsequent joining event to yield DNA of high molecular weight. It was not until later, concurrent with the discovery of DNA ligase, that direct evidence was obtained to support a discontinuous mechanism for DNA replication (8). DNA ligase was first identified in extracts of uninfected and Tphage-infected E. coli in 1967 ( 9 4 4 ) . Initially, the major feature that distinguished the bacterial from phage-induced enzymes was their cofactor specificity; E. coli ligase requires NAD, whereas the phage enzyme requires ATP. Another bacterial ligase, isolated from Bacillus subtilis, also requires NAD (15). DNA ligases have been found in a large variety of eukaryotic cells [see review, Ref. (16)l; all have a requirement for ATP. Of the DNA ligases that have been described, the E. coli and phage T4-induced enzymes have been most thoroughly characterized. There1. Meselson, M., and Weigle, J. J. (1961). PNAS 47, 857. 2. Anraku, N . , and Tomizawa, J . (1965). JMB 11, 501. 3. Boyce, R . P., and Howard-Flanders, P. (1964). PNAS 51, 293. 4. Young, E. T., and Sinsheimer, R. L. (1964). JMB 10, 562. 5. Bode, V. C., and Kaiser, A. D. (1965). J M B 14, 399. 6. Sueoka, N . (1967). In “Molecular Genetics” (J. H. Taylor, ed.), Part 11, p. 1. Academic Press, New York. 7. Nagata, T. (1963). PNAS 49, 551. 8. Okazaki, R., Okazaki, T., Sakabe, K.,Sugimoto, K.,and Sugino, A. (1968). PNAS 59, 598. 9. Gellert, M. (1967). PNAS 57, 148. 10. Weiss, B . , and Richardson, C. C. (1967). PNAS 57, 1021. 11. Olivera, B . M., and Lehman, I. R. (1967). PNAS 57, 1426. 12. Gefter, M. L . , Becker, A . , and Hunvitz, J. (1967). PNAS 58, 240. 13. Becker, A . , Lyn, G . , Gefter, M., and Hurwitz, J. (1967). PNAS 58, 1996. 14. Cozzarelli, N . R., Melechen, N. E., Jovin, T. M., and Kornberg, A. (1967). BBRC 28, 578. 15. Laipis, P. J., Olivera, B. M., and Ganesan, A. T. (1969). PNAS 62, 289. 16. Soderhall, S., and Lindahl, T. (1976). FEBS Lett. 67, 1 .

1 . DNA LIGASES

5

fore, this chapter focuses on these two enzymes and refers to studies on ligases from other sources only when they supplement, or differ from, those obtained with the E. cofi and T4 enzymes. This series (17) and other reviews (18, 19) have already covered earlier studies on DNA ligases. This chapter places major emphasis on (1) the purification and physical properties of the ligases; (2) the properties and substrate specificities of the reactions catalyzed by the enzyme, including the intermediates in the reactions; (3) the in vivo roles of DNA ligases; and (4) the research applications of the enzyme. II.

Isolation and Physical Properties

A. ASSAYS 1. Alteration of the Properties of Polynucleotide Chciins

DNA ligase activities have been measured by a number of procedures. Ligase activity was initially detected in extracts of E. cofi by measuring the conversion of hydrogen-bonded circles of phage A DNA to covalently bonded ones using a sedimentation assay (9). A more rapid assay, which also utilizes the cohesive ends of A DNA, involves the joining of radioactively labeled A DNA to cross-linked unlabeled A DNA (20). Although the cross-linked DNA will renature after treatment with alkali, the labeled DNA will not, unless it has become covalently attached to the crosslinked DNA. The single-stranded and native DNA reaction products are then quantitated by hydroxylapatite chromatography. Another assay ( 1 4 ) that measures the covalent joining of one duplex polymer to another makes use of a polynucleotide chain covalentIy linked to cellulose, thus permitting it to be isolated by sedimentation or filtration. By adding the appropriate complementary polymers to the celluloselinked polymer, a duplex substrate can be prepared with which to measure ligase activity. A rapid and convenient assay measures the conversion of 3H-labeled d(AT) copolymer to a form resistant to exonuclease I11 (21 1. In this reac17. Lehman, I . R. (1974). “The Enzymes,” 3rd ed., Vol. X , Chap. 8, p. 237. 18. Lehman, I. R. (1974). Science 186, 790. 19. Higgins, N. P., and Cozzarelli, N. R. (1979). “Methods in Enzymology,” Vol. 68, p. 50. 20. Zimmerman, S. B . , Little, J. W., Oshinsky, C. K., and Gellert, M. (1%7). PNAS 57, 1841. 21. Modrich, P., and Lehman, I. R. (1970).JBC 245, 3626.

6

MICHAEL J. ENGLER AND CHARLES C. RICHARDSON

tion DNA ligase catalyzes an intramolecular joining reaction with linear self-complementary d(AT) oligomers, leading to the formation of circular molecules (22). 2. Direct Measurement of Phosphadiester Bond Formation

A more direct type of assay for ligase activity measures the conversion of internally located 32P-labeled5’-phosphomonoesters to diesters, which are resistant to E. coli alkaline phosphatase. After limited digestion with pancreatic DNase, duplex DNA contains single-strand breaks bearing 5’-phosphoryl groups. All such phosphomonoesters are, removed by treatment with phosphatase at elevated temperatures, and the external and internal 5’-hydroxyl groups are then radioactively labeled by phosphorylation using [Y-~~PIATP and polynucleotide kinase (10, 2.3). If these 32P-labeled phosphomonoesters are incorporated into phosphodiester linkages in a ligase reaction, they are converted to a phosphatase-resistant form. A similar assay (11, 2 4 ) uses as substrate a double-stranded homopolymer pair consisting of multiple oligo(dT) units labeled with [ 5’3’P]phosphomonoester hydrogen-bonded to a long poly(dA) chain. A novel variation of this type of assay is the covalent joining of a [ 5 ’ 32P](dA-dT)oligomer to yield phosphatase-resistant radioactivity (22 ), a reaction dependent on the ability of poly(dA-dT) to form intramolecular circles. 3 . Detection of Biological Activity

Several biological assays for measuring ligase activity have also been described. Ligase will restore marker activity of transforming DNA that has been inactivated by the introduction of single-strand breaks with pancreatic DNase (15, 25). Similarly, ligase activity has been measured by following the restoration of biological activity in a transfection assay (26). In this case, phage DNA, previously inactivated by a single restriction enzyme cleavage, is repaired by covalent joining via the short cohesive ends generated by the restriction cut. 4. Measurement of

(I

Partial Reaction

More rapid assays, which do not require the preparation of a special DNA substrate, have been used to monitor the purification of DNA ligase. 22. 23. 24. 25. 26.

Olivera, B. M., Schemer, I . E., and Lehman, I. R. (1%8). J M B 36, 275. Weiss, B., Live, T. R., and Richardson, C. C. (1968). JBC 243, 4530. Olivera, B. M., and Lehman, I. R. (1968). J M B 36, 261. Bautz, E. K . F. (1967). BBRC 28, 641. Murray, N. E . , Bruce, S. A . , and Murray, K. (1979). J M B 132, 493.

1.

DNA LIGASES

7

These assays measure the first step in the ligase-catalyzed reaction. Both E. cofi ligase (27) and TCinduced ligase (28) can be assayed by measurement of the formation of the acid-precipitable ligase-AMP intermediate using NAD or ATP, respectively, radioactively labeled in the AMP moiety. T4 DNA ligase has been assayed during purification by measuring the exchange reaction between ATP and 32PPi(29). This assay measures the conversion of "PPi into a form that adsorbs to charcoal. In principle such an assay could also be used to measure the exchange reaction between NAD and NMN in the E. cofi DNA ligase reaction. 5. Other Assriys Although all of the assays discussed above have been used to monitor DNA ligase activity during purification, many other possibilities exist. Such assays are limited only by the expertise and ingenuity of the investigators. For example, the joining of restriction fragments and the reformation of covalently closed circular molecules, reactions catalyzed by DNA ligase, can be followed by such diverse techniques as pycnographic analysis, electron microscopy, and gel electrophoretic analysis. 6 . Choice of an Assay

Which assay should be used to measure DNA ligase activity during purification? For detecting normal amounts of ligase activity in extracts of cells, the most suitable assay procedures are probably those that most directly measure phosphodiester formation by the conversion of a phosphatase-sensitive 32P-labeled 5'-phosphomonoester to a phosphatase-resistant form. Equally satisfactory and sensitive is the conversion of linear poly(dA-dT) copolymer to an exonuclease I11 resistant form. Since extracts ofcells may contain other enzymes that catalyze an ATP-PP, exchange and NAD-NMN exchange, these assays cannot always be used in the early stages of purification. However, the purification procedure developed for the T4 DNA ligase is sufficiently reproducible in the early steps to permit postponing an assay of the enzyme until Step V (chromatography on DEAE-cellulose) when the exchange assay is reliable (29). When overproducing strains of cells are used as a source of enzyme any of the assays should provide a sufficiently reliable method to identify the peaks that contain ligase activity during column chromatography. 27. Zirnrnerrnan, S. B., and Oshinsky, C. K . (1969). JBC 244, 4689. 28. Knopf, K . W. (1977). EJB 73, 33. 29. Weiss, B . . Jacquemin-Sablon, A., Live, T. R . , Fareed, G . C., and Richardson, C. C. (1968). JBC 243, 4543.

8

MICHAEL J. ENGLER AND CHARLES C. RICHARDSON

B. PURIFICATION Of the DNA ligases that have been identified, only the E. coli, T4, and T7 DNA ligases have been purified to homogeneity. In addition, the two most widely used enzymes, theE. coli and T4 ligases, have been amplified in E. coli by cloning the respective ligase genes into a phage A vector. Although partial purification procedures have been described for DNA ligases from eukaryotes as well as other prokaryotes, only the purification of the E. coli and T4 enzymes are discussed here. 1. Escherichia coli DNA Ligase Escherichia coli DNA ligase has been purified approximately 1400-fold from E. coli LC81 (30),a mutant strain that contains four times the normal amount of ligase (31). TheE. coli DNA ligase purified by this procedure is physically homogeneous as judged by polyacrylamide gel electrophoresis in the presence of sodium dodecyl sulfate (30). Purification of the E. coli DNA ligase has been facilitated greatly by the construction of an E. coli strain carrying a hybrid A prophage that bears the ligase overproducing gene, lopll lig+ (32). A simple purification procedure (33) using this strain results in an approximately 40% yield of the homogeneous enzyme; 30 mg of ligase can be obtained from 120 g of cell paste.

2. T4 DNA Ligase The routine purification procedure for T4 DNA ligase from cells infected with wild-type T4 phage consists of seven steps, yielding a 1000fold purified enzyme (29). By using T4 phage mutants defective in DNA replication, it is possible to obtain an excess of the early enzymes, including DNA ligase (28, 34). Using these mutants and either a modification (34) of the original purification procedure or a new four-step purification procedure (28),a homogeneous T4 DNA ligase can be obtained, as judged by polyacrylamide gel electrophoresis in the presence of sodium dodecyl sulfate. The T4 DNA ligase has been amplified in E. coli, allowing one to isolate relatively large quantities of this enzyme. Frequently, however, it is ad30. Modrich, P., Anraku, Y., and Lehman, I. R. (1973). JBC 248, 7495. 31. Gellert, M., and Bullock, M. L. (1970). PNAS 67, 1580. 32. Panasenko, S . M., Cameron, J . R., Davis, R. W., and Lehman, I. R. (1977). Science 196, 188. 33. Panasenko, S. M., Alazard, R. J., and Lehman, I. R. (1978). JBC 253, 4590. 34. Panet, A . , van de Sande, J. H., Loewen, P. C., Khorana, H. G., Raae, A. J., Lillehaug, J. R., and Kleppe, K. (1973). Biochemistry 12, 5045.

1.

DNA LIGASES

9

vantageous to purify T4 DNA polymerase, T4 polynucleotide kinase, and T4 RNA ligase as well as the DNA ligase from a single extract. In these cases all four enzymes can be purified from T4 phage-infected cells by published procedures for the polymerase (351,kinase (36) and RNA ligase (37) using the appropriate side fractions obtained from the ligase purification procedure (29). Simultaneous purification of these enzymes, using these procedures (34) or a novel procedure (38), has been described. A phage A derivative has been constructed that contains the T4 DNA ligase gene (39). The E. coli lysogen that contains this prophage can be induced to synthesize the T4 DNA ligase in amounts considerably greater than that normally obtained in cells infected with phage T4 (26,40). Three procedures have been described that make possible the rapid purification of T4 DNA ligase from the lysogen (26,40,41 ), one of which results in a ligase preparation more than 95% pure (41).

C. PHYSICAL PROPERTIES 1. Molecular Weight

The molecular weight of native E. coli DNA ligase, as estimated by equilibrium sedimentation, is 77,000‘(30). The apparent molecular weight of the denatured and reduced form of the E. coli ligase, as determined by comparison with the mobilities of proteins of known molecular weight on polyacrylamide gels, is 74,000 f 3,000 (30). Thus, the E. coli DNA ligase appears to be a monomer in solution. The s20,w of the enzyme is 3.9 S, a value that is lower than would be expected for a spherical protein, thus suggesting an asymmetric shape. The T4-induced ligase has a native molecular weight of 68,000 f 6,800, as determined by gel filtration (34). The molecular weight of the denatured and reduced form of the ligase is 63,000 f 3,200 (34). Thus, the T4 DNA ligase is also a single polypeptide. The sedimentation coefficient of the native enzyme is 3.5 S, indicating a n elongated shape. The ligase encoded by bacteriophage T7 is somewhat smaller than the E. coli and T4 ligases. Its molecular weight, calculated from the nucleotide 35. Goulian, M., Lucas, S. J., and Kornberg, A. (1968). JBC 243, 627. 36. Richardson, C. C. (1965). P N A S 54, 158. 37. Higgins, N. P., Geballe, A. P., Snopek, T. J . , Sugino, A . , and Cozzarelli, N. R. (1977). Nucleic Acids Res. 4, 3175. 38. Dolganov, G. M., Chestuktin, A. V., and Shemyakin, M. F. (1981). EJB 114, 247. 39. Wilson, G . G . , and Murray, N. E. (1979). J M B 132, 471. 40. Tait, R. C . , Rodriguez, R. L., and West, R. W., Jr. (1980). JBC 255, 813. 41. Davis, R . W., Botstein, D . , and Roth, J. R. (1980). “Advanced Bacterial Genetics,” p. 196. Cold Spring Harbor Laboratory, Cold Spring Harbor, New York.

10

MICHAEL J. ENGLER AND CHARLES C. RICHARDSON

sequence of the gene, is 41,133 (42). The native molecular weight is unknown. Mammalian cells contain at least two different DNA ligases (16). The native molecular weights for each have been calculated from sedimentation coefficients and Stokes radii. DNA ligase I has a molecular weight of approximately 200,000 and DNA ligase I1 is smaller, having a molecular weight of about 85,000 (16). The subunit structure has not been determined for either enzyme but, upon prolonged incubation of crude extracts (43) or during purification (44), an active form of DNA ligase I appears that has a molecular weight of approximately 100,000. 2. Amino Acid Composition

The amino acid composition of the E. coli DNA ligase has been deterand the amino acid sequence of the T7 DNA ligase can be mined (M), deduced from the sequence of the gene (42). The ratio of absorbance at 280 nm to that at 260 nm for the unadenylated form of the E. coli DNA ligase is only 1.42, reflecting the low tyrosine content. Isoelectric focusing of the adenylated T4 DNA ligase reveals five species (28). The major band possesses a PI of 6. The unadenylated form of the enzyme has a PI of 6.2. 111.

Catalytic Properties

A.

MECHANISM OF PHOSPHODIESTER FORMATION

All DNA ligases described thus far use the hydrolysis energy of either NAD or ATP to form a phosphodiester linkage between polynucleotide chains. Enzymes from various organisms may join polydeoxyribonucleotides or polyribonucleotides. Mechanisms by which the E. coli and T4 DNA ligases form phosphodiester bridges in duplex DNA are best understood (Fig. 1). In this overall reaction phosphodiester formation is accompanied by the stoichiometric cleavage of NAD or ATP to yield AMP and either NMN+ or PPi, respectively (Fig. 1). Synthesis of a phosphodiester link at a nick in duplex DNA is dependent on the presence of a juxtaposed 3’-hydro~ylgroup and a 5’-phosphoryl group. The reaction (Fig. 2) proceeds in three discrete steps: (i) Covalent transfer of the adenylyl group of NAD or ATP to an €-amino group of a lysine residue in the enzyme, with the elimination of NMN or PP,; (ii) 42. Dunn, J . J., and Studier, F. W. (1981). J M B . 148, 303. 43. Pedrali-Noy, G . C. F., Spadari, S . , Ciarrocchi, G . , Pedrini, A. M., and Falaschi, A. (1973). EJB 39, 343. 44. Soderhall, S., and Lindahl, T.(1975). JBC 250, 8438.

1.

11

DNA LIGASES ATP or NAD+

ATP

+

AMP

+

.

AMP

PPi

or NMN+

+ PPi 4

0 P ;

to-

FIG. 1. Phosphodiester formation at nicks (A) and at blunt ends (B).

activation of the 5’-phosphoryl terminus at the nick in DNA by transfer of the adenylyl group from the enzyme to the 5’-phosphoryl group to generate a new pyrophosphate moiety; (iii) phosphodiester formation by nucleophilic attack of the 3’-hydroxyl group at the nick on the activated 5’-phosphoryl group with the release of AMP. Each of the intermediates formed in these steps has been isolated and characterized. 1. Ligase-Adenykrte Intermediate

The E. coli ligase catalyzes an exchange reaction between NMN and NAD (45, 4 6 ) , and the T4 and mammalian ligases catalyze an analogous exchange between pyrophosphate and ATP (13,47-49). The DNA ligase of bacteriophage T7 can use either ATP or dATP as a cofactor, and catalyzes an exchange reaction between pyrophosphate and either ATP or dATP (50). The E. coli and T4 ligase-adenylate intermediates (ligase-AMP) have been identified and isolated by such techniques as gel electrophoresis, 45. Olivera, B. M., Hall, Z. W., Anraku, Y., Chien, J. R., and Lehman, I. R. (1968). C S H S Q B 33, 27. 46. Little, J. W., Zimrnerrnan, S. B . , Oshinsky, C. K . , and Gellert, M. (1967). P N A S 58, 2004. 47. Weiss, B . , and Richardson, C. C. (1967). J B C 242, 4270. 48. Weiss, B., Thompson, A., and Richardson, C . C. (1968). J B C 243, 4556. 49. Lindahl, T., and Edelrnan, G. M. (1968). PNAS 61, 680. 50. Hinkle, D. C . , and Richardson, C. C. (1975). J B C 250, 5523.

0-

HO OH

no

OH

FIG.2. Mechanism of the reactions catalyzed by E. coli and T4 phage DNA ligases.

I . DNA LIGASES

13

sedimentation, and gel filtration (30, 45-48). Using homogeneous E. coli DNA ligase it has been shown that one molecule of AMP becomes covalently linked per protein molecule (30). Interestingly, the joining activity of E. coli DNA ligase can be destroyed by limited treatment with trypsin without destroying its ability to react with NAD to form ligase-AMP (51). The ligase-AMP compounds formed are relatively stable to acid and alkali, and the fact that they are stable when precipitated with acid (45, 46, 48) provides an assay for DNA ligase, described in Section II,A,4. Studies carried out with the isolated ligase-AMP compounds clearly show that they are competent intermediates in the overall joining reaction. When the E. coli ligase-AMP is incubated with NMN, the adenylate moiety is released from the compound and appears in NAD (45,46). Incubation of the T4 ligase-AMP with PPi results in the release of the adenylate moiety and the appearance of ATP (48). Both the E. coli and T4 ligaseAMP compounds release AMP when incubated with DNA that contains nicks, and the amount of AMP released is equal to the number of nicks in the DNA (45, 46, 48). In both E. coli and T4 ligase-AMP, the AMP moiety is covalently linked through a phosphoramide bond to the €-amino group of a lysine residue of the enzyme (52).Thus, the initial step in the ligase reaction is most likely a nucleophilic attack of the €-amino group of a lysine on the adenylyl phosphorus of NAD or ATP, leading to the formation of enzyme-bound, lysine-linked AMP. 2. DNA-Adenylcrte Intermediate While the steady-state concentration of the DNA-adenylate (DNAAMP) intermediate is extremely low, it has been accumulated in sufficient amounts for analysis under restricted conditions. Small amounts of the DNA-AMP complex have been isolated after brief incubations at 0" of large amounts of theE. coli enzyme with nicked DNA substrates (53).The DNA-AMP intermediate in the T4 DNA ligase-catalyzed reaction has been isolated by incubating the purified ligase-AMP intermediate with nicked DNA for short periods at p H 5.6 and 0" (54). At this pH the final step of phosphodiester formation appears to be inhibited more than the synthesis of the adenylyl-DNA intermediate. Evidence has also been presented that indicates 5'-adenylated DNA is an intermediate of blunt-end 51. 52. 53. 54. 4523.

Panasenko, S . M . , Modrich, P., and Lehman, I. R . (1976).JBC 251, 3432. Gumport, R. I . , and Lehman, I. R . (1971). PNAS 68, 2559. Olivera, B. M., Hall, Z. W., and Lehman, I. R. (1968). PNAS 61, 237. Harvey, C . L., Gabriel, T. F., Wilt, E. M., and Richardson, C. C. (1971). JBC 246,

14

MICHAEL J. ENGLER AND CHARLES C. RICHARDSON

joining reactions (55) and of the reversal of theE. coli ligase reaction (56). In addition, 5’-adenylated DNA can be formed by E. coli ligase given a dideoxy-terminated DNA substrate (57). In the presence of the enzyme, the isolated DNA-AMP compound will form a proper phosphodiester linkage in the absence of NAD or ATP, releasing AMP (53,54). For each mole of phosphodiester formed, one mole of AMP is released (17, 54). In fact ATP and NAD are inhibitors of the joining of the! DNA-AMP intermediate by the T4 and E. coli enzymes, respectively. This may be the result of competition for the AMP site on the enzyme, which is required for binding the complex (54, 56). The AMP in the DNA-adenylate intermediate has been shown to be in pyrophosphate linkage with the DNA at the 5’-phosphoryl groups (53,54). The structure has been confirmed by studies on chemically synthesized polydeoxyribonucleotide-adenylateintermediates (53,55, 58). There is a high specificity for the adenosine moiety of the DNA-AMP for joining by DNA ligase. When GMP is substituted for AMP, no joining can be detected by T4 ligase reaction (54). When the adenylyl moiety of the intermediate is replaced by dAMP, the rate of joining is only 8% of that observed with AMP. dATP can also exchange with pyrophosphate in the presence of T4 ligase, but at only one two-hundredth the rate of ATP (54). The presence of a pyrophosphate moiety is insufficient for joining, since a synthetic poly(dT) with a triphosphate group at its 5’-terminus is inactive when incubated with E. coli ligase in the absence of NAD (58).

3 . Kinetics of the Reaction Kinetics of the ligase reaction lends additional support to the mechanism shown in Fig. 2. The overall joining reactions catalyzed by both the E. coli and T4 DNA ligase obey ping-pong kinetics, suggesting a covalent intermediate in the reaction (57, 59). The kinetics of the partial reactions catalyzed by the E. coli DNA ligase has been reviewed in detail by Lehman (17) in an earlier volume of this series. B.

REVERSAL OF

THE

LICASEREACTION

The reversal of the sequence of reactions shown in Fig. 2 has been demonstrated using the E. coli DNA ligase (56). In the reversed reaction the ligase acts as an AMP-dependent endonuclease. Incubation of 55. 56. 57. 58. 59.

Deugau, K. V., and van der Sande, J. H. (1978). Biochemistry 17, 723. Modrich, P., Lehman, I. R . , and Wang, J. C. (1972). JBC 247, 6370. Modrich, P., and Lehman, I. R. (1973). JBC 248, 7502. Hall, Z. W., and Lehman, I. R. (1969). JBC 244, 43. Raae, A . J., Kleppe, R. K., and Kleppe, K. (1975). EJB 60, 437.

1.

DNA LIGASES

15

superhelical closed circular DNA with E. coli DNA ligase and AMP results in the formation of two new circular species: Molecules with one single-strand break, and covalently closed molecules that have lost superhelical turns. Among the product molecules that contain single-strand breaks it has been possible to identify the DNA-adenylate intermediate. In addition, ligase-adenylate has been isolated from the reaction mixture, thus establishing the fact that the ligase-AMP to DNAAMP step is also reversible. The ligase-catalyzed relaxation of the supercoiled molecules is slow, but eventually proceeds to completion. This nicking-closing activity of the ligase is a result of the reverse and forward reactions occurring on the same DNA molecule, as expected. The nicking-closing activity is dependent on the presence of AMP and can relax either positive or negative supercoiled molecules.

C. FORMATION OF PHOSPHODIESTERS AT NICKS 1. DNA Substrates

DNA ligases catalyze the joining of polynucleotide strands provided they have juxtaposed 3’-hydroxyl and 5’-phosphoryl end groups aligned in a duplex structure. Examples of such sites in natural DNAs are the annealed ends of lambda DNA (9),the endogenous nicks in T5 DNA (60), the interruptions created by the action of pancreatic DNase ( f O ) , and the annealed fragments generated by the staggered cutting action of some restriction endonucleases (61). Oligonucleotides as short as six or seven in length can be joined if annealed to long complementary deoxyribonucleotides (62 ). The self-complementary polymer, poly(dA-dT), forms a looped-back structure that DNA ligase can join to yield a circular molecule (23). Among the homopolymers that have been tested, oligo(dA) base-paired to poly(dT) is joined at only a fraction of the rate that oligo(dT) is joined when it is annealed to poly(dA) (24, 63). Interruptions in DNA molecules containing 3’-phosphoryl and 5’-hydroxyl termini (13), 3’-hydroxyl and 5‘-hydroxyl termini (20), 3‘-dideoxyribonucleotidesand 5’-phosphoryl termini (57), or 3’-hydroxyl and 5’-triphosphoryl termini (58) are not substrates for the enzymes. Finally, the T4 DNA ligase is able to join polydeoxyribonucleotides with a mispaired base at the 3’-terminus 60. 61. 62. PNAS 63.

Jacquemin-Sablon, A . , and Richardson, C. C. (1970). J M B 47, 477. Mertz, J . E., and Davis, R. W. (1972). PNAS 69, 3370. Gupta, N . K., Ohtsuka, E . , Weber, H . , Chang, S. H . , and Khorana, H. G. (1968). 60, 285. Fareed, G . C . , Wilt, E. M., and Richardson, C . C. (1971). JBC 246, 925.

16

MICHAEL J. ENGLER AND CHARLES C. RICHARDSON

(64, 65). The joining of DNA fragments with fully base-paired ends (blunt-end joining) is discussed in Section II1,D. The E. coli DNA ligase has aK, for 5'-phosphoryl termini of 2.5 x M when the substrate is the alternating copolymer, poly(dA-dT) (57). Values in the range of 2.5 x 10-'M to 5.6 x 10-'M have been reported for the joining of homopolymers (57,66).T4 DNA ligase has a K, for DNA of 6 x lo-' M for either the joining of oligo(dT)lo on poly(dA) ( 5 9 ) or the joining of DNA fragments with a two base-pair overhang generated by a restriction enzyme (67). On the other hand, the K, for internal phosM (29). phomonoesters in nicked natural DNA is 1.5 x The rate of the joining reaction catalyzed by T4 DNA ligase with short oligomers is extremely sensitive to temperature (54, 62, 68). The temperature optima depend on the length of the oligomer, with optimal joining occurring at temperatures above the T, of the substrate. This result suggests the possibility that the enzyme can stabilize a transient duplex structure.

2. R N A - D N A Hybrids ntid R N A Sirbstrates Eschevichi coli DNA ligase is capable of joining the 5'-phosphoryl terminus of a DNA chain to the 3'-hydroxyl terminus of RNA (69). However, neither RNA to RNA joining nor the joining of the 5'-phosphoryl terminus of RNA to the 3'-hydroxyl terminus of DNA has been detected. In addition, ribohomopolymers cannot substitute for deoxyribohomopolymers in the joining of deoxyribohomopolymers (24 ). The T4 DNA ligase, on the other hand, is able to join DNA annealed to RNA (63, 68, 70) and, to a slight extent, even RNA annealed to its complementary RNA strand (71). When oligo(A) is complexed with poly(U), however, joining of this substrate occurs at a much lower rate. The explanation for this observation may be the same as that proposed for the decreased efficiency of the joining of oligo(dA) annealed with poly(dT) as compared with oligo(dT) poly(dA) (63). A triple-stranded structure may be formed with these homopolymer pairs (72, 73). As might be expected 64. Tsiapalis, C. M . , and Narang, S. A. (1970). EBRC 39, 631. 65. Sgararnella, V., and Khorana, H. G. (1972). J M B 72, 427. 66. Olivera, B. M . , and Lehman, I. R. (1967). PNAS 57, 1700. 67. Sugino, A . , Goodman, H. M., Heyneker, H. L., Shine, J . , Boyer, H. W., and Cozzarelli, N . R. (1977).JBC 252, 3987. 68. Harvey, C. L . , and Wright, R. (1972). Biochemistry 11, 2667. 69. Nath, K., and Hurwitz, J. (1974). JBC 249, 3680. 70. Kleppe, K., van de Sande, J . H . , and Khorana, H. G . (1970). P N A S 67, 68. 71. Sano, H . , and Feix, G. (1974). Biochemistry 13, 5110. 72. Blake, R. D., and Fresco, J. R. (1966). J M B 19, 145. 73. Cassani, G., and Bollurn, F. J. (1967). JACS 89, 4798.

1.

DNA LIGASES

17

for an enzyme capable of joining both DNA to DNA and RNA to RNA, the T4 DNA ligase can join RNA to DNA in either orientation (69, 74). Some information on the substrate specificities of DNA ligases from other sources is available. T7 DNA ligase, assayed in crude extracts, is able to join oligo(dT). poly(A) and oligo(A)* poly(dT) (63). Mammalian DNA ligase 1 is unable to join oligo(dT)*poly(rA),and mammalian DNA ligase I1 can join such a hybrid at 5- 10% of the rate of oligo(dT) .poly(dA) joining (16). In addition, the mammalian DNA ligase I catalyzes the joining of the 5' -phosphoryl termini of oligodeoxyribonucleotides to the 3'hydroxyl termini of oligoribonucleotides, but it cannot catalyze their joining if the oligomers are aligned in the opposite orientation (75). 3 . Cofactor Requirement

The E. coli enzyme is highly specific for NAD, which is cleaved in the overall reaction to yield 5'-AMP and NMN (20, 66). NADH and the thionicotinamide derivative of NAD can replace NAD but have significantly higher K, values. The reported K, for NAD ranges from M (19, 57, 66). None o f the compounds tested 3 x lo+ M to 7 x significantly inhibited the activity with NAD (20). Other known NADrequiring DNA ligases are those of B. subrilis (15) and Salmonellri typpkii~iririirm(21). The bacteriophage-induced DNA ligases and the DNA ligases of eukaryotes use ATP as a cofactor in the joining reaction (10, 13, 16). The T4 DNA ligase can use dATP at 0.5% of the rate of ATP; in fact dATP behaves as a competitive inhibitor with regard to ATP of both the PP, exchange and the overall joining reaction (29, 48, 54). The K, for ATP in the exchange reaction is 2 x 10-6M,and the K i of dATP is 1 x 10-5M. In the joining reaction the K , for ATP is 1.4 x low5M , and the K for dATP is 3.5 x M. An additional study of the joining reaction catalyzed by T4 M (59). DNA ligase reports a K, value for ATP of 1 x The T7 DNA ligase can use dATP as cofactor in the joining reaction at approximately one-third to one-half the rate of ATP (50). The K, for dATP is approximately the same as it is for ATP, 6 x M. In the exchange reaction the K , for ATP is 3 x lO-'M, while that of dATP is 10-fold higher, 4 x 10-' M. The rate of PP, exchange with saturating levels of dATP is 2 to 3 times greater than that found with ATP (M. Engler and C. Richardson, unpublished results). Mammalian DNA ligase I has an apparent K m for ATP of 2 x lO-'M to 74. Westergaard, 0..Brutlag, D., and Kornberg, A. (1973). JBC 248, 1361. 75. Bedows, E . , Wachsman, J . T., and Gumport, R. I. (1977). Biochemisnl\~16, 2231.

18

MICHAEL J. ENGLER A N D CHARLES C. RICHARDSON

1.5 x M (49, 76-78), and mammalian DNA ligase I1 has a K m for ATP to 1 x low4M (15, 79). The fact that mammalian ligases may of 4.5 x use dATP as cofactor, but with a 200-fold higher K m , could be explained by a 0.5% contamination of the dATP with ATP (49). 4. pH Optima The joining reaction of E. coli DNA ligase has an optimal pH range of 7.5-8.0 in Tris-HCI buffer, and an optimal pH of 8.0 in sodium phosphate buffer (20). The pH optimum for the NAD-NMN exchange reaction in potassium phosphate buffer is pH 6.5 (46). The exchange rate is approximately 50% lower at pH 5.6 or 7.5. In Tris-HC1 buffer at pH 8.0, the standard condition for the joining reaction, the rate of the exchange reaction is 20% of the maximal. The optimal pH range for the joining of nicks by T4 DNA ligase is 7.2 to 7.8 in Tris-HC1 buffer. At pH 6.9 and 8.0 the enzyme has 46 and 65%, respectively, of its activity at pH 7.6 (29). The pyrophosphate exchange reaction of the T4 DNA ligase, in contrast to that of E. coli DNA ligase, exhibits a pH optimum similar to that for the joining reaction (48). The T7 DNA ligase has a pH optimum extending from pH 7.2 to 7.7 in Tris-HC1 buffer; at pH 8.4 the activity is one-half maximal. The rate in potassium phosphate buffer (pH 7.5) is less than 10% of the rate in TrisHCI at optimum pH (Engler & Richardson, unpublished results). Mammalian DNA ligase I has a pH optimum between 7.4 and 8.0 in Tris-HC1 buffer. The mammalian DNA ligase I1 has a pH optimum at pH 7.8 (80). The L cell DNA ligase I shows more activity in Hepes buffer than it does with Tris-HC1 buffer, the activity being fivefold greater in reactions in the presence of 2-mercaptoethanol, twofold higher with dithiothreitol (75). 5. Requirement for Divalent Cation DNA ligases require a divalent cation for activity. Although Mg2+is commonly used, other metals can fulfill this requirement. In the case of M. the E. coli DNA ligase, the optimal Mg2+concentration is 1-3 x Slightly higher rates are observed when Mn*+is substituted at levels of 2- 10 x 10-4M(20),but MnZ+maybe inhibitory at higher concentrations (I 1). Ca2+was 60% as active as Mg2+in one study ( II ) whereas it was inactive in another, as were Co2+and Ni2+(20). Slight activity was obtained with 76. Bertazzoni, U . , Mathelet, M., and Campagnori, F. (1972). EBA 287, 404. 77. Beard, P. (1972). BBA 269, 385. 78. Young, H., Har, T. S., Morrice, L. A. F., Feldberg, R. S . , and Keir, H. M. (1973). Biorhem. SOC. Trans. 1, 520. 79. Zimmerman, S. B . , and Levin, C. J. (1975). JBC 250, 149. 80. Soderhall, S . , and Lindahl, T. (1973). BBRC 53, 910.

1.

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19

Zn" (20). The optimum concentration of Mg2+for the T4 DNA ligase is 1x M , and at its optimal concentration, Mn2+is only 25% as effective as Mg2+(29, 48). In this study the ligase substrate was nicked duplex T7 DNA. Still, it was found that, in the reactions with hybrids of ribohomopolymers and deoxyribohomopolymers, the rate of joining at the optimal Mn2+ concentration was twice that observed with Mg2+ (63). Mammalian DNA ligases can use Mn2+, but less effectively than Mg2+(80). 6 . Sulflydryl Requirement

The activity of the E. coli DNA ligase does not require the addition of a sulfhydryl reagent (If, 2 0 , 2 / ) .The bacteriophage-induced and eukaryotic DNA ligases require reducing agents such as P-mercaptoethanol or dithiothreitol(f6,48). The T4 DNA ligase is, in fact, much more effective with the latter (48). 7. Activutors ctrid Inhibitors Low concentrations of the monovalent cation, NH:, markedly stimulate the joining reaction of E. coli DNA ligase (57). Saturating concentrations of NH: increase the V,,, twentyfold. The presence of NH: has no effect on the rate of NMN exchange, indicating that the activation occurs at a step subsequent to the formation of the enzyme-adenylate intermediate. It has been proposed that NH: stimulates the rate of association and dissociation of ligase-adenylate and DNA. Other monovalent cations also stimulate the E. coli DNA ligase, and at saturating levels (they have different apparent K , values) K+, Rb+, and NH: give similar maximal velocities. Cs+ and Li+ also stimulate the reaction, but Na+ has no effect in the concentration range 0 to 10 mM. Low concentrations of NH: have no effect on the T4 DNA ligase reactions. Higher levels ( 0 . 2 M ) of Na+, K+, Cs+, and Li+ and NH: inhibit the enzyme almost completely (59). The polyamines, spermine and spermidine, also inhibit the reaction. These inhibitions could be explained by the observed increase in the apparent K , for the DNA substrate caused by these ions. The enzyme is most sensitive to spermine which inhibits joining about 90% at a concentration of 1 mM. However these inhibitions can be overcome by increasing the DNA concentrations. D. BLUNT-END JOINING A surprising result indicated that two DNA molecules with fully basepaired ends could be joined by the T4 DNA ligase (81 ). This reaction (Fig. 81. Sgaramella, V., van de Sande, J. H., and Khorana, H. G. (1970). P N A S 67. 1468.

20

MICHAEL J . ENGLER AND CHARLES C. RICHARDSON

1B)has been given the name “blunt-end joining.” A short DNA duplex that contains a protruding self-complementary sequence at one end and fully base-paired at the other forms not only the expected dimeric molecule, but also higher molecular weight products. Also, much of the 5’-32P at the base-paired ends of such substrate molecules is rendered resistant to phosphatase. The reaction is intermolecular in character, as shown by the covalent joining of two different molecules (82). Additional substrates have confirmed the joining of duplexes at fully base-paired ends (blunt-end joining). Thus the DNA of bacteriophage P22 can be joined at its base-paired ends to yield dimers and higher oligomers (83). Blunt-end ligation has been assayed by the joining of restriction fragments generated by endonucleases that give fully base-paired ends and by the formation of polymers from short, fully self-complementary oligonucleotides (55, 67, 84-87). Ligation of such substrates can be monitored by adaptation of the variety of assay techniques described in Section I1,A. The T4 DNA ligase is the only DNA ligase known that can catalyze blunt-end joining. The E. coli enzyme is totally inactive in this reaction (67, 83). Blunt-end joining proceeds less readily than the sealing of the nicks formed by the annealing of cohesive-end fragments (67, 84). In contrast to the joining of cohesive fragments, blunt-end joining is not linearly dependent on enzyme concentration, requiring large amounts of enzyme. In this regard the T4 RNA ligase (88) has been found to stimulate the rate of blunt-end joining of the T4 DNA ligase, particularly at low concentrations of DNA ligase (67). RNA ligase by itself cannot catalyze blunt-end joining and it does not enable the E. coli DNA ligase to catalyze blunt-end joining. The joining of nicks by T4 DNA ligase is only slightly stimulated by T4 RNA ligase (67). That blunt-end joining is in fact an activity of the T4 DNA ligase now seems certain since it has been shown that the T4 DNA ligase purified from a lambda lysogen that contains the T4 ligase gene, can join blunt-ended restriction fragments (39, 40). The K , for blunt ends is 50 /.LM and is unaffected by the presence of T4 82. Sgaramella, V., and Khorana, H. G. (1972). J M B 72, 493. 83. Sgaramella, V. (1972). PNAS 69, 3389. 84. Marians, K . J . , Wu, R., Stawinski, J., Hozumi, T., and Narang, S. A. (1976). Nature ( L o n d o n ) 263, 744. 85. Heynecker, H . L., Shine, J., Goodman, H. M., Boyer, H. W., Rosenberg, J . , Dickerson, R. E., Narang, S. A,, Itakura, K., Lin, s.,and Riggs, A. D. (1976). Nature (London) 263, 748. 86. Backman, K., Ptashne, M., and Gilbert, W. (1976). P N A S 73, 4174. 87. Sgaramella, V., and Ehrlich, S. D. (1978). EJB 86, 531. 88. Uhlenbeck, O., and Gumport, R. (1981). Chapter 2, this volume.

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RNA ligase (67). This compares with an apparent K , for ends of 0.6 ph4 in nick-sealing reactions (59. 67). The optimal temperature for the joining of the blunt-ends of duplex structures sixteen nucleotides or longer, including long molecules generated by restriction enzymes, is approximately 25” (82, 87). Smaller duplexes require lower temperatures consistent with their melting temperatures (55, 67). Nicked intermediates can be observed in the joining of a self-complementary octanucleotide. This result suggests that blunt-end joining occurs in two discrete steps, the joining of two molecules to form a nicked dimer, followed by sealing of the nick. Kinetic data using this same substrate indicates that the rate-determining step is a bimolecular reaction in which two duplexes are joined to form a nicked dimer (55).

IV.

Role of DNA Ligases in Vivo

The reaction catalyzed by DNA ligase makes it likely to be involved in a number of processes in DNA metabolism. Covalent joining of DNA chains is known to be required for replication, recombination, and repair. However, a more direct and definitive approach to determine the roles of DNA ligase is to examine the biochemical consequences of mutations that directly affect the enzyme. In fact, studies with mutants ofE. coli, phages T4 and T7, and yeast have shown that defects in DNA ligase lead to aberrations in each of these processes. DNA L l G A S E S A. PHAGE-INDUCED 1. Pliuge T4 D N A Ligase

The availability of a collection of conditionally lethal mutants of bacteriophage T4 (89) made possible the identification of a T4 ligase-deficient mutant (90) shortly after the discovery of the T4 DNA ligase. Gene 30 of phage T4 was shown to be the structural gene for DNA ligase by direct assay of extracts of E. coli infected with umber and temperature-sensitive mutants of T4. Although mutants defective in gene 30 were originally classified as DNA-negative (89), subsequent studies established that phage DNA synthesis begins in normal fashion after infection of E. coli 89. Epstein, R. H . , BoIle, A., Steinberg, C. M., Kellenberger, E., Boy de la Tour, E., Chevalley, R., Edgar, R. S., Susman, M . , Denhardt, G. H., and Lielausis, A. (1963). CSHSQB 28, 375. 90. Fareed, G. C., and Richardson, C. C. (1967). PNAS 98, 665.

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MICHAEL J. ENGLER AND CHARLES C. RICHARDSON

with umber mutants defective in this gene (91-97). However, the rate of DNA synthesis rapidly decreases, and synthesis stops within 10 min after infection. The mutant phages produce less than 10% of the amount of DNA produced by wild-type T4 (83, 93, 94). The inviability of T4 gene 30 mutants under nonperrnissive conditions clearly demonstrates an essential role of T4 DNA ligase. However, it has not been possible to identify directly a specific lesion in DNA metabolism that is responsible for this lethality of the gene 30 mutation. Low molecular weight DNA fragments reminiscent of Okazaki fragments, with sedimentation coefficients of 5 to 15 S in alkali, accumulate in cells infected under nonpermissive conditions with T4 phages defective in gene 30 (91, 95-101). The fragments are present in a duplex structure, and can be partially joined in vitro with DNA ligase (91, 98). Equally important, however, is the fact that in the absence of a functional ligase, parental phage DNA, and even the progeny DNA synthesized prior to inactivation of the enzyme, also accumulates fragments (91, 95, 97, 100, 101). It seems likely that the lethality of the gene 30 mutation is the result of multiple defects in DNA metabolism that arise in the absence of a functional ligase. Although the ligase mutation in phage T4 drastically affects DNA synthesis, it apparently has little effect on genetic recombination and the repair of ultraviolet damage. T4 gene 30-deficient mutants exhibit the same frequency of genetic recombination as does wild-type T4 phage (91, 95, I02, 103), although under certain conditions the T4 ligase appears to be required for the formation of covalently linked T4 recombinant molecules in v i t (104). ~ T4 gene 30 mutants show only a slight increase in sensitivity to ultraviolet light compared to wild-type T4 (105). 91. Richardson, C. C., Masamune, Y . ,Live, T. R., Jacquemin-Sablon, A . , Weiss, B . , and Fareed, G . C. (1968). C S H S Q B 33, 151. 92. Hosoda, J. (1967). BBRC 27, 294. 93. Bolle, A . , Epstein, R. H., Sdser, W., and Geiduschek, E. P.(1968). JMB 33, 339. 94. Warner, H . R., and Hobbs, M . D. (1967). Virology 33, 376. 95. Kozinski, A. W. (1968). C S H S Q B 33, 375. 96. Okazaki, R., Okazaki, T., Sakabe, K., Sugimoto, K., Kainuma, R., Sugino,A., and Iwatsuki, N . (1968). C S H S Q B 33, 129. 97. Sugimoto, K., Okazaki, T., and Okazaki, R. (1968). PNAS 60, 1356. 98. Masamune, Y . , and Richardson, C. C. (1968). PNAS 61, 1328. 99. Newman, J., and Hanawalt, P. C. (1968). J M B 39, 639. 100. Hosoda, J . , and Mathews, E. (1968). PNAS 61, 997. 101. Kozinski, A. W., and Kozinski, P. B. (1968). BBRC 33, 670. 102. Bernstein, H . (1968). C S H S Q B 33, 325. 103. Kozinski, A. W., and Kozinski, P. B. (1969). J . Virol. 3, 85. 104. Anraku, N . , and Lehman, I. R. (1969). JMB 46, 467. 105. Baldy, M. W. (1970). Virology 40, 272.

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2. Phage T7 DNA Liguse An amber mutant of phage T7 that fails to induce the T7 DNA ligase was initially obtained by mutagenesis of phage T7 using hydroxylamine (106). Mapping of the first T7 ligase mutant, as well as other point and deletion mutants, established its position to be between genes 1 and 2 on the T7 genetic map; hence the ligase gene is designated “gene 1.3” (107). T7 ligase mutants, including deletion mutants, grow normally in wildtype E. coli but fail to produce progeny phage when grown on ligasedeficient strains ofE. coli (106-108); wild-type T7 grows normally on such strains. T7 gene 1.3 mutants are no more sensitive to ultraviolet light when grown in wild-type E. coli than are wild-type T7 phage (106).No defect in T7 DNA replication is observed in wild-type E. coli infected with T7 ligase mutants. However, during T7 ligase mutant infection of ligase-deficient E . coli strains, there is a marked accumulation of small fragments of newly synthesized DNA (106).

B . Escherichiu coli DNA LIGASE Two classes of ligase mutants have been isolated: Those that produce a defective enzyme (lig), and those that produce increased amounts of a normal enzyme (lop). Both lop and lig map at 52 min on the genetic map (109, 110). The first lop mutants were isolated by selecting mutagenized E. coli cells capable of supporting the growth of T4 gene 30 mutant phages (108). The rationale behind such a screening was that increased levels of E. coli ligase would replace the requirement for the phage ligase. This screening strategy was chosen because T4lig rIt double mutants grow in wild-type E . coli, while T4lig phages do not, thus showing that the requirement for T4 ligase is not absolute (see Section C below). A representative mutant strain, lop8, did indeed produce a normal ligase, but in four to five times the amount of the parental strain. Ligase-deficient strains were then isolated as pseudorevertants of the lop8 strain (108). One of these, lop8, lig4, was found to be temperature-sensitive both in the ability to plate T4 ligase mutants and in DNA joining activity. The lig4 mutation has been separated from the overproducing mutation by P1 transduction, and ligase 106. Masamune, Y., Frenkel, G. D . , and Richardson, C. C. (1971). JBC 246, 6874. 107. Studier, F. W. (1973). J M B 79, 227.

108. Gellert, M . , and Bullock, M. L. (1970). PNAS 67, 1580. 109. Gottesman, M. M . , Hicks, M. L., and Gellert, M. (1973). J M B 77, 531. 110. Bachmann, B. J., and Low, K. B. (1980). M i c r o b i d . R e v . 44, 1 .

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MICHAEL J. ENGLER AND CHARLES C. RICHARDSON

purified from this strain is thermolabile (109, 111). An independently isolated temperature-sensitive mutant, ligts7, has also been shown to be deficient in DNA ligase (111-114). Of the mutations studied, lists7 seems to have the most severe ligase deficiency. The lig4 mutant, which has about 1% of the normal ligase joining activity at 42”, has normal growth rates under all conditions. It is UV-resistant, and is only moderately sensitive to methylmethane sulfonate, indicating that its ability to repair DNA may be less severely impaired (109). The ligts7 mutant has only 5% of wild-type ligase activity at the permissive temperature and less than 1% activity when assayed at 42”. In addition, it is more UV-sensitive than the wild type, and is quite sensitive to treatment with methylmethane sulfonate. Although both ligrs7 and lig4 show approximately 1% of wild-type DNA joining activity at the nonpermissive temperature, the ligts7 enzyme is also temperaturesensitive in regard to enzyme-adenylate formation. The lig4 enzyme, on the other hand, can be adenylated normally, even at elevated temperature. Thus, joining assays may not accurately reflect the effective relative amounts of ligase activity in these strains because they do not necessarily measure turnover of the enzyme (108, 109, 114). The ligis7 mutation is in fact conditionally lethal, but its defect is lethal only after prolonged incubation at 42” (109,111,114). Where then might the defect lie, and how does it relate to the essential function of DNA ligase in the cell? The DNA synthesis in ligase-deficient cells continues at nearly normal rates after a shift in temperature to 42”, even as cell death is occurring (111, 114). The nature of DNA synthesis, however, is abnormal. There is a marked defect in the joining of Okazaki fragments (109, 114). The strains containinglig4 are slow to join short nascent DNA, and in ligts7 strains these pieces accumulate in large quantity. Escherichia coli mutants defective in DNA ligase also show increased recombination and mutation frequencies (115, 116). In fact some E. coli ligase mutants have been isolated on the basis of the “hyper-rec” phenotype (I 15). It is clear, then, that ligase is indispensable for normal cell growth and that inviability of mutants seems to be primarily the result of an inability to seal Okazaki fragments. It would seem that E. coli normally produces 111. Gottesman, M. M., Hicks, M. L., and Gellert, M. (1973). I n “DNA Synthesis in (R. Wells and R. Inman, eds.), p. 107. University Park Press, Baltimore, Maryland. 112. Pauling, C . , and Hamm, L. (1969). PNAS 64, 1195. 113. Modrich, P., and Lehman, I. R. (1971). PNAS 68, 1002. 114. Konrad, E. B . , Modrich, P., and Lehman, I. R. (1973). J M B 77, 519. 115. Konrad, E. B. (1977). J . Bacreriol. 130, 167. 116. Morse, L . S., and Pauling, C. (1975). PNAS 72, 4645.

vitro”

25

DNA LIGASES

1.

an overabundant supply of DNA ligase and that the viability of lig4 at 42” is due to the persistence of significant DNA ligase levels.

c.

P H Y S I O L O G I C A L R E Q U I R E M E N T FOR PHAGE DNA LIGASES

BACTERIAL AND

As previously discussed, the levels of DNA ligase in E. coli cells can vary 500- to 1000-fold without a deleterious effect on cellular growth. Escherichiu coli cells remain viable with a five- to tenfold increase as well as with a 100-fold decrease. Escherichia coli ligase can at least partially substitute for phage T4 and T7 ligases under certain conditions. For example, as already discussed, T4 ligase mutants that cannot grow on wildtype E. coli can grow on the E. coli lop mutants that overproduce the host ligase. Furthermore, whereas T4 lig cannot synthesize DNA normally under nonpermissive conditions, the introduction of an rlZ mutation can phenotypically suppress the ligase mutation (117-119). The suppression is observed with either umber or missense mutations in the ligase gene; it does not occur through a mechanism that restores T4 ligase activity (63). A requirement for the T4 ligase is observed, however, if ligase-deficient strains ofE. coli are used as the host for T4lig rll double mutants (108).T4 rll grows normally in these strains. A separate line of evidence that suggests that the T4 phage ligase may be dispensable is the finding that normal DNA replication can occur in E. coli infected with gene 30 mutants if chloramphenicol is added 3 to 5 min after infection ( 9 5 ) . This result suggests that the ligase is simply acting to repair randomly introduced endonucleolytic breaks in the DNA. T7 ligase mutants are similar to T4 lig rZl double mutants in that they cannot grow on E. coli ligase-deficient mutants. It seems likely that some ligase activity is essential for both T7 and T4 rZZ growth, but that the host ligase is able to substitute for the phage-induced enzymes when the latter are not induced. Thus it appears that, despite different cofactor requirements (NADversus ATP), the E. coli ligase is able to act on the in vivo substrate of the T7 and T4 ligases when the latter enzymes are missing. The similarity between T7 ligase mutants and T4 rll- ligase double mutants became even more striking when it was found that specific mutations of T7 phenotypically suppress a mutation in the T7 ligase gene (120). Phenotypic reversal of the temperature sensitivity of E. coli ligts7 has 117. 118. 119. 120.

Berger, H . , and Kozinski, A . W. (1969). PNAS 64, 897. Ebisuzaki, K., and Campbell, L. (1969). Virology 38, 701. Karam, J. D. (1969). BBRC 37, 416. Pao, C. C., and Speyer, J. F. (1975). PNAS 72, 3642.

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MICHAEL J . ENGLER A N D CHARLES C. RICHARDSON

been obtained with bacteriophage Mu (121). Furthermore E. culi ligts7 lysogenic for Mu can support growth of T4 ligase-negative phage.

D. YEASTDNA LIGASE DNA ligase has been identified in a variety of eukaryotic cells, but ligase mutants have been described only in yeast. The conditionally lethal mutant, cdcl7-K42, of the yeast, Schizosaccharumyces pombe, has no detectable ligase activity at the nonpermissive temperature (122). At the restrictive temperature cells of this mutant enter S phase and undergo a complete round of DNA replication, but mitosis does not follow. Whereas the parental strands of the DNA of the mutant remain intact during this period, the nascent DNA is composed exclusively of short fragments. The mutant is slightly more sensitive to UV radiation than are wild-type cells. However, it is not so sensitive as other radiation-sensitive mutants. A Succharornyces cerevisiue mutant, cdc9 (123), a conditionally lethal cell cycle mutant, has no detectable ligase activity at the permissive or nonpermissive temperatures (124). At the nonpermissive temperature DNA synthesis occurs, but results in the accumulation of small fragments of DNA.The mutant is also more sensitive to UV irradiation. Holding cdc9 cells at the restrictive temperature leads to enhanced levels of recombination in the survivors (125).

V.

Research Applications

DNA ligases are essential reagents in studies on nucleic acid structure and metabolism. Their value derives from the specificity of the reaction and their ability to join polynucleotide chains covalently. In view of the wide applicability of the DNA ligases, it is not feasible to enumerate all of their past and present uses in studies on nucleic acids. However, it is possible to cite a few applications that illustrate their usefulness. The fact that ligases require a bihelical DNA structure containing a single-strand break displaying 3’-hydroxyl and 5‘-phosphoryl end groups in juxtaposition provides a sensitive and specific method for identifying such structures. For example, the conversion of hydrogen-bonded circles 121. 122. 123. 124. 125.

Ghelardini, P. Paolozzi, L . , and Leibart, J . C. (1980). Nircleic Acids Res. 8, 3157. Nasmyth, K. A. (1977). Cell 12, 1109. Culotti, J., and Hartwell, L. H. (1971). E.rp. Ccll Res. 67, 389. Johnston, L. H., and Nasmyth, K . A. (1978). Nutiire (London) 274, 891. Game, J. C., Johnston, L. H., and von Borstel, R. C. (1979). / “ A S 76, 4589.

I.

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27

of phage X DNA to covalently bonded ones using DNA ligase provided the first demonstration that the strands of these circles were continuous over their entire length except for a single phosphodiester interruption (9, 12, 13, 20, 29). Similarly, the finding that the specifically located interruptions found in the individual strands of coliphage T5 DNA are repaired by DNA ligase, demonstrated that the T5 DNA molecule contains single-strand breaks bearing 3’-hydroxyl and 5’-phorphoryl groups without any missing nucleotides (60). DNA ligase has also been used to determine the specificity of endonuclease cleavage. For example, the gene 3 endonuclease of phage T7 was shown to produce single-strand breaks in duplex DNA that could be repaired by DNA ligase (126 ), and the Eco RI restriction endonuclease was found to produce cohesive ends that could be base-paired to form a substrate for DNA ligase (61). The breakdown of ligase-AMP to yield phosphodiester and AMP occurs only in the presence of nicks in duplex DNA displaying 3’-hydroxyl and 5’-phosphoryl groups. Therefore the extent of breakdown of the intermediate is equal to the number of these groups present in a DNA preparation (48). Since ligase-AMP can be easily prepared using radioactively labeled ATP of high specific radioactivity, ligase-AMP can be used as a reagent in an assay to measure the number of nicks in a given DNA preparation. When measured using this technique T5 s t ( 0) DNA contains 3.6 singlestrand breaks per duplex molecule (60). In combination with polynucleotide kinase end-group labeling, DNA ligase can be used to identify 3’- and 5’-end groups at single-strand interruptions by nearest neighbor analysis. Studies of this type include the analysis of sites at which pancreatic DNase introduces nicks into duplex DNA (29) and the 3’ termini of the strands of A DNA (90, 127, 128). DNA ligases can be used to determine the ability of other enzymes to act at nicks and gaps in duplex DNA molecules. The ability of DNA ligase to rejoin a DNA strand provides an extremely sensitive assay to monitor exonuclease action at nicks in DNA. For example, E. coli exonuclease I11 and the exonuclease activity of T4 DNA polymerase, in contrast to phage X exonuclease, can carry out hydrolysis at nicks and thus prevent subsequent joining by DNA ligase (129). Likewise, E. cofi DNA polymerase I can destroy ligase substrates by displacement synthesis starting at nicks, whereas T4 DNA polymerase cannot (130). On occasion it is essential or desirable to eliminate nicks and gaps that have arisen during isolation of Center. M. S . , and Richardson, C. C. (1970). JBC 245, 6292. Wu, R . , and Kaiser, A. D. (1967). P N A S 57, 170. Wu, R . , and Kaiser, A. D. (1968). J M B 35, 523. Masamune, Y., Fleischman, R. A,, and Richardson, C. C . (1971). JBC 246, 2680. 130. Masamune, Y., and Richardson, C. C . (1971). JBC 246, 2692. 126. 127. 128. 129.

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MICHAEL J. ENGLER AND CHARLES C. RICHARDSON

DNA substrates. Most such interruptions can be eliminated by a combination of treatment with DNA polymerase and DNA ligase (129, 131). The DNA ligase reaction has been used to study the primary and secondary structure of DNA molecules. Polymers such as the (dA).(dT) homopolymer pair and the d(AT) copolymer exhibit conformational mobility in that hydrogen bonds between the polynucleotides are broken and shifted constantly, permitting the chains to slip with respect to each other. Information of this nature has been derived in part from their ability to serve as substrates for DNA ligase (22, 24). By measuring the joining of short oligomers by DNA ligase, it is possible to derive information concerning the parameters that effect the secondary structure of nucleic acids. For example, the temperature curves for the joining of short oligomers are characteristic of the length and base composition of the oligomer (54, 6 2 , 6 8 ) . Similarly, the ligase reaction can be used to monitor for the noncovalent formation of circular molecules via cohesive ends produced by restriction endonucleases (132). Finally, DNA ligase can be used to study parameters that effect supercoiling in circular DNA molecules (133). DNA ligase has been an indispensible reagent in the chemical synthesis of double-stranded DNAs of specific nucleotide sequence. For example, the synthesis of the structural genes for alanine and tyrosine tRNA was accomplished by covalently joining overlapping synthetic polydeoxyribonucleotide segments using DNA ligase (134, 135). An important use of DNA ligase is in the preparation of recombinant DNA molecules for use in the cloning of DNA. These techniques are reviewed in detail elsewhere (19, 136-138). These methods allow one to join DNA molecules in vitro and to introduce the resulting recombinant DNA molecules into cells where they are amplified via replication. Hydrogen-bonded recombinant DNA molecules can be generated by annealing two DNA fragments containing complementary and antiparallel single-strand extensions (cohesive ends). Such cohesive ends can be gen131. Kolodner, R., Masamune, Y., LeClerc, J. E., and Richardson, C. C. (1978). JEC 253, 566.

132. Dugaiczyk, A., Boyer, H. W., and Goodman, H. M. (1975). J M E 96, 171. 133. Wang, J. (1981). “The Enzymes,” 3rd ed. Vol. XIV, Chapter 18. 134. Khorana, H. G., Agarwal, K. L., Buchi, H . , Caruthers, M. H . , Gupta, N . K., Kelppe, K . , Kumar, A., Ohtsuka, E., RajBhandary, U . L., van de Sande, J. H., Sgaramella, V., Terao, T., Weber, H . , and Yamada, T. (1972). J M E 72, 209. 135. Brown, E. L., Belagaje, R., Ryan, M. J . , and Khorana, H. G. (1979). “Methods in Enzymology,” Vol. 68, p. 109. 136. Sinsheimer, R. L. (1977). Annrr. Rav. Eiocham. 46, 415. 137. Vosberg, H.-P. (1977). H u m a n Genet. 40, 1 . 138. Morrow, J. F. (1979). “Methods in Enzymology,” Vol. 68, p. 3.

I.

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29

erated by cleavage with type I1 restriction endonucleases or by the addition of complementary homopolymer tails to the appropriate fragments. After annealing and filling any resulting gaps using DNA polymerase, covalent joining of the fragments is accomplished with DNA ligase. Cohesive ends can also be generated by blunt-end ligation of synthetic DNA linkers, 8 to 14 base pairs in length, that contain the recognition sequence for a restriction endonuclease that produces cohesive termini. Alternatively, T4 DNA ligase can be used to catalyze blunt-end joining of two fragments, a reaction stimulated by RNA ligase. T4 DNA ligase must, of course, be used for blunt-end joining, but it is also the enzyme of choice for the joining of cohesive ends since it requires a smaller overlapping sequence than does the E. coli enzyme.

This Page Intentionally Left Blank

T4 RNA Ligase OLKE C. UHLENBECK

I. Introduction

. . .

RICHARD I. GUMPORT

. . . . . .

. , . . . . . . .

,

. AAssays . . . . . . . . . . . . . . B . ProteinIdation . . . . . . . . . . C. Physical Properties . . . . . . . . . 111. Reactions Catalyzed . . . . . . . . . . A. Intermolecular Forward Reaction . . B. Circularization Reaction . . . . . . . C . ATP-Independent Reaction . . . . . D. Reverse Reactions . . . . . . . . . E.Summary . . . . . . . . . . . . . IV. Biological Role . . . . . . . . . . . . V. Applications . . . . . . . . . . . . . A. Oligonucleotide Synthesis . . . . . . B. Nucleic Acid Modification . . . . . . 11. Purification and Properties

1.

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

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

,

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.

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.

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31 33 33 34 35 35 36 41 42 44 46 47 52 53 57

introduction

T4 RNA ligase was discovered in Jerard Hurwitz’s laboratory in 1972 during a study of T4 DNA ligase ( I ) . The enzyme was originally detected as an activity that catalyzes the circularization of homopolyribonucleotides with a 3’-terminal hydroxyl and a 5’-terminal phosphate through the formation of a 3‘ + 5’ phosphodiester bond, with hydrolysis of ATP to AMP and PP, ( 1 , 2 ) . RNA ligase activity can be found inE. coli only after 1. Leis, J., Silber, R., Malathi, V. G., and Hurwitz, J. (1972). In “Advances in the Biosciences” (G. Raspe, ed.), Vol. VIII, p. 117. Pergamon, New York. 2. Silber, R., Malathi, V. G., and Hurwitz, J. (1972). PNAS 69, 3009. 31 THE ENZYMES, VOL. XV Copyright @ 1982 by Academic Press. Inc. All rights of reproduction in any form reserved. ISBN 0-12-122715-4

32

OLKE C. UHLENBECK AND RICHARD I. GUMPORT

infection by T-even bacteriophage. No activity is observed after infection with T-odd or QP bacteriophage or after the induction of bacteriophage A (2). The enzyme was subsequently shown to carry out intermolecular reactions with either RNA ( 3 , 4 )or DNA ( 5 , 6 ) , and also to use analogues of one of its reaction intermediates as substrates (7). In addition, the same protein promotes the noncovalent attachment of the tail fibers to the base plate of bacteriophage T4 (8).Although this morphogenetic function of the protein has a physiological role, the function of the nucleic acid joining activity remains obscure. The removal of internal sequences from primary transcripts of some eukaryotic tRNAs (9) and mRNAs (10) must also involve the intermolecular joining of RNA molecules. In the only example known, the maturation enzyme that joins the two half-molecules of yeast tRNAs operates by a mechanism different from that employed by T4 RNA ligase. In contrast to the bacteriophage RNA ligase this enzyme joins RNA molecules with 3’-phosphate and 5’-hydroxyl termini (I I). Although there were early reports of an eukaryotic RNA ligase activity that used assays involving homopolyribonucleotides (1, 12), these results were probably incorrect (13). The demonstration of the “splicing” enzyme(s) will presumably require specific heteropolymeric substrates that more accurately reflect their in vivo substrates. This chapter focuses on the T4 enzyme because it is the only bacteriophage RNA ligase that has been purified to homogeneity and studied extensively. Because of the catalytic diversity of the enzyme and its potential for studies with nucleic acids, most of the literature deals with applications. However, some studies of the enzyme and its biological functions have appeared. We emphasize the reaction mechanism and biological functions of the enzyme and touch briefly on its applications in 3. Walker, G. C., Uhlenbeck, 0. C., Bedows, E., and Gumport, R. I. (1975). PNAS 72, 122. 4. Kaufmann, G., and Kallenbach, N. R. (1975). Nuture (London) 254, 452. 5. Sugino, A., Snopek, T. J., and Cozzarelli, N. R. (1977). JEC 252, 1732. 6. Moseman McCoy, M. I., and Gumport, R. I. (1980). Biochemistry 19, 635. 7. England, T. E., Gumport, R. I., and Uhlenbeck, 0. C. (1977). PNAS 74, 4839. 8 . Snopek, T. J., Wood, W. B., Conley, M. P., Chen, P., and Cozzarelli, N. R. (1977). PNAS 74, 3355. 9. Ogden, R. C., Knapp, G., Peebles, C., Kang, H. S., Beckman, J. S . , Johnson, P. F., Fuhrmann, S. A., and Abelson, J. N. (1974). In “Transfer RNA: Biological Aspects’’ (D. Soll, J. N. Abelson, and P. R. Schimmel, eds.), p. 173. Cold Spring Harbor Laboratory, Cold Spring Harbor, New York. 10. Abelson, J. (1979). Annu. R e v . Biochem. 48, 1035. 1 1 . Knapp, G . , Ogden, R. C., Peebles, C. L., and Abelson, J. (1979). Cell 18, 37. 12. Cranston, J. W., Silber, R., Malathi, V. G., and Hurwitz, J. (1974). JEC 249, 7447. 13. Bedows, E . , Wachsman, J. T., and Gumport, R. I. (1975). EBRC 67, 1100.

33

2. T4 RNA LIGASE Adenylylotion of Enzyme ATP Sile

2

p-p-p-A

u Acceptor Site

Donor Site

v

--

I

Activotion of Donor

-2

Formotion of Phosphodiester Bond p-A

,

A-p-A-p-4,

P-C-P

-

A-p-A-p-A-p-C-p I

u

FIG. 1. Three-step mechanism of the RNA ligase-catalyzed intermolecular reaction of ABand pCp to form A&p.

nucleic acid chemistry. A more detailed guide to practical applications is available ( M ) , and summaries of some of its properties have also appeared (15, 16). II.

Purification and Properties

A. ASSAYS The enzyme is usually assayed by the conversion of [5‘-32P]poly(A)into a circular form in which the label becomes insensitive to a phosphomonoesterase (1, 2). This assay can be used at all stages of enzyme purification if precautions are taken to correct for the destruction of the labeled substrate in crude extracts. [See Ref. (14) for a discussion of the problems associated with quantitating the assay.] The first step of the reaction mechanism (see Fig. 1) is the basis for an 14. Gumport, R. I., and Uhlenbeck, 0. C. (1981). In “Gene Amplification and Analysis. Analysis of Nucleic Acid Structure by Enzymatic Methods” (J. G. Chirikjian and T. S. Papas, eds.), Vol. II., p. 313. ElseviedNorth-Holland, New York. 15. Kornberg, A. (1980). “DNA Replication,” p. 261. Freeman, San Francisco, California. 16. Higgins, N. P., and Cozzarelli, N. R. (1979). “Methods in Enzymology,” Vol. 68, p. 50.

34

OLKE C. UHLENBECK AND RICHARD I. GUMPORT

ATP-[32PplPPi exchange assay ( 1 2 ) . The enzyme can also be measured by the formation of the covalent RNA ligase-AMP complex using labeled ATP (12, 17). Adenylylated enzyme can be separated from free enzyme by polyacrylamide gel electrophoresis in the presence of sodium dodecyl sulfate (18). Thus, an estimate of the proportion of enzymically active protein in a homogeneous preparation can be obtained. B. PROTEIN ISOLATION RNA ligase is synthesized in large amounts throughout the latent period of the infectious cycle (8). The preferred source of RNA ligase is E. coli infected with DNA-negative (DO) mutants of T4 that allow the synthesis of enzyme but do not lyse the cells. Bacteriophage T4 with amber mutations in genes 43 (19), 44 (20, 21), and 45 (18, 2 2 ) have all been used successfully for this purpose. A DIO mutant in either gene 44 or 45 that contains, in addition, a mutation in the regulatory gene regA has been reported to overproduce the enzyme two- to threefold with respect to the corresponding DO single mutant (18). For example, the isolation of 8.5 (18), 7.2 (19), 12 (22), and 20 (20)mg of homogeneous enzyme per 100 g of cells infected with DIO oraO -regA mutants has been reported. Recoveries of activity ranged from 13 to 22% in these purifications. Specific activity measurements suggest that as much as 1% of the protein in sonicated and centrifuged extracts is RNA ligase (19). The major problem in purifying RNA ligase is to remove the trace contaminants of nucleases that remain after apparent homogeneity is attained. Various affinity resins are useful for this purpose (19, 22-24). A typical purification of enzyme that is greater than 90% homogeneous as judged by SDS-gel electrophoresis is shown in Table I. The enzyme is free of RNase activities but would require a second Affi-Gel Blue chromatography step for use in DNA joining reactions (19). 17. Vasilenko, S . K., Veniyarninova, A. G . , Yamkovoy, V. I . , and Maiyorov, V. I. (1979). Bioorg. Khim. 5 , 621.

18. Higgins, N . P., Geballe, A. P., Snopek, T. J . , Sugino, A., and Cozzarelli, N . R. (1977). Nucleic Acids R e s . 4, 3175. 19. Moseman McCoy, M. I., Lubben, T. H., and Gumport, R. I. (1979). BBA 562, 149. 20. Last, J. A., and Anderson, W. F. (1976). ABB 174, 167. 21. Snopek, T. J., Sugino, A . , Agarwal, K . , and Cozzarelli, N. R. (1976). BBRC 68, 417. 22. Gurnport, R. I., Manthey, A. E., Baez, J. A . , Moseman McCoy, M. I., and Hinton, D. M . (1981). PRC-FRG Joint Symp. Nitcleic Acids Proteins, Shcit?g/icii, p. 237. 23. Sugiura, M . , Suzuki, M., Ohtsuka, E., Nishikawa, S . , Uemura, H . , and Ikehara, M. (1979). FEBS L e f t . 97, 73. 24. Hu, M . , Wang, A., Hua, H . , Chen, Y., and Xue, C. (1980). Sci. R e p . Beijing Unit,. 4, in press.

35

2. T4 RNA LIGASE TABLE I PURIFICATION OF RNA LIGASE" Total activity Fraction

(u x

Crude extract Streptomycin supernatant DEAE-cellulose Atfi-Gel Blue Matrex Gel Red A Hydroxylapatite

10-3)

185

163 74.6 60.8

41.0 34.1

Yield

(ro) 100 88

40 33 22 18

Total protein (mi4 6970 3 140 511 70 40 17.5

a Adapted from Ref. ( 2 2 ) . E . c d i BB (146 g) infected with T4 was the source of the material.

UUI

Specific activity (U/mg) 26 52

147 863 1035

1945 E10x3 (gene 45-)

C. PHYSICAL PROPERTIES Relatively little information is available on the physical properties of the protein. Electrophoresis on polyacrylamide gels in the presence of SDS gave apparent molecular weighfs of 41,000 (21), 43,000 ( 1 9 ) , and 45,000 (20) for the enzyme. When examined under nondenaturing conditions, apparent molecular weights of 47,000 by gel filtration and 48,200 by sedimentation equilibrium ultracentrifugation were observed, suggesting that the enzyme exists as a monomer in solution up to concentrations of 0.5 mglml (20). An isoelectric point of 6.1 has been reported for the enzyme (17). The ultraviolet spectrum of the free enzyme shows a A,, of 279 nm with an Az80/Azso of 1.98. The absorbance of a solution of 1 mg/ml of RNA ligase is 1.3 at 280 nm (14).

Ill.

Reactions Catalyzed

The overall reaction of RNA ligase is the formation of a 3' + 5 ' phosphodiester bond between nucleotide residues of oligomers that bear 3'-hydroxyl and 5'-phosphate groups. This reaction is coupled to the pyrophosphorylytic cleavage of ATP. Two covalent complexes have been isolated from reaction mixtures. In one AMP is attached to the enzyme by a phosphoamide bond, and in the other AMP is attached to the 5 ' phosphate of one of the reacting oligomers by an anhydride linkage. The demonstration that these isolated complexes are competent to react and are, therefore, likely to be intermediates, along with other evidence that will be cited below, has led to the formulation of a three-step mechanism

36

OLKE C. UHLENBECK AND RICHARD I. GUMPORT

for the reaction (Fig. 1). Variations in substrates or products, substitutions of analogues of one of the covalent intermediates, or partial reactions lead to an apparent variety of reaction types catalyzed by the enzyme. We discuss these reactions in separate sections but they can all be understood in terms of the central three-step mechanism. A. INTERMOLECULAR FORWARD REACTION Figure 1 illustrates the mechanism of the intermolecular forward reaction. The 3’-hydroxyLterminated oligoribonucleotide As, called the acceptor, is joined to the 5’-phosphorylated donor nucleotide pCp. ATP is hydrolyzed to AMP and PP, in the course of the reaction. Studies on the substrate specificity of the enzyme led to the conclusion that there are at least three nucleotide residue binding sites on the enzyme (Fig. 1). The first binds ATP, the second binds the acceptor that contains the reactive 3’-hydroxyl, and the third binds the donor that bears the 5’-phosphate to be joined to the acceptor. 1. Enzyme-AMP Formation

The initial event in the mechanism is the formation of the adenylylated enzyme from ATP with the release of PPi. This reaction can occur in the absence of acceptor and donor and the adenylylated enzyme can be discharged by PPi to reform ATP (12). The adenylylated enzyme has been isolated by gel filtration, polyacrylamide gel electrophoresis, and velocity sedimentation (12). It can be separated from free enzyme on the analytical level by polyacrylamide gel electrophoresis in the presence of SDS (18), and preparatively by dye-affinity chromatography on a Matrex Gel Red A column (22). One mole of AMP residue is bound per mole of enzyme (12, 18). Studies of the hydrolysis of a nucleotide peptide isolated from protease digests of the adenylylated enzyme suggest that the AMP residue is linked via a phosphoamide bond. The isolated material is resistant to hydrolysis by snake venom phosphodiesterase (25), a property of model compounds when the nucleotide is linked to the peptide by a P-N bond but not by a P-0 bond ( 2 6 , 2 7 ) .In addition, the isolated nucleotide-peptide is cleaved by acidic hydroxylamine, which is a treatment diagnostic for P-N link25. Juodka, B. A . , Markuckas, A. Y., Snechkute, M. A. Zilinskiene, V. J . , and Drigin, Y. F. (1980). Bioorg. Khirn.. 6, 1733. 26. Rothberg, P. G . , Harris, T. J . , Nomoto, A., and Wimmer, E. (1978). PNAS 75,4868. 27. Juodka, B . , Kirveliene, V., and Liorancaite, L. (1979). J . Carb. Nucleosides N N cleotides 6 , 333.

2. T4 RNA LIGASE

37

ages (28). Studies with the specific lysine-directed reagent 2,4pentanedione suggest that the AMP residue is linked to the €-amino group of a lysine residue of the enzyme (25). The catalysis of an ATP-PP, exchange reaction is consistent with the formation of the adenylylated enzyme. The apparent K,,,for ATP in the reaction is 12 pM. Of the common nucleoside triphosphates, only dATP substitutes for ATP, suggesting that the ATP site is highly specific in its recognition of the adenosine group. The specific activity of the exchange reaction is 50-fold greater than that of the sealing reaction (12). No conclusions can be drawn from these data concerning the kinetic competence of the adenylylated enzyme in the overall mechanism, however, because the turnover number for the sealing reaction has not been determined. The inability to saturate the enzyme with oligonucleotide substrates has made thorough steady state kinetic studies unattractive. No ATP-AMP exchange occurs with the enzyme alone (12) but the catalysis of such an exchange, when an appropriate oligonucleotide and PP, are present, can be inferred from the AMP-dependent reverse reactions (Section 111,D). 2. Adenylylated Donor Formcrtion In the second step of the mechanism the adenylyl group is transferred from the enzyme to the 5’-phosphate of the donor. The formation of a 5‘ + 5‘ anhydride linkage activates the 5‘-phosphate of the donor for subsequent reaction (Fig. 1). Significant amounts of adenylylated donor can accumulate in some RNA ligase reactions. Its structure was proved by chemical and enzymatic hydrolyses of material isolated from reaction mixtures containing labeled ATP and/or 5’-32P-labeleddonor (29-31). In addition, its structure was confirmed by chemical synthesis (30). Oligoand polynucleotides with 5’-phosphate, but not 5’-hydroxyl, termini discharge isolated adenylylated-RNA ligase (12). This finding suggests that the enzyme-AMP is a direct precursor of the adenylylated donor, however the formation of activated donor from isolated RNA ligase-AMP has not been demonstrated directly. The formation of the activated donor is stimulated by the presence of acceptor ( 5 , 32) although, with high concentrations of enzyme, the reaction to form adenylylated donor can proceed to high yield in its absence 28. Shabarova, Z. A. (1970). P r o g r . Niicleic Acid Res. M a / . Biol. 10, 145. 29. Kaufrnann, G., and Littauer, U . Z. (1974). PNAS 71, 3741. 30. Sninsky, J. J . , Last, J. A . , and Gilharn, P. T. (1976). Nucleic Acids Res. 3, 3157. 31. Ohtsuka, E., Nishikawa, S . , Sugiura, M . , and Ikehara, M. (1976). Niicleic Acids Res. 3, 1613. 32. Uhlenbeck, 0. C . , and Cameron, V. (1977). Nircleic Acids Res. 4, 85.

38

OLKE C. UHLENBECK AND RICHARD I. GUMPORT

(33).The stimulation of adenylyl group transfer from enzyme to donor by the acceptor may serve the purpose of facilitating a concerted reaction by ensuring that an acceptor is on the enzyme surface when the donor is activated (16). However, during reactions with poor acceptors the activated donor can dissociate from the enzyme and accumulate (30-32). In addition, the acceptor that stimulates the adenylylation of the donor is not necessarily the same one to which it is ultimately joined. This latter phenomenon was demonstrated by reactions in which the circularization of an oligodeoxythymidylate, a DNA to DNA joining event, was accelerated by the addition of the oligoribonucleotide acceptor As ( 5 ) . The substrate specificity for the donor in the adenylylation reaction has been indirectly determined by studying the overall joining reaction. A ribonucleoside 3’,5’-bisphosphate will serve as a substrate, whereas a 5’-nucleoside monophosphate will not, even though the latter contains the requisite 5’-phosphate. Thus, at least one nucleotide residue and a 3‘phosphate are required by the enzyme for reactivity as a donor. The second phosphate must be on the 3’-hydroxyl group because ribonucleoside 2’,5’-bisphosphates are unreactive (34) and, in addition, only poorly inhibit the reaction with the 3’,5’-bisphosphates (23). Since longer donors also function efficiently the 3’-phosphate needs only one charge. The donor binding site probably does not extend beyond the initial 5’-pNp region of the donor because the rates of reaction of a homologous series of pA,p donors with a given acceptor are very similar ( 3 4 ) .Donors as long as dX174 DNA serve as substrates (35). The effect of the nucleotide composition of donor on the reaction rate is not excessive, with pyrimidine pNps being two- to tenfold better donors than their purine counterparts. Some modifications of the sugar, the base, and the 5’-phosphate are tolerated; e.g., 2’-O-methyl (36), /inbenzoadenosine ( 3 6 ) ,and 5’-thiophosphoryl(37) pNp derivatives are substrates (see Section V, and Table 111). As a generality, it appears that the larger the base portion of the pNp the less reactive it is as a substrate; i.e., the donor efficiency is pyrimidine > purine > modified purine. The structural requirements for donors previously discussed apply only to the formation of activated donor by the adenylylated enzyme and not to the 33. Hinton, D. M., Baez, J. A., and Gumport, R. I. (1978). Bioc/iemistry 17, 5091. 34. England, T. E . , and Uhlenbeck, 0. C . (1978). Biochemistry 17, 2069. 35. Higgins, N. P., Geballe, A . P., and Cozzarelli, N. R. (1979). Niccleic Acids Res. 6 , 1013. 36. Barrio, J . R . , Barrio, M. G., Leonard, N. I., England, T. E . , and Uhlenbeck, 0. C. (1978). Biochemistry 17, 2077. 37. Bryant, F. R., and Benkovic, S. J. (1981).JACS 103, 696.

2. T4 RNA LIGASE

39

subsequent formation of the phosphodiester bond, which is discussed in the next section.

3 . Phosphodiester Bond Formation In the third step of the mechanism, the 3‘-hydroxyl of the acceptor displaces AMP from the activated donor and forms the phosphodiester bond. The activated donor, isolated either from RNA ligase reaction mixtures or chemically synthesized, reacts with an appropriate acceptor in the absence of ATP to form a phosphodiester bond and release AMP (5, 30). The reaction releases AMP coordinately with the formation of the oligonucleotide product, and one AMP is produced per phosphodiester bond formed (5). In addition, the chemically synthesized activated donor forms product in the same yield as obtained when acceptor and unactivated donor react in the presence of ATP (30). When the donor is dTlo the isolated adenylylated derivative formed by RNA ligase and ATP serves as a substrate for T4 DNA ligase in the absence of ATP, if it is aligned on a poly(dA) template. This finding indicates that the adenylylated donor is very likely an intermediate in the RNA ligase reaction and, in addition, demonstrates the identity of the intermediates for the two enzymes ( 5 ) . The formation of the phosphodiester bond is a direct nucleophilic displacement on the 5’-phosphorus of the adenylylated donor with an inversion of the stereochemical configuration about the phosphorus. This conclusion was derived from experiments in which it was found that only one of the two possible stereoisomers of a chemically adenylylated donor containing a 5’-thiophosphoanhydride group will react with an acceptor to form product. Analysis of the product thiophosphodiester bond showed its phosphorus to have opposite chirality with respect to that of the 5 ‘ thiophosphoanhydride bond of the activated donor. Thus, no covalent intermediate is likely to be involved in the third step of the mechanism (37). In addition, when a donor that contains a 5’-thiophosphate group reacts with RNA ligase and ATP, the stereoisomer of the adenylylated donor made by the enzyme is of the same chirality as the active isomer made chemically. These experiments demonstrate that the enzyme maintains a preferred chirality at the relevant phosphorus through the activation and displacement steps of the reaction. A stereochemical study of enzyme-AMP formation has not been reported but should be possible with a-thiophosphate analogues of ATP (38). The formation of the phosphodiester bond of the product requires free enzyme. Addition of ATP to reactions of isolated adenylylated donor and 38. Eckstein, F. (1979). Acwirnrs Clrcrn. Res. 12, 204.

40

OLKE C. UHLENBECK AND RICHARD I . GUMPORT

acceptor inhibits product formation, suggesting that enzyme-AMP is inactive in the third step (5). The minimal size of an oligonucleotide that will always serve as an acceptor is a trinucleoside diphosphate (4. 33, 34). Several dinucleoside monophosphates including GpG and Up1 (39) and pA2 (4) can also be joined to certain donors. Acceptors containing more than three residues do not show a dramatic increase in rates or yields, suggesting that the acceptor site probably recognizes three nucleosides and two phosphates. Acceptors as long as ribosomal and viral RNAs will react (40). There are striking compositional effects upon acceptor efficiency. With homopolymers, oligomers that contain A residues are the best acceptors; those with C or I intermediate, and with U are the worst (34). Because the trimers UAG and AUG are both poorer acceptors than AAG, the enzyme appears to discriminate against uridine residues in any of the three nucleoside acceptor subsites. The magnitude of the inactivating effect of uridine residues is illustrated by the fact that it takes 30 times more enzyme to add a donor to Usthan to A3 (34). An even larger effect is seen when DNA is tested as an acceptor. With oligonucleotides, dA, reacts over 200 times slower than rA, with a given donor ( 6 ) . Under conditions optimized for DNA joining it has not been possible to attain turnover numbers greater than two per hour (41). The reason for this low reactivity of DNA is not clear. The 2‘-hydroxyl is probably not directly involved in the mechanism since on oligonucleotide acceptor containing a terminal 2’-0-methyl group will react (36).Addition of a single 3‘-terminal ribonucleotide to an oligodeoxyribonucleotide increases its reactivity and, conversely, the addition of a deoxynucleotide to the 3‘ end of an oligoribonucleotide acceptor renders it less reactive (22). In either case, the mixed composition molecules do not react like their pure DNA or RNA analogs, suggesting that the enzyme prefers ribonucleotides in at least two of the three subsites of the acceptor site. Whether these effects are due to direct recognition of the sugar or to secondary conformational effects is unknown. Duplex structures often inhibit the RNA ligase reaction. In studies of the circularization reaction with homopolymers the addition of the complementary strand either had little effect or inhibited the reaction (12). Intermolecular reactions involving tRNA as a donor have indicated that if the 5’-terminal nucleotide is in a duplex structure the reaction will not 39. Collaboration Group of Nucleic Acid Synthesis. (1980). PRC-FRG Joint Svtnp. Nrdric. Acids Proteins, Shnnpfitii, p. 254.

40. England, T. E., and Uhlenbeck, 0. C. (1978). Nature (London) 275, 560. 41. Hinton, D. M., and Gurnport, R . I. (1979). Nucleic Acids Res. 7, 453.

2. T4 RNA LIGASE

41

occur (42). For instance, yeast tRNAPhehas the 5’-terminal pG residue in a base-paired structure in the duplex acceptor stem of the molecule and is inactive as a donor. On the other hand, in E. coli, tRNAfMet the 5’-terminal pG residue is opposite a partner with which it cannot base-pair and, as a result tRNAfMetacts as a donor. Duplex structure between an oligodeoxyribonucleotide acceptor and donor inhibits their joining. This inhibition can be overcome by the single-strand DNA binding protein RNase A (6). In contrast, however, blunt-end DNA fragments produced by Hoe111 restriction endonuclease are active donors (35). In addition, duplex structures at the 3’-hydroxyl end of the double-strand RNA of reovirus are active as acceptors (40). Thus, it appears that in some cases structured substrates cannot bind to the enzyme donor or acceptor sites, while in other cases the structures can be easily disrupted, or can be accommodated in the binding sites. However, no clear picture of these phenomena has emerged.

B. CIRCULARIZATION REACTION The circularization reaction of oligonucleotides is a special case of the intermolecular reaction in which the donor and acceptor are parts of the same molecule. When oligoribonucleotides are circularized no intermediates are observed, but if their 3’-terminal ribose structure is destroyed by periodate oxidation, or if the poor acceptor oligodeoxythymidylic acid is used, the anticipated adenylylated 5’-phosphate is observed (5). This finding along with the stoichiometry of one molecule of ATP being cleaved to AMP and PP, per molecule circularized ( I ) , and the ability of 5‘phosphate-terminated oligoadenylate to discharge isolated adenylylated enzyme ( I 2 ) ,support the identity of mechanisms in the intra- and intermolecular reactions. Because the juxtaposition of the ends of the molecule is facilitated in the intramolecular reaction, phosphodiester bond formation occurs at higher rates with less enzyme than during the intermolecular reaction. In a reaction mixture where both intra- or intermolecular reactions can occur, the circularization reactions predominate ( 4 , 42). Both compositional and length effects are seen in the circularization reaction. The composition effects (12) mirror those in the intermolecular reaction. The length effects are seen in the limits where the oligoribonucleotide is too short to circularize on the surface of the enzyme, or so long that the ends have difficulty coming into contact with one another. A study of the circularization of pA,s where n = 6- 100, indicated that As was the 42, Bruce, A. G., and Uhlenbeck, 0. C. (1978). Nucleic Acids Res. 5, 3665.

42

OLKE C. UHLENBECK AND RICHARD I. GUMPORT

shortest oligomer to react, and that the initial velocity of the reaction increased to a maximum at n = 10-16 and fell as the chain length increased further (43). Other oligomers probably behave similarly, e.g., pdTs is the shortest pdT, to circularize, and the rate increases as n increases from 8 to 20 and is decreased at n = 30 ( 5 ) . The circularization reaction requires ATP with an apparent K m of 0.2 p M . As observed with the exchange reaction, only dATP substitutes for ATP ( 2 ) . There is no evidence of saturation of the enzyme with pAzoat concentrations as high as 10 pM (12). In another study, however, apparent K m values for oligo(A) substrates have been reported at 10 pM for pAe and 1 pM for pA10,1580.0r (43). These results indicate that the kinetics with respect to the oligonucleotide substrates of RNA ligase reactions are complex. Velocity measurements in both the circularization (20) and intermolecular (3, 32) reactions are sometimes not proportional to the amount of enzyme. In addition, in some intermolecular reactions both the initial velocities and final yields depend upon the enzyme concentration (6, 32, 3 4 ) . Attempts to overcome the effect on yields by adding protein stabilizing components to the reaction mixture have failed. The reason that yields are proportional to enzyme concentrations is unknown.

C. ATP-INDEPENDENT REACTION The third step of the reaction mechanism provides an explanation for an ATP-independent reaction of RNA ligase (Fig. 1). As indicated in Section III,A,3, isolated adenylylated donors can react with acceptors in the presence of free enzyme to form product and release AMP. A large number of P-substituted ADP derivatives (Ado-5’PP-X) can serve as analogues of the adenylylated donor (Ado-5’PPS’-donor) when reacted with acceptors and enzyme. AMP is eliminated in equimolar amounts to the phosphodiester bond formed between the P-X group and the acceptor (7). In the usual ATP-dependent reaction, the 5’-terminal nucleotide of the donor must have a 3’-phosphate to accept the adenylyl group from the enzyme (Section III,A,2). In contrast, after the adenylyl group is attached to the 5’phosphate of the donor the 3’-phosphate is no longer needed for reaction with the acceptor. This is demonstrated by the high reactivity of A-5’ pp5’-N, where N is any of the four common ribonucleosides (7, 44). The relaxation of specificity for the group transferred to the acceptor is further illustrated in Table 11. Nucleotide analogs altered in both base and sugar portions, nucleotides with a-N -glycosidic linkage, and phosphate 43. Kaufmann, G., Klein, T., and Littauer, U. Z. (1974). FEBS Leu. 46, 271. 44. Ohtsuka, E . , Miyake, T., Nagao, K., Uemura, H., Nishikawa, S., Sugiura, M., and Ikehara, M. (1980). Nucleic Acids Res. 8, 601.

43

2. T4 RNA LIGASE TABLE I1 @-SUBSTITUTED ADP-DERIVATIVES As SUBSTRATES OF RNA LIGASE" Substrate

Reaction

Ado-S'PPS'-NU'' Ado-S'PPS-(l-BrAdo) Ado-5' PP5'-(2'-FI Ado) A~O-~'PPS'-C~NU Ado-5'PP5'-Rib Ado-5' PP5'-riboflavin Ado-5' PP4-pantetheine Ado-S'PP6-cyanoethanol P-2'-Ado-S'PPS'-Nir P-3'-Ado-5'PP4-pantetheine 1110-5'PP5'-Nir rAdo-S'PPS'-Nir cCyd-S'PPS'-Nir 8-BrAdo-5'PP5'-(8-BrAdo) FI-2'Ado-5' PP5'4 2 ' - FI Ado) Data taken from Refs. (7) and (44). Plus (+) indicates reaction with acceptor to form product . ' Nir represents nicotinamide riboside. a

esters of ribose, riboflavin, pantetheine, and cyanoethanol can be added to an acceptor (7). The common structural feature of successfully reacting ADP derivatives is an ester linkage between the P-phosphate of the nucleotide and a primary hydroxyl of the X group. The structural variability in the groups that can be transferred to ribonucleotide acceptors is also illustrated in Fig. 4. The nonreactivity of NADP', coenzyme A, deaminoNAD+, ENAD+, ENCD+,and the symmetrically disubstituted pyrophosphates of 8-bromoadenosine and 2'-fluoroadenosine indicate that the enzyme shows great specificity for the adenosine moiety at several sites in both its base and sugar groups. This stringency of recognition in the group eliminated is presumably a reflection of the high specificity of the ATP site of the enzyme. These results suggest that the adenylyl group of @substituted ADP derivatives is sufficient to bind them to the enzyme for subsequent reaction with the acceptor. This is consistent with the apparent K , values in the *A4 range for ATP in the circularization reaction, and for AMP in the reverse transfer reaction (see Section 111,D). When the P-substituent of the ADP derivative is a good leaving group, the products of the reaction of A-Spp-X and the acceptor are the acceptor

44

OLKE C. UHLENBECK A N D RICHARD I. GUMPORT

with a 2’,3’-cyclic phosphate at its 3‘ terminus, the X group, and AMP. This product is formed with the @substituted ADP derivatives of p-nitrophenol, 4-methylumbelliferone, fluorine, and glucose (45). The reaction probably proceeds with the P-X group being initially transferred to the acceptor in a phosphodiester linkage. However, because X is a good leaving group, and because the 2’-hydroxyl of the terminal nucleotide of the acceptor is ideally situated to attack the phosphorous in the diester linkage, the reaction proceeds to eliminate the X group and to form the 2’,3’-cyclic phosphate-terminated product (45). This conclusion is based on the finding that when the relatively poor leaving group p- methoxyphenol is the P-substituent, both the 2’ ,3‘-cyclic phosphateterminated acceptor and acceptor terminated with p-methoxylphenyl phosphate are found. The enzyme is not involved in the second part of the reaction since isolated A3Cp-p-methoxyphenol converts to A3C > p at the same rate with or without enzyme (46). Becausep-nitrophenyl ADP forms the 2’,3‘-cyclic phosphate product (45) and o-nitrobenzyl ADP transfers o- nitrobenzylphosphate to the acceptor to form a phosphodiester linkage (47), it can be concluded that a methylene group can shield the phosphorus of the X group from nucleophilic attack of the 2‘-hydroxyl of the extended acceptor and thereby prevent the nonenzymatic reaction.

D. REVERSEREACTIONS The reversal of the third step of the RNA ligase reaction was first noted in oligonucleotide synthesis reactions because of the appearance of unexpected products (48). A simplified reaction used to detect reversal of the third step involves the incubation of 3H-labeled ACG and A3Cp with RNA ligase, and measuring the formation of [3H]ACGCp. Since this reaction involves the transfer of the 3’-terminal pCp residue from one oligomer to another, it is termed a reverse rransfer reaction. The formation of ACGCp continues until an equilibrium is reached, which is dictated by the molar ratio of ACG and A3Cp present at the start of the reaction. The rate of the reverse transfer reaction is greatly stimulated by AMP but not by any of the other 5’-nucleoside monophosphates. These facts suggest that the reverse transfer reaction occurs by the formation of the adenylylated intermediate A-S’ppS’-Cpfrom AMP and A3Cp in a reversal of the third 45. Gumport, R. I., Hinton, D. M., Pyle, V. S. , and Richardson, R. W. (1980). Nucleic Acids Res., S y m p . Ser. N o . 7 , 167. 46. Pyle, V. S., and Gumport, R. I . (1981). Unpublished observations. 47. Ohtsuka, E., Uemura, H., Doi, T., Miyake, T,,Nishikawa, S . , and Ikehara, M. (1979). Nucleic Acids Res. 6 , 443. 48. Krug, M . , and Uhlenbeck, 0. C. (1982). In preparation.

45

2. T4 RNA LIGASE

step of the reaction mechanism and the subsequent transfer of the pCp residue to ACG in the ATP-independent forward reaction AsCp + PA A-S’ppS‘-Cp + ACG AaCp

+ ACG

-,A3 + A-S‘ppS‘-Cp + ACGCp +

+ pA

A3 + ACGCp

This explanation is strongly supported by the detection of 32P-labeled A-S’pp5‘-Cp in reaction mixtures containing either [5’-32P]AMP or A3[3’+ 5’32P]pCp. Presumably A3Cp can bind to both the donor and acceptor sites of the enzyme such that the last internucleotide linkage is attacked by AMP binding in the ATP site (Fig. 2). This view is supported by the fact that ATP inhibits the reverse transfer reaction by forming adenylylated enzyme. The apparent K m for AMP in the reaction is about 5 p M , which is similar to the K m of ATP in the forward reaction (2). Presumably the slow rate of reverse transfer reaction observed in the absence of added AMP is due to residual AMP present in the enzyme. A comparison of several oligonucleotide substrates in the reverse transfer reaction suggests that although reversal can occur at any internucleotide bond, there is a strong preference for 3’-phosphorylated termini. Thus, if a long oligonucleotide with a 3‘-phosphate is incubated with RNA ligase and AMP, the 3’-terminal internucleotide bond is the preferred site of cleavage. However, cleavage at other internal phosphodiester bonds can be observed to a much lesser extent. The reverse transfer reaction can lead to difficulties in the use of RNA ligase in oligonucleotide synthesis (Section V). Reverse Reoction

Adenylylation of Donor 3’-Phosphaie

dT-p-dT-p-dC-p U U

dT-p-dT-p-dC-b U

U

FIG.2. Other reactions of RNA ligase. The reverse reaction to generate an oligonucleotide with a 3‘-OHis shown at the top. The modification of the 3’-P of an oligodeoxyribonucleotideis shown at the bottom. The brackets indicate the substrate binding sites illustrated in Fig. 1 .

46

OLKE C. UHLENBECK A N D RICHARD I. GUMPORT

Reversal of both the second and third steps in the RNA ligase reaction mechanism can be detected by incubation of AMP, A,Cp, and [5’-32PlpCp with RNA ligase and measuring the formation of A3[3’ -+ 5’32P]Cp.Since the reaction involves the exchange of the 3’-terminal pCp of A3Cp with free labeled pCp, it is called the reverse exchange reaction. Again, the extent of this reaction is determined by the molar ratio of pCp to A&p, and the reaction rate is greatly stimulated by AMP and not by other 5’-ribonucleoside monophosphates. The reverse exchange reaction presumably proceeds by reversal of the last two steps of the RNA ligase reaction, and subsequent forward reaction as follows:

where the (*) represents radioactive phosphorus. Evidence for this pathway is provided by the detection of 32P-labeledadenylylated enzyme on SDS gels when [32P]AMPand A3Cp are incubated with the enzyme. Since RNA ligase catalyzes an ATP-pyrophosphate exchange reaction, all. three steps of the enzyme mechanism are reversible. In addition, the detection of the same covalent intermediates in the reverse and forward reactions strengthens the view that the postulated mechanism is correct.

E. SUMMARY The three substrate binding sites of the enzyme have different specificities (Fig. 1). The donor site binds one nucleoside residue and its two associated phosphate groups with little specificity for the nucleoside. The acceptor site probably binds three nucleosides and two phosphates and strongly prefers ribonucleosides to deoxyribonucleosides. Finally, the ATP site is highly specific for the adenosine portion of the molecule. During attempts to join oligodeoxyribonucleotides at high enzyme and ATP concentrations, an unanticipated product was found (6). The 3’phosphate of both donor and product were found to be adenylylated. To illustrate this reaction, the incubation of dT,dCp with ATP and RNA ligase results in the formation of dT4dC-3’pp5’-Awith the adenylyl group bound to the 3’-phosphate by an anhydride linkage (49). This reaction probably occurs by the 3‘-phosphate-terminated oligodeoxyribonucleo49. Hinton, D. M., Brennan, C. A., and Gumport, R . I. (1982). In preparation.

2. T4 RNA LIGASE

47

tide binding to the acceptor site and receiving an adenylyl group from the activated enzyme (Fig. 2). This conclusion is strengthened by the finding that the addition of a 3’-hydroxyLterminated acceptor to the reaction mixture decreased the formation of the 3’-modified product, presumably by competing for the same site (6). If this interpretation is correct, it corroborates the expectation that the 3’ end of the acceptor site is very near to the 5’ end of the donor site. The mechanism of the RNA ligase reaction is analogous to that of T4 DNA ligase (50). In both cases an enzyme-AMP complex is formed with the nucleotide linked by a phosphoamide bond. Both enzymes activate the 5‘-phosphate of the donor by transferring the AMP group to it to form an anhydride bond. With both enzymes the third step involves the displacement of AMP and the formation of the phosphodiester bond in the product. The major contrasting characteristics of the mechanisms are that RNA ligase aligns the ends of the reacting polynucleotides on its surface, whereas DNA ligase requires a base-paired template strand to accomplish this end. In addition, RNA ligase prefers RNA substrates, whereas DNA ligase preferentially joins DNA molecules. Although both enzymes can react with heterologous substrates, the rates are slow with respect to those with homologous substrates. The observation that RNA ligase stimulates the blunt-end joining reaction of DNA ligase by increasing the V,,, of the reaction suggests that these two enzymes might interact in vivo (51). However, purified T4 DNA ligase obtained from bacteria that contain cloned T4 gene 30 DNA catalyzes blunt-end joining. Since the enzyme produced in this way cannot be contaminated with T4 RNA ligase, it is clear that this enzyme is not essential for blunt-end joining (52). IV.

Biological Role

A good deal of physiological and genetic information is available concerning RNA ligase, nevertheless its biological function remains obscure. Since RNA ligase activity is first detected about 3 minutes after phage infection (2) it is considered an early protein. In mutants that are defective in DNA replication, RNA ligase accumulates (12) along with several other early proteins. There is evidence (18, 53) that like several other early 50. Lehman, I . R. (1974). 5rienc.r 186, 790. 51. Sugino, A . , Goodman, H. M . , Heyneker, H . L., Shine, J., Boyer, H . M., and Cozzarelli, N. R. (1977). JBC 252, 3987. 52. Murray, N . E., Bruce, S. A . , and Murray, K. (1979). J M B 132, 493. 53. Karam, J . , McCulley, C . , and Leach, M. (1977). Virology 76, 685.

48

OLKE C . UHLENBECK A N D RICHARD I. GUMPORT

proteins, RNA ligase is under control of the w g A gene (54), a posttranscriptional repressor of early functions. Thus, it appears that RNA ligase may be needed for a function early in T4 infection, perhaps in DNA replication or host cell shutoff. Mutants in the RNA ligase gene were first obtained by screening survivors of phage stocks heavily mutagenized with hydroxylamine for RNA ligase activity (8). One of the survivors that induced decreased levels of RNA ligase activity was back-crossed with wild-type T4 and examined in detail. This mutant, A5x4, had about 8% of the wild-type level of RNA ligase activity and showed a completely normal latent period and burst size upon infection of E . coli B. Preliminary mapping of the A5x4 mutant located it in the region of gene 63. It was then noted that bacteriophage with limber- mutations in gene 63 completely lacked RNA ligase activity when grown on nonpermissive cells. Revertants of these limber mutants regained normal RNA ligase activity. Additional genetic tests confirmed that RNA ligase is the product of gene 63. T4 gene 63 has been extensively studied as part of the phage morphogenesis pathway (55-57). Its product (gp 63) is member of an unusual class of proteins that promote phage assembly but are not themselves part of the intact bacteriophage (58).Amber and missense mutants in gene 63 are defective in the attachment of the proximal end of tail fibers to the phage base plate. The attachment of tail fibers is noncovalent (59) and can occur at low rates in the absence of gp 63 (57). The addition of purified gp 63 to extracts infected with a gene 63 amber mutant stimulates the tail fiber attachment rate 50-fold and leads to high titers of viable phage. This assay of tail fiber attachment (TFA) activity has been used to purify gp 63 to near homogeneity (57). Examination of pulse-labeled proteins on SDS gels (58-60) shows that gp 63 is made both early and late in infection. This suggests that the late synthesis of gp 63 serves the purpose of tail fiber attachment, which is the last step in phage morphogenesis. Purified TFA protein and purified RNA ligase are the same protein. They have identical specific activities when measured with both assays (8). The ratio of specific activities remains constant throughout purifica54. Wiberg, J. S . , Mendelson, S . , Warner, V., Hercules, K., Aldrich, C., and Munro, J. L. (1973). J . Virol. 12, 775. 55. Wood, W. B . , and Henninger, M. (1969). JMB 39, 603. 56. Wood, W. B . , and Bishop, R. J. (1973). fn “Virus Research” (C. F. Fox and W. S. Robinson, eds.), p. 303. Academic Press, New York. 57. Wood, W. B . , Conley, M. P., Lyle, H . L . , and Dickson, R. C. (1978). JBC 253, 2437. 58. Vanderslice, R. W., and Yegian, C. D. (1974). Viro/og.v 60, 265. 59. Ward, S . , Luftig, R. B . , Wilson, J. H . , Eddlernann, H . , Lyle, H., and Wood, W. B. (1970). J M B 54, 15. 60. Wiberg, J. S. (1981). Personal communication.

2. T4 RNA LIGASE

49

tion and is similar in several different enzyme preparations. The two proteins comigrate on SDS-gel electrophoresis. Thus, the same gene product has two apparently distinct activities. It is unlikely that the TFA activity involves the ligation of RNA molecules. The attachment of the tail fibers to the base plate is noncovalent and no phosphate is present in any of the T4 structural proteins (61). Furthermore, tail fiber attachment does not require ATP or magnesium ion, which are both necessary for RNA ligase activity. Addition of 1 M (NH&S04 completely inhibits RNA ligase but stimulates TFA activity sevenfold (8).Finally, one umber mutant (MN23), which maps close to the carboxyl-terminus of the protein, shows residual TFA activity and no detectable RNA ligase activity. Thus, RNA ligase and TFA are two independent activities of the same protein. Only the TFA activity of gp 63 is important for T4 infection in wild-type E. coli. Infection of E. coli B with umber mutants in gene 63 show normal protein synthesis, phage DNA replication, and packaging (55). Only the tail fiber attachment reaction is affected. In extracts of gene 63 amber mutants, tail fiber attachment can occur effectively upon addition of TFA protein under conditions where RNA ligase is not active (55, 57). Thus, any putative early RNA ligase function can apparently either be carried out by cellular enzymes or is not strictly needed for phage infection. The unrelatedness of the RNA ligase and TFA activities has been confirmed (62). A number of missense mutants, termed ipk mutants, were isolated by their inability to grow on E. coli CTrSx, a hybrid strain derived from E. coli K12 and a clinical isolate, E. coli CT196 (63).Theipk mutants map in gene 63 but unlike most other gene 63 mutants, grow normally on E. coli B or K, showing normal tail fiber attachment. Since extracts of E. coli B infected with several different ipk nutants show no detectable RNA ligase activity (62), it appears that RNA ligase is indeed not essential for T4 infection of wild-type cells. However, since all mutants in gene 63 are unable to grow on E. coli CTrSx, there is a clear correlation between lack of growth on E. coli CTrSx and lack of RNA ligase activity. The availability of a restrictive host has also allowed the relative placement of the ipk and other gene 63 mutants on the T4 genetic map (Fig. 3). Infection of the nonpermissive E. coli CTrSx by an ipk mutant results in an early interruption of T4 development. T4 DNA replication is greatly inhibited, and the DNA that is made is of lower molecular weight (62). Possibly as a result of the reduced DNA replication, little or no synthesis of late proteins is observed as well. This early defect arising from an RNA 61. Dickson, R . C. (1973). J M B 79, 633. 62. Runnels, J . , Soltis, D., Hey, T., and Snyder, L. (1982). J M B , in press. 63. Depew, R. E., and Cozzarelli, N. R. (1974). J . Vivol. 13, 888.

50

OLKE C . UHLENBECK AND RICHARD I. GUMPORT

/

p I

N R

h Nk

z z

63- RNA I igase 43,000

alclunf

I

57.1% aJ,w

2 2 2 c 2," I ' "

"

'

Pset-Kinase/ Phosphotase

? NY

33,000

-

FIG.3. Location of the T4 RNA ligase and polynucleotide kinase-3'-phosphatase genes. The various mutations are arranged in their correct positions, but no map distances are available. [Data taken from Ref. (62) and (65). The designation ipk has recently been changed to rli (62).]

ligase mutation is consistent with the early appearance of RNA ligase after infection. It is not known why E. coli CTrSx cannot support an infection of T4 that is defective in RNA ligase activity. One simple explanation is that wildtype E. coli have an activity, missing or reduced in the CTrSx strain, that can perform the same function as RNA ligase and thus permit T4 infection. This view is supported by the fact that if a strongamber suppressor is introduced into E. coli CTrSx, it is no longer restrictive for any T4 ipk mutant (62). This suggests that E. coli CTrSx contains anamber mutation in the putativeE. coli activity that replaces RNA ligase. More evidence for a substitutingE. coli activity is indicated by the isolation of several missense mutants of E. coli B, which are phenotypically very similar to E. coli CTrSx (64). These mutants map at two loci, 1itA and litB, and require further characterization. The biochemical identity of the missing CTrSx function (or lit function) remains obscure. Despite an intensive search by several groups, no RNA ligase activity has been detected in E. coli extracts. Another class of T4 mutants, called PseT mutants, show many similarities to ipk mutants (63,65).PseT mutants cannot grow on E. coli CTrSx or 64. Cooley, W., Sirothen, K., Green, R., and Snyder, L. (1979). J . Bucteriol. 140, 83. 65. Sirotkin, K . , Cooley, W., Runnels, J . , and Snyder, L. R. (1978). J M B 123, 221.

2. T4 RNA LIGASE

51

E. coli &A but do grow normally on E. cnli B. PseT infection of E. coli CTrSx or lirA results in a similar inhibition of DNA replication and absence of late proteins. Introduction of an umber suppressor into E. coli CTrSx also permits a T4 PseT infection. Finally, a T4 extragenic suppressor of PseT mutants, called srp ( 6 3 , also suppresses ipk mutations (64). The T4 PseT gene maps very close to the ipk gene (Fig. 3), probably separated from it only by the alciirnf gene (66, 67). PseT is known to be the structural gene for T4 polynucleotide kinase3‘-phosphatase (63, 65, 68). This enzyme has two activities. Polynucleotide kinase catalyzes the transfer of the y-phosphate of ATP to the 5‘ terminus of DNA or RNA (69). The 3’-phosphatase activity removes phosphates specifically from the 3’ terminus of DNA and, less effectively, RNA (68, 70). Both activities are probably necessary for physiological function since neither PseTI, a mutant missing only the 3’-phosphatase activity (63, 65), nor PseT47, a mutant missing only the polynucleotide kinase activity (65), is able to grow on E. coli CTrSx. Furthermore, a mixed infection of PseTI and PseT47 will not grow on E. coli CTrSx even though both kinase and phosphatase activities are present in the infected cells. It is therefore possible that both activities must be present on the same protein molecule for biological function. The function in E. coli that permits T4 PseT mutants to grow and is missing in the CTrSx strain or the lirA mutant is again unknown. Although 3’-phosphatases have been demonstrated (70) no polynucleotide kinase activity has been found in E. coli. No reduction in the major E. coli 3’-phosphatase is observed in E. coli litA or CTrSx (64). Both genetic and biochemical data suggest that RNA ligase and kinase-phosphatase are part of the same biochemical pathway. The strikingly similar phenotype and host range of ipk and PseT mutants suggests that the same pathway in T4 infection may be interrupted by defects in either enzyme. Since both enzymes carry out reactions at the 5’ and 3‘ termini of nucleic acids, it is not difficult to imagine that they might act sequentially. For example, polynucleotide kinase could phosphorylate a 5’ terminus and dephosphorylate a 3’ terminus of an RNA (or DNA) in preparation for inter- or intra-molecular joining by RNA ligase. The in vivo substrates of these enzymes remain unknown. The substrate specificities provide few clues. RNA ligase prefers 3’-terminal ribonucleotides and 66. Snustad, D. P., Tigges, M. A . , Parson, K . A . , Borsch, C. J. H., Caron, F. M., Koerner, J. F., and Tutas, P. J. (1976). J . Virol. 17, 622. 67. Snyder, L., Gold, L., and Kutter, E. (1976). PNAS 73, 8098. 68. Cameron, V., and Uhlenbeck, 0. C. (1977). Bioclwmisre 16, 5120. 69. Richardson, C. C . (1981). “The Enzymes,” 3rd ed., Vol. XIV, p. 299. 70. Becker, A., and Hurwitz, J . (1967). JBC 242, 936.

52

OLKE C. UHLENBECK AND RICHARD I. GUMPORT

shows no preference at the 5’ terminus. Kinase-phosphatase prefers 3‘terminal deoxyribonucleotides and shows no preference at the 5’ terminus. It is interesting to note that a single E. coli function, supplied by the lirA gene, is apparently able to complement defects in both the T4 RNA ligase and the T4 polynucleotide kinase-3’-phosphatase genes. Two general models have been suggested for the biological role of RNA ligase. The first suggests that host RNA modification is involved ( 7 1 ) . This is supported by the fact that several host RNAs that are specifically labeled by polynucleotide kinase can react with RNA ligase in permeable TCinfected cells. The identity and function of these RNAs remains unclear. The second model suggests that T4 DNA replication is involved. This is supported by the fact that aberrant DNA replication is observed in nonpermissive infected cells that do not contain RNA ligase. Both models are at present incompletely substantiated. V.

Applications

Applications of T4 RNA ligase have been in two general areas, the synthesis of oligonucleotides and the modification of RNA molecules. An enzymatic method to join single-stranded oligonucleotides is an important addition to the available chemical and enzymatic procedures for oligomer synthesis. This is especially the case for oligoribonucleotide synthesis where RNA ligase is extremely efficient and chemical methods are not as well developed as they are for oligodeoxyribonucleotide synthesis. Oligonucleotides of defined sequence are important for a wide variety of applications in molecular biology (72). In addition, physical measurements on synthetic oligonucleotides are useful for obtaining a better understanding of the structure of DNA and RNA (73) and of the thermodynamics of nucleic acid interactions (74). RNA ligase can be used to modify RNA molecules by adding nucleotides to either terminus or by replacing nucleotides in internal positions in the polynucleotide chain. The extension of RNA molecules at the 3’ terminus is a useful method for labeling RNA in vitro and allows determination of sequence and secondary structure in the neighborhood of the label. Replacing or adding nucleotides to RNA molecules permits studies that relate their structure to their function. The remarkably broad substrate specificity of RNA ligase should permil 71. David, M . , Vekstein, R., and Kaufmann, G. (1979). P N A S 76, 5430. 72. Itakura, K . , and Riggs, A. D. (1980). Science 209, 1401. 73. Borer, P. N., Kan, L. S.,and Ts’o,P. 0. P. (1975). Biochernisfry 14, 4847. 74. Borer, P. N., DengIer, B., Tinoco, I., and Uhlenbeck, 0.C . (1974). JMB 86, 843.

2. T4 RNA LIGASE

53 TABLE 111

NUCLEOSIDE 3',5'-BISPHOSPHATES ACTIVEWITH RNA LIGASE" Uridine and derivatives

Cytidine and derivatives

Adenosine and derivatives

Guanosine and derivatives

Uridine Deoxyuridine 2'-O-Methyluridine 5-Bromouridine 5-Bromodeoxyuridine 5-Fluorouridine 5-Iodouridine Dihydrouridine Pseudouridine 4-Thiouridine 3-Methyluridine Deazauridine

Cytidine Deoxycytidine 2'-O-Methylcytidine c-Cytidine 5-Iodocytidine

Adenosine Deoxyadenosine €-Adenosine /in-Benzoadenosine NR-Hexylaminoadenosine Purine

Guanosine Deoxyguanosine 2'-O-Methylguanosine Inosine 2-Aminopurine 1-Methylguanosine c-Guanosine p-Guanosine

"

Taken from Refs. (34) and ( 3 6 ) .and from unpublished data of N. Pace and W. Wittenberg.

subtle future applications in RNA synthesis and modification. A wide variety of base-modified, and several sugar-modified, nucleoside 3' 3bisphosphates have been shown to be active donors with RNA ligase (Table 111). In fact, no examples of inactive 3' ,5'-bisphosphates have been found. By incorporation of these donors into oligonucleotides or RNA molecules, a series of RNAs with single, defined chemical modifications can be prepared. These molecules can be used for detailed structurefunction studies. In addition, since many @-substituted ADPs are also substrates for RNA ligase, a variety of complex nonnucleotide groups can be attached to the 3' terminus of oligomers or RNA molecules. Several examples of these are shown in Fig. 4 (see Refs. 45, 47, 75). Thus the potential for RNA ligase to introduce an almost unlimited variety of chemical complexity into RNA chains should permit a better understanding of the structure and function of RNA.

SYNTHESIS A. OLIGONUCLEOTIDE The strategy for the construction of long oligonucleotide fragments from shorter ones by using RNA ligase generally involves developing a highly branched synthetic pathway. Short oligomers made by chemical (76) or 75. Hecht, S. M., Alford, B. L., Kuroda, Y.,and Kitano, S. (1978). JBC 253 4517. 76. Koester, H. (ed.), (1980). "Nucleic Acids Synthesis: Applications to Molecular Biology and Genetic Engineering," Nicclric Acids Re.s.. S y m p . Ser., No. 7. Information Retrieval, Ltd., London.

54

OLKE C. UHLENEECK AND RICHARD I. GUMPORT

R-0-P-u-r-u

FIG.4. P-Substituted ADPs used in applications of RNA ligase. 1,4, and 5 are from Ref. (4S), 2 is from Ref. ( 4 7 ) , and 3 is from Ref. (7.5). The R’ groups of 3 are the side chains of valine, threonine, isoleucine, and phenylalanine.

enzymatic (77) methods are joined to make intermediate-size oligomers, which are subsequently joined to make longer oligomers. A branched pathway not only makes the most efficient use of the smaller precursor oligomers, but also permits the convenient preparation of variant sequences that differ from the parent by one or more nucleotides. The choice of which positions to ligate is governed by two considerations. First, the substrate specificity of RNA ligase dictates that acceptors with uridine residues at or near the 3‘ terminus should be avoided and that donors with 5’-terminal pyrimidines be chosen, although the latter choice is less critical. Second, since a series of similar oligomers will generally be made, joining points are chosen so that the interesting variants can be constructed with a minimal number of changes in the total pathway. For example, if one desires a series of RNA molecules that differ only in the modification of one of the nucleotides in the sequence, it is best to design 77. Thatch, R. E. (1966).1n “Procedures in Nucleic Acid Research” (G. L. Cantoni and P. R. Davis, eds.), Vol. I, p. 520. Harper and Row, New York.

2. T4 RNA LIGASE

55

the synthetic pathway such that the important nucleotide is added as a nucleoside 3‘,5’-bisphosphate as late as possible in the pathway. Ideally, each variant would be made by only one or two alternative synthetic steps using identical precursor oligomers . In order to make the most efficient use of oligonucleotides, a given intermolecular joining reaction is best carried out at approximately equal concentrations of donor and acceptor. A removable blocking group on the 3‘ terminus of the 5’-phosphorylated donor molecule is needed to prevent donor cyclization, donor dimerization, or sequential additions of donors to the acceptor. The product of such an intermolecular reaction is an oligomer with a 5’-terminal hydroxyl and a blocked 3‘ terminus. This product can be either 5’-phosphorylated with T4 polynucleotide kinase (69) to form a new donor molecule, or deblocked to form a new acceptor molecule. Thus, synthesis can proceed in either direction. The choice of the donor blocking group is an important one. The use of a 3’-terminal phosphate appears to have several advantages. It is present on oligonucleotides prepared by ribonucleases and can easily be removed by phosphomonoesterases. Although T4 polynucleotide kinase has an associated 3’-phosphatase activity that could remove the blocking group (68), the kinase from the PseTl. mutant lacks this activity (78). However, the reversal of RNA ligase at 3’-phosphorylated termini (Section II1,D) severely limits the use of phosphates as blocking groups. The AMP released during the course of the forward reaction is sufficient to promote reversal at the 3’ terminus of the donor or the product and therefore lead to removal of the blocking group and to the formation of a variety of undesirable products (48). Several other removable 3’-blocking groups have been developed that would avoid these difficulties (30, 47, 79). In joining reactions where the donor is a comparatively poor acceptor, no blocking of the 3‘ terminus is needed (80). The conditions for optimal intermolecular joining vary for each donoracceptor pair. Since the goal of most reactions has been to obtain the intermolecular product in reasonable amounts, systematic studies of reaction conditions have not usually been reported. However, in one carefully studied reaction, joining could effectively be carried out at oligomer concentrations as high as 1 mM with modest amounts of enzyme (32). Several large RNA fragments have been made with the aid of RNA ligase. By joining smaller fragments made by chemical methods the entire 78. Cameron, V., Soltis, D., and Uhlenbeck, 0. C. (1978). Nircleic Acids Res. 5, 825. 79. Ohtsuka, E., Nishikawa, S . , Markham, A. F., Tanaka, S., Miyake, T., Wakabayashi, T., Ikehara, M . , and Sugiura, M. (1978). Biochemistrv 17, 4894. 80. Collaboration Group of Nucleic Acids Synthesis. (1978). Sci. Sin. 21, 687.

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75 nucleotides of E. coli tRNAP' (79, 81 -83), the 41-nucleotide 3'-half of yeast tRNAAla(84), and several decameric fragments of bacteriophage MS2 RNA (85, 86) have been prepared. Using small oligomers made by enzymatic methods, a 21-nucleotide fragment of bacteriophage R17 RNA (87), a 15-nucleotide fragment of a viroid RNA (88), and several 13nucleotide model anticodon loops (89) have been made. RNA ligase reaction yields varied substantially (5-95%) in these syntheses. However, it may be possible to improve the yields in many cases by a systematic attempt to optimize each particular reaction. RNA ligase can also be used to join oligodeoxyribonucleotides (6, 33, 41 ); however, since deoxyribonucleotides are such poor acceptors, much higher enzyme concentrations and longer incubation times are required for successful joining. Conditions of deoxynucleotide joining have been carefully optimized. MnZ+is more effective than Mg2+in the reaction and low temperatures are required (6, 33). In addition, careful regulation of the ATP concentration in the reaction is needed. Because of the poor reactivity of oligodeoxyribonucleotide acceptors, the third step of the reaction mechanism is rate-limiting and large amounts of adenylylated donor accumulate in reactions that contain high levels of ATP. Furthermore, since high ATP concentrations reduce the amount of free, unadenylylated enzyme needed for the third step in the reaction, the overall reaction is extremely slow. To provide an optimal concentration of both adenylylated donor and free enzyme, oligodeoxyribonucleotide joining reactions are run at low ATP concentrations in the presence of an ATP regeneration system (6, 41 1. These precautions are not necessary in the oligoribonucleotide joining reactions since RNAs are much better acceptors. 81. Ohtsuka, E., Nishikawa, S . , Fukumoto, R., Uemura, H., Tanaka, T., Nakagawa, E., Miyake, T., and Ikehara, M. (1980). EJB 105, 481. 82. Ohtsuka, E., Markham, A. F., Tanaka, S . , Miyake, T., Nakagawa, E., Wakabayashi, T., Taniyama, Y . , Fujiyama, K . , Nishikawa, S . , Fukumoto, R., Uemura, H . , Doi, T., Tokunaga, T., and Ikehara, M. (1980). Nircleir Acids R e s . , Symp. Ser. N o . 7 , p. 335. 83. Ohtsuka, E . , Doi, T., Uemura, H . , Taniyama, Y . , and Ikehara, M. (1980). Nitcleic Acids R e s . 8, 3909. 84. Wang, T. P. (1980). N r d e i c Acids Re.s., Symp. Ser. N o . 7 . 325. 85. Neilson, T., Kofoid, E. C . , and Ganoza, M. C. (1980). Nircleic Acids R e s . , Symp. Ser. N o . 7 , p. 313. 86. Neilson, T., Gregoire, R. J., Fraser, A. R., Kofoid, E. C . , and Ganoza, M. C. (1979). EJE 99, 429. 87. Krug, M., deHaseth, P., and Uhlenbeck, 0. C . (1982). In preparation. 88. Seliger, H., Haas, B., Holupirek, M . , Knable, T., Todling, G . , and Phillipp, M. (1980). Nircleic Acids R e s . , Symp. Ser. N o . 7 , p. 191. 89. Studencki, A . , and Uhlenbeck, 0. C. (1982). In preparation.

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B. NUCLEIC ACID MODIFICATION The 3' termini of most RNA molecules are active acceptors in the RNA ligase reaction (40, 90). By introducing radioactivity into a short donor molecule, usually a 3',5'-bisphosphate such as pCp, a convenient method for in vitro radiolabeling of RNA is achieved. Although the label is usually introduced as [5'-"'PlpCp, other isotopes can be incorporated as well (42). The product of the reaction is an RNA molecule one nucleotide longer with a label near the 3' terminus. RNA ligase labeling therefore complements "'P-labeling of the 5' terminus of RNA with polynucleotide kinase. RNA molecules labeled at the 3' terminus can be used to determine the sequence of the RNA by either the enzymatic (91) or chemical (92) rapid gel-sequencing methods. It should be noted that if [5'-32P]pCpis used to introduce the label, an ambiguity in the sequences of the shorter fragments is often seen due to the internal position of the [32P]phosphate.This can be avoided by the use of [3'-P3'P]pGpp as a donor (93). In a related application, 3'-terminally labeled RNA can be used to obtain information about the structure of the RNA by partial enzymatic (94) or chemical (95) degradation of the molecule. The convenience of 3' end labeling of RNA by RNA ligase has made this the most widespread application of the enzyme to date. Functional events at the 3' terminus of RNA molecules can also be studied with RNA ligase. One or more successive rounds of periodate oxidation and p-elimination will remove residues from the 3' terminus of RNA. The missing 3'-terminal nucleotides can then be replaced using RNA ligase and the function of the altered RNA molecule examined. These procedures have been used primarily on tRNA (75) and 5 S RNA (96) but larger molecules would be of interest as well. The ATPindependent RNA ligase reaction has been used to "chemically aminoacylate" an incorrect amino acid or amino acid analogue onto the 3' terminus of tRNA (75). 90. England, T. E . , Bruce, A. G . , and Uhlenbeck, 0. C. (1980). "Methods in Enzymology," Vol. 65, p. 65. 91. Lockard, R . E., Alzner-Deward, B . , Heckman, J . , MacGee, J . , Tabor, M. W., and RajBhandary, U. L. (1978). Nifrleic Acids R m . 5, 3 7 . 92. Peattie, D. A . (1979). P N A S 76, 1760. 93. Simoncsits, A. (1980). Niicleic Acids Res. 8 , 4111. 94. Vournakis, J . N . , Celantano, J . , Finn, M . , Lockard, R . , Mitra, T., Pavlakis, G., Troutt, A , , van den Berg, M., and Worst, R . (1981). I n "Gene Amplification and Analysis. Analysis of Nucleic Acid Structure by Enzymatic Methods" (J. G . Chirikjian and T. S. Papas, eds.), Vol. 11, p. 267. ElsevieriNorth-Holland, New York. 95. Peattie, D. A., and Gilbert, W. (1980). P N A S 77, 4679. 96. Stahl, D. A , , Meyhack, B . , and Pace, N . R. (1980). P N A S 77, 5644.

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The 5' termini of most RNA molecules are often not active donors in the RNA ligase reaction, presumably due to their secondary structure ( 4 2 ) . However, if this difficulty is overcome, RNAs can be extended at the 5' terminus. RNA molecules extended at the 5' terminus are expected to be useful in the study of RNA processing enzymes. For example, oligonucleotides attached to the 5' terminus of B . sirhtilis 5 S RNA are removed by the 5 S RNA processing enzyme, RNase M5 (97). This has allowed a study of substrate specificity (96) and the development of a convenient assay for the enzyme. Since it is characteristic of most processing enzymes not to recognize structural elements outside the mature RNA, synthetic substrates for other processing enzymes could be constructed by adding any small oligonucleotide to the 5' terminus of a mature RNA by using RNA ligase. Modification of internal positions of an RNA molecule is also possible using RNA ligase. A procedure is first required to make a specific cleavage in the RNA chain. This can be done either by partial digestion with nucleases or by RNase H-directed cleavage (98).Modifications at the nick or gap in the RNA chain can be carried out using RNA ligase, provided the normal termini of the RNA are blocked or inaccessible to the enzyme. Subsequent joining of the half-molecules would be expected to be quite efficient since the proximity of the two termini to be ligated is ensured by the secondary structure of the RNA chain (29). As an example of internal modification of any RNA, nucleotides 34-37 of yeast tRNAPhehave been replaced by A& (99). The procedure is quite general and any oligonucleotide sequence can be inserted into the anticodon loop for various structure-function studies.

97. Meyhack, B . , Pace, B . , Uhlenbeck, 0. C . , and Pace, N . (1978). P N A S 75, 3045. 98. Donis-Keller, H. (1979). Nliclric Acids Res. 7, 179. 99. Bruce, A. G . , and Uhlenbeck, 0 . C. (1982). Bioc.lzemi.stry, in press.

Section II

RNA Polymerases and Related Enzymes

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Bacterial DNA-Dependent RNA Polymerases MICHAEL J. CHAMBERLIN

1. Background . . . . . . . . . . . . . . . . . . . . . . . . . . . 11. Molecular Properties . . . . . . . . . . . . . . . . . . . . . . .

A. Purification . . . . . . . . . . . . . . . . . . . . . . . . . B. Enzyme Assay . . . . . . . . . . . . . . . . . . . . . . . . C. Structure of the Enzyme . . . . . . . . . . . . . . . . . . . D. Other Molecular Properties . . . . . . . . . . . . . . . . . . 111. Catalytic Properties . . . . . . . . . . . . . . . . . . . . . . . A. Variety of Reactions Catalyzed by Bacterial RNA Polymerase . . B. Outline of the DNA-Directed Reaction . . . . . . . . . . . . .

I.

61 64 64 68 14 81 82 82 84

Background

Transcription of genetic sequences in bacteria is mediated by DNAdependent RNA polymerase. The enzyme catalyzes the initiation, elongation, and termination of polyribonucleotide chains that employ ribonucleoside triphosphates as substrates. The synthetic reaction shows an absolute requirement for a divalent metal ion, and normally requires the presence of DNA or a polydeoxyribonucleotide to serve as a template in the reaction. Incorporation of nucleotidyl residues from ribonucleoside triphosphates into an RNA-like material was reported in 1959 by Weiss and Gladstone in rat liver nuclei (I). The bacterial enzyme was identified in 1. Weiss, S ., and Gladstone, L. (1959).JACS 81, 4118.

61 THE ENZYMES,VOL. XV Copyright @ 1982 by Academic Press. Inc. All rights o f reproduction in any form reserved. ISBN 0-12-122715-4

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several laboratories shortly thereafter and was shown to be DNAdependent (2-6). The enzyme from E. coli has been most extensively studied, although the RNA polymerases of other bacteria including species of Azotobacter, Bacillus, Pseudomonas, Micrococcus, and Cauiobacter have also been characterized (7). DNA-dependent RNA polymerase has been found in all bacterial species where it has been sought and its distribution, taken with its sensitivity to drugs that inhibit bacterial transcription, indicates that it is the enzyme responsible for transcription in the bacterial cell. The bacterial RNA polymerases are large molecules (molecular weights between 400,000 and 500,000) and have complex subunit structures. Preparations of the enzyme generally contain two kinds of active molecules (8): core RNA polymerase, which is catalytically active but is unable to interact normally with template promoter sites; and RNA polymerase holoenzyme, which contains an additional subunit, sigma, which is a specificity factor that determines the promoter specificity of the enzyme. Bacterial cells can contain more than one kind of sigma factor (9-11), hence there are multiple RNA polymerase holoenzyme species (12). 2. Hurwitz, J., Bresler, A., and Diringer, R. (1960). BBRC 3, 15. 3. Stevens, A. (1960). BBRC 3, 92; (1961). JBC 236, PC43. 4. Ochoa, S., Burma, D. P., Kroger, H., and Weill, D. (1961). P N A S 47, 670. 5 . Weiss, S. (1960). PNAS 46, 1020. 6. Chamberlin, M., and Berg, P. (1962). P N A S 48, 81. 7. Burgess, R. R. (1976). In "RNA Polymerase" (R. Losick and M. Chamberlin, eds.), p. 69. Cold Spring Harbor Laboratory, Cold Spring Harbor, New York. 8. Burgess, R. R., Travers, A. A., Dunn, J., and Bautz, E. K. F. (1969). Nature (London) 221, 43. 9. Haldenwag, W. G., and Losick, R. (1980). PNAS 77, 7000. 10. Haldenwag, W. G., Lang, N., and Losick, R. (1981). Cell 23, 615. 11. Wiggs, J. L., Gilman, M. Z., and Chamberlin, M. J. (1981) PNAS 78, 2762. 12. The existence of multiple RNA polymerase species in bacteria leads to certain problems in nomenclature. The term RNA polymerase or RNA polymerase holoenzyme is used to designate the form of enzyme containing the predominant form of sigma, and this is referred to simply as sigma subunit. The sigma subunits of RNA polymerases from different bacterial species can differ considerably in size (see Section C), but all of them seem to dictate recognition of a class of related promoter sequences (Wiggs, J . L., Bush, J. and Chamberlin, M. J. (1979). C ~ l l16, 97.) A species of RNA polymerase containing a minor bacterial or phage-coded sigma factor that dictates recognition of a distinct class of promoters is designated by reference to the apparent molecular weight of that factor (e.g.,B. subrilis a2"-RNA polymerase or a*"polymerase), or by reference to the gene coding for that factor (e.g., SPOl phage uypdw RNApolymerase or ugprN polymerase). This form can also be used to refer to the predominant species of sigma factor, sigma subunit (e.g., B . subrilis We originally proposed that species of bacterial RNA polymerase bearing minor sigma factors be designated hdoenzyme 11, 111, etc. (ll), but this nomenclature is confusing since the core polymerase is the same for the different polymerases.

3. BACTERIAL DNA-DEPENDENT RNA POLYMERASES

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The overall enzymatic reaction catalyzed by bacterial RNA polymerase consists of an intricate series of steps in which the enzyme locates sites on the DNA template, initiates an RNA chainde n o w , extends the chain, and finally terminates the chain and is released from the template. Whereas purified RNA polymerase holoenzymes acting alone can carry out all of these steps in v i m , there are additional protein factors involved in the reaction in vivo. Some of these play well-defined roles in a particular step, such as rho termination factor (13), however, it is likely that there are other factors, still unknown, involved in assuring the specificity and efficiency of the transcriptional reaction in v i v a In addition to carrying out a precise complementary copying of the DNA template, the enzyme appears to recognize a variety of genetic signals on the bacterial chromosome and translates the signals into the biochemical events involved in the processes of DNA site selection, RNA chain initiation, RNA chain termination, and enzyme release. Many of the regulatory processes in the bacterial cell appear to be linked directly to transcription, either through direct interaction of the bacterial RNA polymerase with genetic sequences or metabolites, or through interaction of the enzyme or DNA template with accessory factors. This chapter neglects the regulatory properties of bacterial RNA polymerases and focuses primarily on the isolation and structure of the enzymes. A variety of recent reviews are available that deal with RNA polymerase and its role in bacterial transcription. These include several general reviews of enzyme specificity and its role in regulation (14-171, as well as more specialized reviews on the genetics of RNA polymerase and regulation of RNA polymerase synthesis (18-2O), enzyme structure, isolation, and reconstitution (7, 21, 22), promoter and terminator specificity and interactions (23-32), catalytic mechanisms (33-35), and inhibitors 13. Roberts, J. (1969). Nature (London) 224, 1168. 14. Chamberlin, M. (1974). Annu. R e v . Biochem. 43, 721. 15. Chamberlin, M. (1976).In “RNA Polymerase” (R. Losick and M.Chamberlin, eds.), p. 17. Cold Spring Harbor Laboratory, Cold Spring Harbor, New York. 16. Doi, R. (1977). Bacteriol. Rev. 41, 568. 17. Travers, A. A. (1978). In “Biochemistry of Nucleic Acids 11” (B. F. G. Clark, ed.), Vol. 17, p. 233. Univ. Park Press, Baltimore, Maryland. 18. Yura, T., and Ishihama, A. (1979). Annu. Reis. Genet. 13, 59. 19. Scaife, J. (1976). I n “RNA Polymerase” (R. Losick and M. Chamberlin, eds.), p. 207. Cold Spring Harbor Laboratory, Cold Spring Harbor, New York. 20. Matzura, H. (1980). Cuw. Topic F Cell R e g . 17, 89. 21. Ishihama, A. (1980). Advcrtt. Biop/zys. 14, 1 . 22. Zillig, W., Palm, P., and Heil, A. (1976). In “RNA Polymerase” (R. Losick and M. Chamberlin, eds.), p. 101. Cold Spring Harbor Laboratory, Cold Spring Harbor, New York. 23. Gilbert, W. (1976). I n “RNA Polymerase” (R. Losick and M. Chamberlin, eds.), p. 193. Cold Spring Harbor Laboratory, Cold Spring Harbor, New York.

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(36, 37). A monograph dealing with RNA polymerases has recently appeared (38)as well as symposia dealing with selected aspects of transcription (39, 40). II. Molecular Properties

A. PURIFICATION The method of preparation is unusually important for bacterial RNA polymerases since the subunit content, specificity, purity, degree of contamination by inhibitors, and the fraction of RNA polymerase protein that is active can all vary with different methods sE isolation. A detailed review of different fractionation procedures has been presented by Burgess (7); earlier methods were reviewed previously in this series (41). For the E. coli RNA polymerase an efficient procedure based on the precipitation of the enzyme with polyethyleneimine (polymin P) was introduced by Zillig el a/. (42) and has been modified by Burgess and Jen24. Losick, R . and Pero, J. (1976). In “RNA Polymerase” (R.Losick and M. Chamberlin, eds.), p. 227. Cold Spring Harbor Laboratory, Cold Spring Harbor, New York. 25. Chamberlin, M. (1976). In “RNA Polymerase (R. Losick and M. Chamberlin, eds.), p. 159. Cold Spring Harbor Laboratory, Cold Spring Harbor, New York. 26. Siebenlist, U., Simpson, R. B., and Gilbert, W. (1980). Cell 20, 269. 27. Roberts, J. R . (1976). In “RNA Polymerase” (R.Losick and M. Chamberlin, eds.), p. 247. Cold Spring Harbor Laboratory, Cold Spring Harbor, New York. 28. Adhya, S., and Gottesmann, M. (1978). Annu. Rev. Biochem. 47, %7. 29. Rosenberg, M., and Court, D. (1980). Annu. Rev. Genet. 13, 319. 30. Bujard, H. (1980). TIES 5, 274. 31. Platt, T. (1981). Cell 24, 10. 32. Losick, R., and Pero, J . (1981). Cell 25, 582. 33. Krakow, J., Rhodes, G., and Jorm, T. M. (1976). I n “RNA Polymerase” (R. Losick and M. Chamberlin, eds.), p. 127. Cold Spring Harbor Laboratory, Cold Spring Harbor, New York. 34. Mildvan, A. S., and Loeb, L. (1979). CRC Crif. Rev. 6, 219. 35. Scheit, K . (1979). I n “Antimetabolites in Biochemistry, Biology and Medicine” (J. Skoda and P. Langen, eds.), p. 127. Pergarnon, New York. 36. Riva, S . , and Silvestri, L. G. (1972). Annu. Ret,. Microbial. 26, 199. 37. Horwitz, S . (1974). FP 33, 2281. 38. Losick, R., and Chamberlin M., (eds.). (1976). “RNA Polymerase” Cold Spring Harbor Laboratory, Cold Spring Harbor, New York. 39. Osawa, S ., Ozeki, H., Uchida, H., and T. Yura (eds.) (1980). “Genetics and Evolution of RNA Polymerase tRNA and Ribosomes.” Univ. of Tokyo Press, Tokyo. 40. Rodriguez, R., and Chamberlin, M. (eds.). (1982). “Promoters, Structure and Function”. Praeger, New York. 41. Chamberlin, M. (1974). “The Enzymes” Vol. X, p. 333. 42. Zillig, W., Zechel, K., and Halbwachs, H. (1970). Hoppe-Seyler’s Z. Physiol. Chem. 351, 221.

3.

BACTERIAL DNA-DEPENDENT RNA POLYMERASES

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drisak (43). The procedure appears to be quite reproducible and gives fractions that contain the polymerase subunits p’, p, a, and u together with the w polypeptide as the only major polypeptide components. The total recovery ofE. coli RNA polymerase based on analysis of the p’ + p content of cell extracts and final fractions is quite good. The use of polymin P also has proved very effective in obtaining highly purified RNA polymerases from a large number of other bacterial species, including Mycobacteria, Rhodospirillum, Salrnonellu, Bacillus, and Clostridia, as well as T6infected E. coli and the blue-green alga, A. nidulans ( 4 4 4 7 , 47cr). The fact that these different RNA polymerases behave similarly in certain fractionation steps probably reflects their common subunit structure, large size, and DNA binding properties (7). It should be cautioned, however, that the actual solution conditions needed to stabilize the different kinds of bacterial RNA polymerases can vary quite substantially (7, 44, 47).

Certain specific fractionation steps have also proved valuable in fractionation of different bacterial RNA polymerases, for separating particular contaminants, or for separating different forms of the enzyme. These include chromatography on DNA cellulose or DNA agarose (7, 48) and on Blue Dextran columns (49). Noteworthy here is heparin-agarose column chromatography (50), which can give a rapid and efficient purification of the normal B. subrilis RNA polymerase (51) as well as the B. subtilis uZ8 polymerase ( 1 1 ), The exact method of preparation of the heparin-agarose appears to be quite critical (51). For rapid preparation of small amounts of E. coli RNA polymerase from many bacterial strains, a modified form of the polymin procedure has been suggested (52). A similar efficient preparation method employs stepwise elution of the enzyme from a heparin-agarose column (51) after adsorption directly from a cell extract (53).These methods are especially suitable for analysis of RNA polymerases from mutant strains for alterations in struc43. Burgess, R. R., and Jendrisak, J . J . (1975). Biochemistry 14, 4634. 44. Wiggs, J . , Bush, J . , and Chamberlin, M. (1979). Cell 16, 97. 45. Schachner, M . , and Seifert, W. (1971). Hoppr-Sey/er‘,y2. PA.vsiol. Chem. 352, 734. 46. Herzfeld, F., and Zillig, W. (1971). EJB 24, 242. 47. Stetter, K., and Zillig, W. (1974). EJB 48, 527. 47a. Murray, C . , and Rabinowitz, J . C. (1981). JBC 256, 5153. 48. Alberts, B., and Herrick, G . (1971). “Methods in Enzymology,” Vol. 21D, p. 198. 49. Hailing, S . , Sanchez-Anzaldo, F., Fukuda, F., Doi, R., and Meares, C. F. (1977). Biochemistry 16, 2880. 50. Sternbach, H., Englehardt, R., and Lezius, A. G . (1975). EJE 60, 51. 51. Davison, B., Leighton, T., and Rabinowitz, J. C. (1979). JBC 254, 9220. 52. Gross, C., Engbaek, F., Flammang, T., and Burgess, R. (1976). J . Bacteriol. 128, 382. 53. Chamberlin, M., Gilman, M . , and Kingston, R . (1982). “Methods in Enzymology,” in preparation.

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MICHAEL J . CHAMBERLIN

ture or transcriptional properties when used with gel electrophoresis and transcriptional analysis (44). Most purification procedures give RNA polymerase fractions that contain both core polymerase and RNA polymerase holoenzyme; that is, sigma is not present in stoichiometric amounts. Furthermore, it is now clear that, at least in B. subtilis, there are multiple sigma factors, although one species (a”) predominates (9-11, 32). These sigma factors combine with B. subtilis core polymerase to give several distinct kinds of RNA polymerase holoenzymes with different promoter specificities. Since the transcriptional properties, particularly the promoter specificity, of core RNA polymerase and an RNA polymerase holoenzyme are quite different, it is frequently desirable that holoenzyme be prepared free of core polymerase. Chromatography on single-stranded DNA agarose columns (54) or on phosphocellulose in 50% glycerol (55) separates the E. coli enzyme into fractions containing predominently holoenzyme and core polymerase, respectively, as judged by SDS gel analysis of the fractions. However the presence of modest amounts (<5%) of core polymerase in the holoenzyme fractions would not be detected by most assay procedures, and in fact the former method can give holoenzyme fractions containing as little as 0.6 equivalents of a (56). These small amounts of core polymerase can be serious contaminants for experiments intended to study the specific properties of an RNA polymerase holoenzyme. Eschrrichiu coli RNA polymerase holoenzyme can also be reconstituted by adding purified a subunit (56) to partially a-saturated holoenzyme or core polymerase preparations (43). Although reconstitution is rapid, it is not a simple reaction. In the absence of Mg” ion an enzyme form is obtained that differs in its properties from normal holoenzyme (57). Since it cannot be assumed a pviori that holoenzyme reconstituted in v i m is identical to that isolated directly, the possibility of differences should be kept in mind if reconstituted holoenzyme is used. Escherichiu coli core RNA polymerase is usually obtained by phosphocellulose or Bio-Rex-70 chromatography, which releases the u subunit from holoenzyme (56). However, there are generally small traces (< 1%) of a in these preparations, and because u can recycle during transcription a major part of the activity of such fractions can be due to holoenzyme when duplex DNA templates are used (58, 59). This has given rise to 54. Nusslein, C., and Heyden, B. (1972). BBRC 47, 282. 55. Gonzalez, N . , Wiggs, J . , and Chamberlin, M. (1977). ABE 182, 404. 56. Lowe, P., Hager, D., and Burgess, R . R . (1979). Biochemisrry 18, 1344. 57. Fisher, R . , and Blumenthal, T. (1980). JBC 255, 11056. 58. Tjian, R., Stinchcomb, D., and Losick, R. (1974). JBC 250, 8824. 59. Kingston, R., and Chamberlin, M. (1981). Unpublished studies.

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several reports that core RNA polymerase can show promoter-specific transcription. Eschrrichirr coli core polymerase can be freed of the last traces of u by chromatography on poly(rC) cellulose columns (53, 5 9 ) ; these enzyme fractions show no detectable transcription from the strong T7 A promoters, but transcribe actively from single-strand breaks and ends. The release of E. coli u subunit from RNA polymerase fractions upon chromatography on phosphocellulose or Bio-Rex-70 columns provides a useful way of purifying that factor (7, 5 6 ) . Escherichiri coli u is a very stable protein and can also be isolated in active form directly by eluting a-containing bands from SDS-polyacrylamide gels (60, 6 1 ) . For bacterial RNA polymerases other than the E. coli enzyme, isolation of holoenzyme, core polymerase, or sigma is often quite a different matter. For several other bacterial RNA polymerases the predominant u subunit is not released by chromatography on phosphocellulose (62, 6 3 , and this is also true of several of the minor B. sirbtilis u subunits (64, 6 5 ) . This makes isolation of these factors or of core polymerases difficult, and there are no general methods known. However, like the E. coli sigma subunit, the minor B. siihtiliJ u factors can be eluted in active form from SDS-polyacrylamide gels (9-11); this is a highly useful method for confirming the identity of such factors. Some bacterial RNA polymerases are quite unstable and dissociate even during purification procedures (such as column chromatography) that are perfectly suitable for the E . coli RNA polymerase (44, 47). This can severely limit the choice of purification steps for a particular RNA polymerase. Because of the structural complexity of the bacterial RNA polymerase molecules, and because of the intricate set of reactions they catalyze, it is not advisable to characterize a preparation simply on the basis of enzymatic activity and protein content [see Section II,B and Ref. ( 6 6 ) ] .Instead it is important to monitor all fractions by SDS-polyacrylamide gel electrophoresis (7) and determine the content of active and specific RNA polymerase by using an appropriate quantitative assay ( 6 6 ) if one is available. In particular, it is possible to obtain preparations that are highly homogeneous as judged by protein analysis by using SDS gels, which have a very low fraction of the active enzyme, or show reduced efficiency in specific transcription cycle steps. 60. Weber, K . , and Kutter, D. (1971). JBC 246, 4505. 61. Hager, D., and Burgess, R. (1980). A n d . Biochem. 109, 76. 62. Johnson, J . , Debacker, M . , and Boezi, J . (1971). JBC 246, 1222. 63. Amemiya, K . , Wu, C. W., and Shapiro, L. (1977). JBC 252, 4157. 64. Haldenwag, W., and Losick, R . (1979). Notrtre (London) 282, 256. 65. Jaehning, J . , Wiggs, J., and Chamberlin, M . (1979). P N A S 76, 5470. 66. Chamberlin, M . , Nierman, W., Wiggs, J . , and Neff, N . (1979). JBC 254, 10061

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MICHAEL J . CHAMBERLIN

A number of contaminating enzymatic activities can be detected in all but the most highly purified preparations of bacterial RNA polymerases (7). In many instances these activities can interfere with applications of the enzyme or can give misleading information as to its properties. A partial list of important contaminants in this regard are deoxyribonucleases (exo- and endo-), ribonucleases, nucleoside triphosphatases (67, 6 8 ) , polynucleotide phosphorylase (69), polyphosphate kinase (70, 71), and polyadenylic acid polymerase (72, 73).

B. ENZYME ASSAY 1, Qualitative and Qrrantitative Assay Procedures

While the homogeneity of a purified RNA polymerase preparation is easily monitored by sodium dodecyl sulfate gel electrophoresis of the protein, the definition and quantitative determination of enzymatic activity is extremely difficult. The presence of active enzyme in a preparation is revealed by its ability to direct DNA-dependent incorporation of labeled nucleotide into acid-precipitable material. In general, such procedures employ nucleoside triphosphates labeled in the nucleoside residue with 3H or I4C or in the a-phosphate with 32P, and follow their conversion to an acid-insoluble form (RNA) in a 10- or 20-min incubation. These are useful procedures for following total RNA polymerase activity during purification or where the enzyme is used as a preparative reagent. However, because of the complex series of steps that are involved in synthesis of a single RNA chain, the amount of such incorporation generally bears no simple relationship to the fraction of RNA polymerase molecules that are active in a preparation, nor will it normally distinguish between enzyme fractions that may differ greatly in their specificity. In addition, because of the intricacy of the partial reactions carried out by RNA polymerase in the transcription cycle, no single parameter can be used to describe the “activity” of an enzyme preparation. For example, consider two enzyme preparations that contain equal concentrations of active RNA polymerase molecules that have identical rates of site selection, chain initiation, and chain elongation. If one preparation is unable to Paetkau, V., and Coy, G . (1972). C m . J . Biochem. 50, 142. Ishihama, A., Ikeuchi, T., and Yura, J. (1976). J . Biochem. (Tokyo) 79, 917. Kimhi, V., and Littauer, U . (1968). JBC 243, 231. Kornberg, A . , Kornberg, S.,Simms, E. (1956). BRA M , IS. 71. McConnell, D., and Bonner, J. (1972). Biochrniistry 11, 4329. 72. August, J., Ortiz, P., and Hunvitz, J . (1962). JBC 237, 3786. 73. Sippel, A . (1973). EJB 37, 31. 67. 68. 69. 70.

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read termination signals on the template and the second preparation can do so, the two fractions can differ substantially, both in the amount of nucleotide incorporation that they catalyze and in the genetic sequences that are read during transcription. Because of these considerations, studies of in vitro transcription that follow the effect of different reaction conditions, factors, or inhibitors on total nucleotide incorporation generally signify very little except that they show that the RNA polymerase is, or is not, active. This is especially true where complex templates are used (calf thymus, E . coli, T4 DNAs). Similarly, studies of the physical or chemical properties of an RNA polymerase carried out with an enzyme preparation that has not been characterized as to its basic catalytic properties, or under conditions where the enzyme is not known to be active, seem of very limited value. Several groups have described procedures to measure the concentration of active bacterial RNA polymerase molecules in preparations. These include measurements of the number of [y-32P]nucleosidetriphosphates incorporated with DNA templates, where each RNA polymerase initiates only a single RNA chain (74), measurement of covalent addition to the end of DNA chains ( 7 3 , and measurement of the concentration of enzyme by titrating its ability to form template complexes that carry out the abortive initiation reaction (76). There are also specific assays for each of the steps in the transcription cycle [see Ref. (14) for a review]. However, in general, these procedures do not give an overall picture of the transcriptional parameters of an RNA polymerase preparation, and many require specialized reagents. One approach to the quantitative characterization of bacterial RNA polymerase preparations involves the use of a DNA template that bears a single well-defined transcriptional unit, where the values of different transcriptional parameters can be altered by following labeled nucleotide incorporation during a single cycle of transcription. The method requires that template binding and RNA chain initiation be completed in a time period that is short, as compared to the transit time for the transcriptional unit involved. This allows all active RNA polymerase molecules to initiate a chain and begin chain elongation. Under conditions where the mean chain elongation rate is constant and there is no termination of RNA chains or cessation of transcription, the concentration of active RNA polymerase molecules is given by the rate of incorporation divided by the mean chain elongation rate. 74. Nierman, W.. and Chamberlin, M. (1979). JBC 254, 7921. 75. Wickner, S., Hurwitz, J., Nath, K . , and Yarbrough, L. (1972). BBRC 48, 619. 76. Cech, C . , and McClure, W. (1980). Biochemistry 19, 2440.

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The basic features of the procedure have been set forth using bacteriophage T7 DNA and E. coli RNA polymerase (66). Here a single strong transcriptional unit is used during free RNA chain initiation in vitro, and the rates of all of the different steps in the transcriptional cycle are known from separate studies, The procedure gives the amount of active E. cofi RNA polymerase in the preparation together with values of several of the parameters for the different steps of the transcriptional cycle, including (a) the rate of promoter location plus chain initiation, (b) the mean rate of RNA chain elongation, and (c) the efficiency of chain termination at the strong early T7 termination signal. This quantitative assay using T7 DNA as template is rapid, employs commonly available reagents, and can be utilized even with rather impure fractions of RNA polymerase. In addition, the assumptions on which the assay procedure is based also appear to be valid in most cases for bacterial RNA polymerases from a wide range of taxonomically different bacterial strains. This follows from the observation that these diverse bacterial RNA polymerases all transcribe T7 DNA selectively, and generally use the same promoter and terminator sites that are utilized by the E. coli enzyme (12). RNA polymerases from Rhodosporillum rubrum, Mycobacterium smegmatis, Caulobacter crescentus, Azotobacter vinelundii, and Bucillus sirbtilis give single-cycle transcriptional curves with T7 DNA similar to those of the E. coli polymerase, although the exact values of the assay parameters differ significantly among the different enzymes (66). In principle, this kind of quantitative assay for an RNA polymerase can be devised with any template and RNA polymerase for which the transcriptional parameters fill the requirements set forth above. Practically speaking, the major difficulties that must be considered are: (a) The promoter site or sites used must be strong enough to give rapid initiation, and there must be no competing promoter sites or tight-binding sites that cause loss of enzyme. (b) The transcriptional unit must be reasonably long to allow a phase of transcription in which all of the active polymerase is growing chains. The rate of elongation can be reduced by lowering the temperature or the substrate concentrations (77), and this will help lengthen the elongation phase. However, these conditions can also slow promoter complex formation andlor chain initiation (78) and may extend the initiation phase unacceptably. In addition, low nucleotide concentrations can lead to extensive “pausing” of transcriptional complexes and give nonlinear kinetics of chain elongation (79). (c) There must be no ,

77. Neff, N . , and Chamberlin, M. (1980). Biochemistry 19, 3005. 78. Chamberlin, M., Mangel. W., Rhodes, G . , and Stahl, S. (1976). AlfredBenron S y m p . I X . Munksgaard Press, Copenhagen. 79. Kassavetis, G . , and Chamberlin, M. (1981).JBC 256, 2777.

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premature termination. This latter requirement has proved to be a major problem in devising quantitative assay procedures with cloned promoters. For example, the vector pBR322 (80) bears a partially effective transcriptional terminator located near 3100 bp on the standard map of pBR322, which interrupts what would otherwise be a very satisfactory transcription unit read from the tet promoter or from promoters on fragments cloned into the EcoRI site or neighboring sites (81). The existence of partial terminators on templates is especially a problem with B. subtilis RNA polymerase, which appears to respond much more strongly to such sites than the E. coli enzyme (77). It is often important to determine the amount of RNA polymerase holoenzyme in a preparation that contains a mixture of holoenzyme and core polymerase. This can be done using T7 DNA and a rifampicin challenge procedure (66), or by titrating the enzyme for its ability to form complexes that are able to carry out abortive chain initiation with dAT (82). 2. Defectiori and Asscry qf RNA Polyrna.rrses o j Novel Spec8citg The quantitative assay procedures discussed above generally depend on having an enzyme of defined promoter specificity. However the specificity and the efficiency of utilization of promoter and terminator sites can be changed for different forms of bacterial RNA polymerase (8-11), or even for RNA polymerase mutants (77, 83). Hence the investigator is often faced with the problem of determining whether an RNA polymerase fraction has the specificity of the normal E. coli holoenzyme or is significantly different. This information can be decisive either in allowing purification or RNA polymerases of novel specificity or in characterization of purified fractions. Ideally one would like an assay procedure that gives a quantitative measurement of the amount and specificity of each distinct RNA polymerase species present in a fraction. In practice, the specificity of such enzyme species is usually not known a priori and such assays are not available. Also, there is often a large background of transcription by other forms of RNA polymerase in the extract. One approach to this problem is to monitor the transcriptional specificity of RNA polymerase fractions by transcribing well-defined templates and analyzing the RNA transcripts by polyacrylamide gel electrophoresis. This is most effective where there is a promoter site on the template that is 80. 81. 82. 83.

Sutcliffe, J . G. (1979). CSHSQB 43, 77. Gilman, M., and Chamberlin, M. (1981). Unpublished observations. Hansen, U . , and McClure, W. (1980). JBC 255, 9556. Yanofsky, C., and Horn, V. (1981). J . Bacterial. 145, 1334.

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MICHAEL J. CHAMBERLIN

coupled with a strong, rho independent terminator to give a transcription unit of well-defined size that can be sharply resolved by gel analysis (44). Alternatively the DNA template can be cleaved at a site downstream from the promoter, using an appropriate restriction endonuclease. This gives a discrete “run off” transcript terminated at the cleavage site [see for example, Refs. (64,84-87)]. Although this procedure generally does not give a quantitative analysis of the amount of RNA polymerase specific for a given promoter, it can be highly effective as a qualitative test for such an activity. Advantages of the procedure are that the presence of a promoter-specific enzyme can often be picked up against a large background of nonspecific transcription, or transcription from different promoters. This of course depends on the size and strength of the transcriptional unit. Short transcription units (up to 500 n) are advantageous since the resolution of the gel system is better for smaller than for larger fragments, and even the presence of small amounts of contaminating nucleases may not preclude detection of the transcript. Furthermore the procedure need not depend on a priori knowledge of the promoter location (10). An extension of this approach involves using a template that bears a collection of different promoter sites with different transcriptional properties. Here, by following transcription from each promoter, one can monitor not only large changes in promoter specificity (promoter recognition identity) but also detect minor changes that affect the relative utilization of promoters that share a common specificity (promoter strength). Such a promoter test system has been described using bacteriophage T7 DNA as template (44). Here in addition to the strong early promoter sites, there are several weaker promoters that function effectively in vitro, yet differ substantially in their actual DNA sequences, their rates of transcription, and in the biochemical properties of their RNA polymerase promoter complexes. With the deletion mutant T7 D111, only one of the strong T7 A promoters remains on the template, and transcripts from this site, which terminate at the T7 early terminator (18.1%), are easily resolved from those initiated at the weaker T7 C, D, and E promoters. By using appropriate transcription protocols involving prebinding of enzyme to DNA or .free initiation of transcripts with limiting RNA polymerase, one can distinguish between RNA polymerases that differ from one another in promoter recognition identity or in promoter strength interactions, respectively. 84. Meyer, B., Kleid, D., and Ptashne, M. (1975). P N A S 72, 4785. 85. Gilbert, S., de Boer, H . , and Nomura, M . (1979). CeII 17, 211. 86. Young, R., and Steitz, J. (1979). CeI/ 17, 225. 87. Davison, B . , Murray, C . , and Rabinowitz, J. (1980). JBC 255, 8819.

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This analysis has been used quite effectively to show that the predominant forms of RNA polymerase holoenzyme in a large variety of true bacteria share a common promoter recognition identity, but can differ strikingly in the strength of initiation with particular promoter sites (44). Similarly, a form of B . sirhtilis RNA polymerase that shows a novel promoter recognition identity with this test system was purified and shown to have a unique structure, involving a new sigma-like subunit that confers the altered recognition properties ( I I , 65). Two other useful procedures to detect and analyze transcription from specific promoters or transcription units should be mentioned. The presence of transcripts derived from a particular region of a DNA template in a mixture of in vitro transcripts can be detected by cleavage of the template using appropriate restriction endonucleases, followed by gel electrophoresis to separate DNA fragments, and hybridization of labeled transcripts to the fragments by the Southern method (88). Alternatively, RNA chains can be terminally labeled with [y-32P]nucleotidesand subsequently cleaved with an appropriate ribonuclease to generate a mixture of 5’-y-32P-labeled oligonucleotides. These can be separated and the amount of transcription from particular promoter sites can be analyzed (89). Both of the above procedures are applicable even where there is enough ribonuclease or deoxyribonuclease in the enzyme fractions to preclude gel analysis of the intact RNAs. However each has its own weaknesses. Random transcription by core polymerase will be detected in the first procedure and must be distinguished from selective transcription. The use of y-32P-labelingrequires rather active polymerase fractions and high specific activity nucleotides to get sufficient incorporation to study, and the presence of ATPases or phosphatases can pose problems. Not all changes in transcriptional specificity of an RNA polymerase involve promoter-polymerase interactions. Alterations can potentially affect the rate and efficiency of RNA chain initiation, RNA chain elongation (90), and RNA chain termination and release (77,831. Hence, there is no general assay procedure for detecting all such changes. Some of these changes can be detected using quantitative assay procedures with templates like T7 DNA (66). There are also test systems to screen in vitro for changes in RNA polymerase-terminator efficiency analogous to the promoter test systems (77, 90), and Yanofsky has devised an imaginative in vivo system for a similar test (83). 88. Southern, E. M. (1975). J M B 98, 503. 89. Miller, J . , and Burgess, R. (1978). Biocherrrisfr-v 17, 2054. 90. Neff, N., and Chamberlin, M. (1978). JBC 253, 2455.

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C. STRUCTURE OF THE ENZYME 1. Subunits and Subunit Structure of the Escherichia coli Enzyme Escherichia coli RNA polymerase holoenzyme as normally isolated contains four species of polypeptide chain designated p’ , p, u,and a in order of decreasing size. These are all functional subunits of the enzyme since mutants are known that alter each polypeptide and affect cellular transcription (18-20). In addition, all four components are required to reconstitute active holoenzymein vitro from separated components (21,22). There is good evidence that E. coli core polymerase binds tightly to an E. coli protein specified by the nusA gene (91) and that nusA is involved in transcriptional elongation and termination in vivo. Because of its properties the nusA protein should probably be considered a subunit of elongating E. coli RNA polymerase (91) (see discussion in Section II,C,3). Escherichia coli RNA polymerase preparations also often contain significant amounts of a smaller polypeptide (about 8000- 10,000 MW) referred to as w (8, 92). This can be removed without affecting any of the known properties of the enzyme, and is not required for reconstitution of active holoenzyme (21, 22). It is, therefore, not known whether w is involved as a transcriptional component in v i i ~ oor is an adventitious contaminant. The E. coli genes for the four subunits of RNA polymerase holoenzyme have been mapped and cloned, and complete DNA sequences are available for rpoD (ugene) (93) and rpoB ( p gene) (94, 9 3 , as well as partial sequences for rpoA (96) and rpoC (97). These sequences give exact size and molecular weight for u (613 amino acids, MW 70,263) and p (1342 amino acids, MW 150,619). The complete amino acid sequence of a (329 91. Greenblatt, J . , and Li, J. (1981). Cell 24, 421. 92. Berg, D., and Chamberlin, M. (1970). Biochemistry 9, 5055. 93. Burton, Z., Burgess, R., Lin, J . , Moore, D., Holder, S., and Gross, C. (1981). Nticleic Acids Res. 9, 2889. 94. Ovchinnikov, Yu., Monastryskaia, G., Gubanov, V., Guriev, S., Chertov, O . , Modyanov, N . , Grinkevich, V., Markova, I . , Marchenko, T., Polovnikova, I., Lipkin, V.,and Sverdlov, E. (1980). Dokl. Acad. N m k . USSR 253, 994. 95. Ovchinnikov, Yu., Monastryskaia, G . , Gubanov, V., Guriev, S., Chertov, O . , Modyanov, N . , Grinkevich, V., Markova, I . , Marchenko, T., Polovnikova, I., Lipkin, V., and Sverdlov, E. (1981). EJB 116, 621. 96. Post, L., and Nomura, M. (1979). JEC 254, 10604. 97. Squires, C., Krainer, A., Barry, G., Shen, W.-F., and Squires, C. L. (1981). Nitclric Acids Res. 9, 6827. 98. Ovchinnikov, Yu., Lipkin, V. A . , Modyanov, N., Chertov, O . , and Smirnov, Yu. (1977). FEES Lett. 76, 108.

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amino acids, MW 36,512) is also known (98). The correct molecular weights are close to those originally determined by SDS-gel electrophoresis (7) except for (T, which is much smaller than expected from its mobility (56). In addition to the four normal polypeptide subunits, RNA polymerase contains two atoms of ZnzC,bound in the p’ subunit (34). This is thought to play a functional role in the catalytic action of the enzyme and is generally found in other RNA polymerases as well (34). The protomeric form of RNA polymerase holoenzyme has the composition P’Pa2a(8. 92). Since the P’ subunit has a MW of about 160,000 (7), the MW for the E. coli holoenzyme is quite close to 454,000, which is consistent with the value of 380,000 determined for core polymerase by sedimentation equilibrium ( 9 2 ) . The RNA polymerase protomer can aggregate to form dimers and higher multimers (92, 99), and this aggregation is greatly favored at lower salt concentrations. Hence the question is raised as to the nature of the active unit. Quantitative studies of the efficiency of RNA chain initiation (74) and quantitative assays of the number of active polymerase molecules in different preparations (66) leave no doubt that each protomer is a functional unit. Electron microscopy of RNA polymerase bound at a variety of promoter sites also shows clearly that the enzyme binds as a protomer (100-102). Finally, although detailed studies of the aggregation reaction have not yet been done, it appears that preparations of pure RNA polymerase holoenzyme that lack significant amounts of core polymerase aggregate very little, even under conditions (low ionic strength) that favor dimerization, and electron microscopy of such preparations at protein concentrations below 20 pg/ml reveals no dimers or higher aggregates [see, for example, Refs. (100, 103)l. The organization of the subunits in RNA polymerase holoenzyme has been probed by crosslinking (104), by protease digestion (57, 105, 106), low-angle X-ray scattering (107), antibody binding (108), and studies of enzyme assembly from isolated subunits [for reviews, see Refs. (21, 2 2 ) ] . Studies by Ishihama and his collaborators ( 2 1 ) have shown that RNA 99. Richardson, J . P. (1966). P N A S 55, 1616. 100. Williams, R. C. (1977). P N A S 74, 2311. 101. Hirsch, J., and Schleif, R. (1976). J M B 108, 471. 102. Kingston, R., Gutell, R., Taylor, A . , and Chamberlin, M. (1981). JME 146, 433. 103. Kadesch, T., Williams, R. C., and Charnberlin, M. (1980). J M B 136, 79. 104. Hillel, Z., and Wu, C . (1977). Biochemistry 16, 3334. 105. Lill, H . , and Hartmann, G . (1975). W E 54, 45. 106. King, A , , Lowe, P., and Nicholson, B. (1974). Biochem. Soc. Trans. 2, 76. 107. Meisenberger, O., Pilz, I . , and Henmann, H. (1980).FEES L e f t . 120, 57. 108. Stender, W. (1980). Nircleic Acids Res. 8, 1405.

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polymerase is assembled in vitro and in vivo in an ordered sequence of reactions involving four main steps. 2a

+ a2

a2

i-

P

P'Pa2

--

+

ffza + P'

+0

Pa2

P'PS P'P'a*ff

The sequence suggests that a is likely to be present as a dimer and makes significant contacts with p; these conclusions are confirmed by crosslinking studies (104). Cross-linking studies also reveal important close contacts between p and p' and between u and both p and p' (104, 106), and these have led to a general model for the subunit organization in the holoenzyme (104). A series of studies has been carried out on the small angle X-ray scattering properties of the E. coli holoenzyme and isolated subunits and subassemblies; these studies have also led to a suggested model for subunit organization and shape (107). The role of each of the subunits in the enzymatic reaction has been studied in some detail, and is reviewed in several articles (16, 18, 21, 22, 26. 32). 2. Subunits and Subitnit Structure of Other Buctericil RNA Polymeruses RNA polymerases from all of the normal bacteria studied, as well as blue-green algae (46) and streptomyces (109), appear to contain subunits homologous to the E. coli p', p, u,and a subunits and share a common protomer structure (7, 16, 22). The homology between comparable subunits is most clearly shown by the reconstitution of active RNA polymerase from heterologous mixtures of subunits. Sigma subunits of different bacterial polymerases are generally interchangeable (44, 110109. Jones, G. (1978). BBRC 84, 962. 110. Whiteley, H., and Hemphill, H. (1971). BBRC 41, 647. 111. Shorenstein, R., and Losick, R. (1973). JBC 248, 6170. 112. Lill, U., Behrendt, E., and Hartmann, G. (1975). EJB 52, 411. 113. Fukuda, R., Ishihama, A., Saitoh, T., and Taketo, M. (1977). hlnl~c..Gcn. Gmcr. 154, 135. 114. Gragerov, A., and Nikiforov, V. (1980). FEBS Lett. 122, 17. 115. Zillig, W., Stetter, K . , and Tobien, M. (1978). EJB 91, 193. 116. Zillig, W., Stetter, K . , and Janekovic, D. (1979). EJB 96, 597. 117. Halling, S., Burtis, K., and Doi, R. (1977). JBC 252, 9024. 118. Ruger, A,, and Ruger, W. (1980). FEBS Lett. 120, 233. 119. Tjian, R., and Pero, J. (1976). Nature (London) 262, 753. 120. Duffy, J., and Geiduschek, E. P. (1977). Nuture (Lotidun) 270, 28. 121. Talkington, C . , and Pero, J. (1978). PNAS 75, 1185.

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122). This is also true of the core subunits, although not in all possible combinations (112). A wide variety of bacterial RNA polymerases also cross-react with antibody to the E. coli RNA polymerase subunits ( I 13, 114). Taken together these findings indicate that the structure of bacterial RNA polymerases is highly conserved in evolution; this also appears to be true of the major promoter recognition specificity as well ( 4 4 ) . This conservation of structure and specificity may be a consequence of the need for heterologous genetic exchange in bacterial growth and evolution (44). This homology among bacterial RNA polymerases may not extend to bacterial species that are classified as Archybacteria, such as thermophiles and halophiles. Here highly purified RNA polymerase preparations contain up to 10 polypeptide components, and are of sizes and in molar ratios that appear quite different from the normal /3‘/3a2upolymerase structure (115, I f 6 ) . More detailed studies of the structures of these enzymes and their properties is needed to establish the composition of the actual active RNA polymerase. The molecular weights of the core polymerase subunits from most true bacteria are generally close to those of the E . cnii polymerase as judged by mobility on SDS-polyacrylamide gels (7), although the B. sirbtilis enzyme differs from the E . coli enzyme in that the p subunit is larger than the /3‘ subunit (117). The size of the major sigma subunit, however, is quite different for gram-positive bacteria, including Bacillus ( II I ), Lactobacillus (47), and Clostridiirm (44) species, which have apparent MW values of 55,000, 44,000, and 60,000, respectively, by SDS gel electrophoresis. It should be noted that most published MW values for sigma subunits estimated from SDS gel electrophoresis are likely to be in error for several reasons. First, the mobility of E . coli sigma has proved to be much lower than expected for its known molecular weight of 70,000 (93), and early estimates of the size (86,0OO-95,000) were therefore much too high. Second, the actual mobility obtained for sigma can be quite different for different gel buffer systems ( 5 6 ) . This anomalous electrophoretic behavior may be due to some general structural feature of sigma subunits (56), but is probably not due to the presence of carbohydrate since sigma does not seem to be a glycoprotein (118).Third, many workers have used E . coli RNA polymerase subunits as standards for SDS gel measurements of subunit sizes, assuming MW = 96,000 for E. coli sigma (8) which is much larger than appropriate for its apparent mobility in most gel systems (7). All of these considerations cast a great deal of uncertainty on values for 122. Whiteley, H . , Spiegelman, G . , Lawrie, J . , and Hiatt, W. (1976). I n “RNA Polymerase” (R.Losick and M. Chamberlin, eds.), p. 587. Cold Spring Harbor Laboratory, Cold Spring Harbor, New York.

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both subunit and protomer molecular weights for RNA polymerases other than E. coli, and most of the values should be considered only as rough estimates. RNA polymerase from B. subtilis has been extensively studied both as an example of a gram-positive RNA polymerase, and because of the interest in regulation of transcription during the differentiation processes of endospore formation (16,32). In addition to the difference in subunit sizes noted above, the B. subtilis enzyme has several novel features not currently found for the E. coli enzyme. In addition to the predominant B. subtilis sigma subunit (a55), which confers a specificity for promoter recognition homologous to other bacterial polymerases, there are three or more B. subtilis polypeptides that can replace a55and confer a distinct and novel promoter recognition specificity on the resulting holoenzyme (811). Other distinct polypeptides replace B. subrilis as5 after infection with SPOl or SP82 bacteriophages, and also switch the specificity of the enzyme to recognition of new classes of bacteriophage promoters (119-122). These modifying polypeptides have been termed “sigma factors” because of their functional homology with the sigma subunit. Two sigma factors, a’’ and a3’,are found in vegetatively growing B. subtilis (9, 11). These factors are present in low amounts relative to sigma subunit ($7, and the levels do not change appreciably during the initial stages of endospore formation (10, 123). Synthesis of at least one sigma factor (a’? is induced early during sporulation ( l o ) ,and there is evidence for yet another such factor of distinct specificity appearing somewhat later (124).

It is not yet known that the minor sigma factors are determined by genes but distinct from that of 1~~~rather than by cleavage or modification of ass, the specificity of their action strongly suggests that they are different proteins. The a’* polymerase, for example, does not use promoter sites used by the cr55polymerase, and the 8*promoter sites show quite different sequences from promoter sites in regions associated with control of polymerase-promoter interaction (125). This is also true of promoters used by B. subtilis a3’polymerase (126) and B. slrbtilis polymerase bearing the SPOl gene 28 sigma (127). This directly implicates the sigma factor in binding and recognition of specific promoter sequences (125-127).

123. 124. 125. 126. 127.

Wiggs, J . , and Chamberlin, M. J. (1981). Unpublished observations. Losick, R. (1981). Personal communication. Gilman, M., Wiggs, J . , and Chamberlin, M. (1981). Nrrcleic Acids. R e s . , 9 , 5991 Moran, C., Lang, N . , and Losick, R . (1981). Nrrdeic. Acids R e s . , 9 , 5979. Lee, G., and Pero, J . (1981). J M E 152, 247.

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It is likely that some of the B. sirbtilis minor sigma factors play important roles in regulation of transcription during endospore formation since they control transcription of sporulation specific sequences ( l o ) ,and there may be additional roles for these factors in other cellular processes as well. Minor sigma factors that confer altered promoter specificity analogous to those of B. sirbtilis have not yet been reported for other bacterial species (128). Bmillirs sirhtilis RNA polymerase preparations normally also contain substantial amounts of a polypeptide designated delta (8) of MW 21,000 (129, 130). Delta can be removed by chromatography on phosphocellulose and is not required for catalytic activity or promoter recognition by B. subrilis RNA polymerase. However, S does have a substantial effect on transcription by the polymerase, and is very likely to be a functional subunit of the B. subtilis polymerase. Delta appears to increase the accuracy of transcription by various forms of the B. sirbtilis enzyme (130, 131) and can reduce binding by a5’polymerase at both specific and nonspecific sites (132). A possible role for delta is suggested by the finding that E. coli a 7 0 acts both to dictate specific promoter recognition and to suppress nonspecific DNA binding by a very large factor (14). Since B. subtilis a55and delta together are roughly the size ofE. coli a’”, it might be imagined that in B. subtilis the two functions of E. coli r 7 O have been separated onto different polypeptides (130). This would be advantageous if the 6 function of suppressing nonspecific binding were essential, and if B. subtilis transcription involved a number of different sigma factors, which is the case (32). However assignment of a definite function for delta must await isolation of mutants and appropriate in ~ ~ i vstudies. o

3. Trunscriptioncil Fucrors and Missing Srhrnits Purified RNA polymerase holoenzyme is the only component required for accurate initiation and termination of certain phage transcription units (for reviews, see 14, 16, 17). However for other phage and many bacterial transcription units, there is genetic and sometimes biochemical evidence that other protein factors are required for transcription in vivo, usually in 128. A second form of E . coli cr has been reported and designated u’but the promoter specificity has not been shown to differ from the normal bacterial u. See Fukuda, R., Iwakura, Y., and Ishihama, A. (1974). J M B 83, 353. 129. Pero, J . , Nelson, J . , and Fox, J. (1975). P N A S 72, 1589. 130. Tjian, R . , Losick, R . , Pero, J . , and Hinnebush, A. (1977). EJB 74, 149. 131. Spiegelman, G . , Hiatt, W., and Whiteley, H. (1978). JBC 253, 1756. 132. Whiteley, H . , Achberger, E., and Hilton, M . (1982). I n “Promoters, Structure and Function” (R. Rodriguez and M . Chamberlin, eds.). (1982). Praeger, New York.

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addition to the normal components of holoenzyme. In some cases these additional components bind tightly and specifically to RNA polymerase and can reasonably be considered to be subunits of the polymerase. Examples include phage and bacterial sigma factors (9-1 1, 119-122), E. coli nusA protein (91), and B. subtilis delta protein (129-132). In other instances, such as the E. coli catabolite regulatory protein CAP (133-1351, nraC protein (136) and rho protein (28, 29), the association or interaction with RNA polymerase is more transient, or may not even be direct. These components should probably be considered simply as transcriptional factors, rather than subunits. One can set several criteria for judging whether a newly identified component is a functional part of the bacterial transcription apparatus (41). Ideally, the criteria should require that (a) the component should be purified and shown to be distinct from known polymerase subunits or transcriptional factors; (b) the component should be required for some step in selective transcription in vitro (in the case of a new subunit, for example, reconstitution of the enzyme in the presence or absence of the component should give an activity that possesses or lacks the new specificity, respectively); ( c ) bacterial or phage mutants, or cells grown under conditions where the new component is not observed, should lack the corresponding in vivo function. These are stringent criteria and only a few of the many components believed to be involved in transcription in vivo meet all three; examples are the B. subtilis phage SPOl gene 28 sigma factor (119, 127), E. coli CAP and crraC proteins (133), and E. coli rho protein (28, 29). Other, well characterized transcription factors fill only two of the three criteria. For example, purified bacteriophage A gene N protein is clearly a transcriptional factor by genetic criteria and has been purified to homogeneity (137, 138). However purified N protein is not active in a purified transcription system with E. coli RNA polymerase alone, probably because additional components are needed for its action (139). In this category would also go the B. subtilis minor sigma factors (9-11) and delta protein (129), E. coli nusA protein ( 9 1 ) , and T4 phage gene products 33 and 55 (140). 133. Pastan, I . . and Adhya, S. (1976). Bacteriol. Rev. 40,527. 134. Majors, J. (1975). Nuture (London) 256, 672. 135. Taniguchi, T., O’Neill, M., and decrombrugghe, B. (1979). PNAS 76, 5090. 136. Lee, N . , Wilcox, G . , Gielow, W., Arnold, J . , Cleary, P., and Englesberg, E. (1974). P N A S 71, 634. 137. Greenblatt, J . , Malnoe, P., and Li, J. (1980). JBC 255, 1465. 138. Ishii, S., Sugino, Y.,and Imamoto, F. (1980). Gene 10, 291. 139. Greenblatt, J. (1981). CeN 24, 8. 140. Rabussay, D., and Geiduschek, E. P. (1977). In “Comprehensive Virology” (H. Fraenkel-Conrat and R. Wagner, eds.), Vol. 8, p. 1. Plenum, New York.

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At the other extreme, there are many published reports of “transcriptional factors,” RNA polymerase “subunits,” or “heterogeneity of RNA polymerases” based simply on the ability of a protein fraction to stimulate incorporation of nucleotide in an in vifro assay, or on the binding of a polypeptide to the polymerase as judged by cochromatography. While this type of observation may be consisfenfwith a functional association of a component with RNA polymerase and certainly merits further analysis, it is not a necessciry relationship, and it is inappropriate and misleading to represent it as such. Bacterial RNA polymerases contain a number of binding sites for charged macromolecular ligands, and it should be assumed that a polypeptide bound to polymerase is adventitiously bound until proved otherwise. Nor does an effect of a component on total polymerase activity in vitvo signify a functional interaction; at low protein concentrations bovine serum albumin can give a twentyfold enhancement of E. coli RNA polymerase activity! D.

OTHERMOLECULAR PROPERTIES

As in studies of subunit structure and reaction mechanism, most physical studies of bacterial RNA polymerases have been carried out with the E. coli enzyme. Studies of circular dichroism in the far UV have been reported for the E. coli enzyme and its subunits (141, 142). These studies have allowed estimates of a-helical content and p-sheet structure for holoenzyme of about 43 and 27%, respectively. Sigma seems to be unusually high in a-helical content (55-60%) (93, 141). Ultraviolet spectra of the E. coli RNA polymerase holoenzyme have been combined with measurements of nitrogen content (141) or low wavelength UV absorption (7) to calculate extinction coefficients E A Z n m of 6.7 and 6.2, respectively. A value of 6.7 was obtained by Richardson (9) using the refractive increment method with an enzyme preparation that was probably not saturated with sigma, however core polymerase is reported to have a significantly lower extinction coefficient (7). An average value of 6.5 has been used by some workers for holoenzyme, and it is not clear which of the different values is likely to be more accurate. In view of the importance of this parameter in the determination of the actual concentration of RNA polymerase, it is unfortunate that there has not been a more detailed study. Despite the uncertainty of the exact value for E&$inm,determination of RNA polymerase concentrations by UV absorption is probably the most 141. Levine, 9.. Orphanos, P., Fischman, B., and Beychok, S. (1980). Eioclrrmistry 19, 4808. 142. Ishihama, A . , Aiba, H., Saitoh, J., and Takahashi, S. (1979). Biochemistry 18, 972.

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precise method of obtaining the RNA polymerase protein concentration. However substantial errors can result from light scattering due to aggregation of the polymerase (7), and this is particularly true in the extreme instance of core polymerase at lower salt concentrations. Therefore it is always necessary to determine the absorption at both 280 and at 300-350 nm, and correct using the Leach and Scheraga (143) procedure if there is substantial absorbance above 300. As a general rule, if the absorbance at 320 is less than 2% that at 280, the correction at 280 will be less than 5% (66 ) . RNA polymerase concentrations can also be determined using the Lowry protein method in the BSA as standard. The method should be calibrated using a sample of RNA polymerase for which the concentration is known from UV absorption. In our hands a solution of RNA polymerase having a Lowry protein concentration of 1 mg/ml has a true concentration of 0.8 mg/ml using an extinction coefficient Ed&,,,, of 6.5 [Ref. (66); note that this relationship is incorrectly stated in the reference]. There is an accumulating amount of evidence that bacterial RNA polymerases can exist in a variety of conformational states. This is best shown by the formation of inactive or structurally altered complexes during reconstitution, which can have physical and chemical properties quite different from the starting enzyme (21, 142). Preparations of holoenzyme can contain up to 90% inactive polymerase (66) and there is some evidence that termination efficiency may vary among different preparations as well (66). Because of this variability it seems essential that measurements of basic physical parameters be carried out on preparations that have been fully characterized for homogeneity and composition, and are also fully active in the overall transcription reaction. Measurements that depend on other properties of the enzyme seem far less desirable; for example, the azP subassembly can bind DNA and rifampicin, but is catalytically inert (47, f f 2 ) , and catalytically inactive RNA polymerase may still bind DNA. It is not evident that the physical or biochemical properties of catalytically inactive RNA polymerase are the same as those of fully active, native polymerase. 111.

Catalytic Properties

CATALYZED BY BACTERIAL A. VARIETYOF REACTIONS RNA POLYMERASE Bacterial RNA polymerase carries out several distinguishable kinds of polynucleotide synthesis, as well as a variety of reactions that represent 143. Leach, S., and Scheraga, H. (1960). JACS 82, 4790,

3. BACTERIAL DNA-DEPENDENT RNA POLYMERASES

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steps in the pathway of polynucleotide synthesis. For the purposes of discussion three kinds of overall reactions are distinguished: (1) Template-directed formation of complementary polyribonucleotides. (2) Template-directed formation of homopolymers or repeating polymers (reiteration). (3) Unprimed synthesis of polyribonucleotides. The first reaction is best represented by the DNA-directed synthesis of a complementary polyribonucleotide (RNA). When a helical DNA is employed as template, the product is free RNA and the template DNA is left unaltered after synthesis. The RNA product is precisely complementary to the region of the DNA employed as template and is antiparallel in sequence. In most cases only certain regions of the DNA template are transcribed as a result of the restriction imposed by the location of sites on the DNA required for the initiation and termination of RNA chains (14, 23-32).

The second reaction is observed with single-stranded polyribonucleotides, polydeoxyribonucleotides, or DNA as template, and involves the formation of a polyribonucleotide homopolymer or simple alternating polymer (144, 14.5). The reaction involves the repeated (reiterative) copying of a restricted region of the template, and the product is far longer than the template sequence. Thus, short runs of thymidylate residues in single-stranded DNA serve as templates for synthesis of long chains of poly(rA) (144), and this reaction can also be directed by poly(dT) oligomers as short as (pT), (14.5). Reiterative synthesis of poly(rA) with poly(rUX) templates probably requires a sequence of at least two U residues (146). However, synthesis of poly(rG) is observed with the alternating polymer poly(dTC) as template, where there are no sequences of C residues greater than one in length (147). In the latter instance G-T base pairing (“wobble” pairing) may account for the reaction. Reiterative synthesis of repeating sequences of two or three nucleotides has been observed (148) and could possibly lead to difficulties in the accurate copying of short DNA fragments. Reiterative copying of a template sequence is suppressed when a substrate triphosphate that is complementary to the template base distal to the reiterative sequence is included in the reaction. For example, reiterative synthesis of poly(rA) directed by the sequence TTTTTTC is suppressed by inclusion of GTP in the reaction, even at micromolar concentrations (144). The third reaction involves synthesis of polyribonucleotides by bacte144. 145. 146. 147. 148.

Chamberlin, M., and Berg, P. (1964). J M B 8, 708. Falaschi, A , , Adler, J . , and Khorana, H. (1963).JBC 238, 3080. Adman, R., and Grossman, L. (1967). J M B 23,417. Paetkau, V., Coulter, M., Flintoff, W., and Morgan, A. (1972).J M B 71, 293. Nishimura, S., Jacob, T., and Khorana, H. (1964). P N A S 52, 1494.

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rial RNA polymerase in the absence of an added template or primer. Two major products have been identified: poly(rA) .poly(rU), which is formed when ATP and UTP are added as substrates (149, /50), and alternating poly(rIC), formed when ITP and CTP are used as substrates (151). The unprimed synthetic reaction appears to be an intrinsic property of bacterial RNA polymerase and is not the result of contamination of the enzyme preparations by a bacterial poly(A) polymerase or by polynucleotide phosphorylase, neither of which could account for synthesis of alternating poly(r1C) in any event. Both of the latter enzymes are capable of giving rise to poly(rA) synthesis in RNA polymerase reactions since they are common contaminants of the enzyme. The unprimed synthesis of poly(rA) poIy(rU) and poly(r1C) by bacterial RNA polymerase is associated with homogeneous, electrophoretically purified RNA polymerase and, in fact, the reaction can be employed as an in situ assay for the enzyme in acrylamide gels (152). In addition, inhibitors of bacterial RNA polymerase inhibit unprimed poly(rA) .poly(rU) synthesis (153). The multiple sources of poly(rA) synthetic activity in RNA polymerase preparations have led to some confusion in the literature, and probably in some experimental systems as well. It has been claimed that only E. coli RNA polymerase that bears a novel sigma factor (d) is active in unprimed polynucleotide synthesis (154). However, since core polymerase reconstituted from purified subunits is fully active (112) this is unlikely to be correct. Unprimed synthesis of poly(rA) poly(rU) and poly(r1C) by bacterial RNA polymerase requires Mn2+as divalent metal ion and usually shows a lag period of at least 30 min. Rather high concentrations of enzyme favor the reaction. Attempts to find traces of endogenous primer in enzyme preparations have not been successful (150).

-

B. OUTLINE OF

THE

DNA-DIRECTED REACTION

The DNA-directed synthesis of RNA by bacterial RNA polymerase may be broken down into a number of sub steps. Each of these steps is complex and depends on the nature of the DNA template as well as on the RNA polymerase involved. The commonly accepted steps involved are: 1. Template binding, in which the RNA polymerase attaches to the 149. Mehrotra, B . , and Khorana, H. (1965). JEC 240, 1750. 150. Smith, D., Ratliff, R . , Williams, D., and Martinez, A. (1967). JEC 242, 590. 151. Krakow, J . , and Karstadt, M. (1967). PNAS 58, 2094. 152. Krakow, J . , Daley, K . , and Fronk, E. (1968). BBRC 32, 98. 153. Krakow, J . , and Von der Helm, K . (1970). C S H S Q B 35, 73. 154. Iwakura, Y., Fukuda, R., and Ishihama, A. (1974). J M B 83, 369.

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BACTERIAL DNA-DEPENDENT RNA POLYMERASES

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Negative control factors: repressors Positive control factors: activators RNA Polymerase holoenzyme n -

Sigma factors

-4

Core polymerase

I. Template Binding

d

0

+ ATP+XTP 2. RNA Chain Initiation

4. RNA Chain Termination and Enzyme Release

3. RNA Chain Elongation

Antitermination factors

FIG.I .

The transcription cycle.

DNA and ultimately locates a promoter site at which chain initiation can occur. 2. R N A chuin initiurion, in which the enzyme catalyzes the coupling of a ribonucleoside triphosphate, usually ATP or GTP, with a second ribonucleoside triphosphate to eliminate inorganic pyrophosphate and form a dinucleoside tetraphosphate. 3 . R N A chain elongntion, in which successive ribonucleoside monophosphate residues are added to the 3’-OH terminus of the nascent RNA chain from ribonucleoside triphosphate substrates. 4. RNA chain termination und relense, in which chain elongation halts, and the nascent RNA chain and RNA polymerase are released from the template. The overall synthetic reaction consists of at least one such synthetic cycle, which has been called the Transcription Cycle [Ref. (14), Fig. 11. With helical DNA templates and RNA polymerase holoenzyme, initiation and termination of transcription in tivo and in vitro take place almost

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entirely at discrete DNA sequences, termed promoters and terminators. These define units of transcription, and transcription on such templates is restricted to these regions and is therefore selective (14). Both promoter and terminator sites can vary considerably in their structure and in the efficiency and specificity of interaction with RNA polymerase. Thus the rate and efficiency of interaction of RNA polymerase with these loci is of major importance in determining which genetic sequences are expressed in the cell, and at what rate. There have been extensive studies of the individual steps of the transcription cycle, especially those involved in promoter binding and chain termination. A discussion of the mechanism and specificity of these reactions is not possible in the space available here. The reader is referred to reviews and monographs on promoter structure and binding (15, 23-26, 29, 30), RNA chain initiation (22, 23), RNA chain elongation (33, 35, 79), and RNA chain termination (27-29, 31, 155). With single-stranded DNA templates there is little or no specificity in the sites used for RNA chain initiation, and RNA chain termination occurs randomly throughout the reaction (156) to give rather short RNA chains bonded to the template in a DNA-RNA hybrid (157, 158). The process may well be brought about by formation of the hybrid structure, leading to premature chain termination, since a very similar reaction occurs with poly(dG) poly(dC) template when poly(rG) is synthesized (90). The product of this reaction is a poly(rG). poly(dC) hybrid with displacement of the poly(dG) strand (159). The random initiation-termination transcription process on single-stranded M13 DNA can be suppressed by E. coli single-strand binding protein, leading to selective initiation at a single promoter site (160), however there have been relatively few studies of in vitro transcription with well-defined single-stranded templates.

155. Yanofsky, C. (1981). Nature (London) 289, 751. 156. Maitra, U., Nakada, Y.,and Hurwitz, J. (1967). JBC 242, 4908. 157. Chamberlin, M., and Berg, P. (1%4). JMB 8, 297. 158. Sinsheimer, R., and Lawrence, M. (1964). J M B 8, 289. 179. Chamberlin, M. (1965). Ff 24, 1446. 160. Schaller, H., Uhlmann, A., and Geider, K. (1976). f N A S 73, 49.

4

Bacteriophage DNA-Dependent RlvA Pol'ymeraases M. CHAMBERLIN

T. RYAN

I . Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . 11. T7-Like RNA Polymerases . . . . . . . . . . . . . . . . . . . A. Molecular Properties . . . . . . . . . . . . . . . . . . . .

B. Catalytic Properties . . . . . . . . . . . . . . . . . . . . . 111. Other Bacteriophage RNA Polymerases . . . . . . . . . . . . . .

A. Bacteriophage PBS2 RNA Polymerase. . . . . . . . . . . . . B . Bacteriophage N4 RNA Polymerase . . . . . . . . . . . . . .

1.

87 89 89 98 105 105 106

Introduction

Infection of a bacterial cell by a bacteriophage leads to a progressive reprogramming of the biosynthetic capabilities of the cell toward synthesis of bacteriophage components. An early and fundamental step is the establishment of a regulated transcription program for the bacteriophage genome. Most bacteriophages-with notable exceptions-depend on the host transcriptional machinery for transcription of genes used early in infection. Late bacteriophage transcription, however, can employ either the host RNA polymerase or an independently synthesized RNA polymerase coded for by the bacteriophage genome. This review deals entirely with the latter enzymes. Other related reviews cover phage trans87 THE ENZYMES, VOL. XV Copyright @ 1982 by Academic Press. Inc. All rights of reproduction in any form reserved.

ISBN 0-12-122715-4

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cription involving bacterial RNA polymerases that have been reprogrammed to read phage-specific transcriptional units (14,and the genetics and physiology of T7 and the T7-like bacteriophages, including genetics of the T7-like RNA polymerases and their role in bacteriophage growth and development (4, 5 ) . Viral-coded DNA-dependent RNA polymerases were first identified in mammalian viruses as activities carried in the viral particle (6, 7). Subsequently, an RNA polymerase coded for by T7 bacteriophage and specific for T7 DNA as template (8) was isolated from extracts of infected E. coli. Similar enzymes are induced after infection by a variety of T7-like bacteriophages, and are characterized by having only a single polypeptide chain and by being highly specific for the homologous phage DNA template (4, 9-12). Two other kinds of bacteriophage-specified DNA-dependent RNA polymerases are known. Both are under the control of bacteriophages that can induce active transcription in bacterial cells even in the presence of rifampicin, which blocks transcription by bacterial RNA polymerase. An RNA polymerase activity specified by one of these phages, N4, is carried in the bacteriophage particle, and this enzyme, rather than the E. coli host polymerase, is responsible for transcription of N4 DNA immediately after infection (13, 14). It is a very large protein, consisting apparently of only a single polypeptide chain of molecular weight about 350,000 (15). The structure of another RNA polymerase, induced by bacteriophage PBS2, which grows in Bacillirs sirbfilis, is more complex. The enzyme seems to rilis, is more complex. The enzyme seems to contain at least five phage1 . Losick, R., and Pero, J. (1976). In “RNA Polymerase” (R. Losick, and M. Chamberlin, eds.). p. 227. Cold Spring Harbor Laboratory, Cold Spring Harbor, New York. 2. Losick, R., and Pero. J . (1981). Cell 25, 582. 3 . Rabussay, D., and Geiduschek, E. P. (1977). I n “Comprehensive Virology” (H. Frankel-Conrat and R. Wagner, eds.), Vol. 8, p. 1. 4. Hausmann, R. (1976). Citrr. Tupics Micruhiol. Immirnul. 75, 77. 5. Kruger, D., and Schroeder, C. (1981). Microbid. R e v . 45, 9. 6. Kates, J., and McAuslan, B. (1967). P N A S 58, 134. 7. Baltimore, D., Huang, A., and Stamfer, M. (1970). PNAS 66, 572. 8. Chamberlin, M., McGrath, J., and Waskell, L. (1970). Nufirre (Londun) 228, 227. 9. Dunn, J . , Bautz, F., and Bautz, E. (1971). Nature N e w B i d . 230, 94. 10. Towle, H., Jolly, J., and Boezi, J. (1975). JBC 250, 1723. 11. Korsten, K., Tomkiewicz, C., and Hausmann, R. (1979). J . G e n . Virol. 43, 57. 12. Butler, E. (1978). Ph.D. Thesis, University of California, Berkeley, California. 13. Rothman, Denes, L., and Schito, G. (1974). Virology 60,65. 14. Falco, S . , Vanderlaan, K., and Rothman-Denes, L. (1977). PNAS 74, 520. 15. Falco, S . , Zehring, W., and Rothman-Denes, L. (1980). JBC 255, 4339.

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contain at least five phage-coded components (16, 17), and it does not seem to be involved in the early phase of phage transcription as originally thought. Its role in the physiology of PBS2 infection is still not well understood. This review is divided into sections concerning the T7-like RNA polymerases, about which there is extensive information, and the RNA polymerases from other phages, about which there is somewhat less information. II. T7-Like Bacteriophages

A.

MOLECULAR PROPERTIES

1. Purijication

A number of bacteriophages specify DNA-dependent RNA polymerases similar to the T7 enzyme; these are all morphologically similar to T7 and often show genetic homology as well (4, 11, 18). The T7 RNA polymerase is the most thoroughly studied and will be taken as representative of the general class, although individual phage polymerases may show significant differences. A variety of procedures has been used to purify T7 RNA polymerase, however there is no really satisfactory method that gives high yields of homogeneous and active polymerase. This is primarily due to several factors. The enzyme is rather unstable and loses activity during purification, especially when protein concentrations are low. In addition, there is not a large amount of enzyme induced in infected cells under conditions of wild-type infection. The original method of Chamberlin et al. (8) for purification of T7 polymerase employed streptomycin sulfate precipitation to remove nucleic acids, precipitation and extraction with ammonium sulfate, followed by column chromatography on DEAE-cellulose and phosphocellulose. The peak fractions from phosphocellulose were over 90% T7 polymerase protein as judged by SDS gel analysis (8). However, the yields and specific activities obtained by this method are relatively low and there is often significant variation in the early steps. Niles el al. (19) introduced a 16. 17. 18. 19.

Clark, S . , Losick, R., and Pero, J. (1974). Ncrture (London) 252, 21. Clark, S. (1978). J . Virol. 25, 224. Hyman, R ., Brunovskis, I . , and Summers, W. (1974). Virology 57, 189. Niles, E., Conlon, S., and Summers, W. (1974). Biochemistry 13, 3904.

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modified procedure in which nucleic acids and T7 polymerase are precipitated from the extract with polyethyleneimine, and the T7 polymerase is then extracted from the precipitate with salt. These fractions are then fractionated with ammonium sulfate, and then by column chromatography on phosphocellulose, DEAE-cellulose, and hydroxylapatite, respectively. This procedure gives good yields of enzyme activity through the phosphocellulose step and is quite reproducible (20). These fractions are only of moderate specific activity and contain contaminating peptides, but they give normal amounts of the large T7 transcripts, as measured by RNA gel analysis of the products, and are quite adequate for transcriptional analysis (19, 20) or preparation of specific labeled RNAs. Such fractions have been kept at -20" in 50% glycerol solutions without substantial loss in activity for many years (20). Further purification of these fractions by heparin agarose chromatography gives enzyme of very high specific activity (20), but these fractions have been somewhat less stable, possibly due to the low protein concentrations involved. An alternative modification of the Niles er a / . (19) procedure was reported to give homogeneous T7 polymerase (21), although no yields or specific activities were described. However, subsequent studies indicate that these fractions may be contaminated with a single-strand-specific endonuclease, and gel analysis suggests that as much as 30-50% of the protein can be in peptides other than T7 polymerase (22). Chromatography of these fractions on T7 DNA cellulose may give homogeneous enzyme (22), but no yields are reported and the capacity of the column is said to be quite low. Similar procedures are generally applicable for the purification of T3 RNA polymerase. Bailey and McAllister (23) have isolated the T3 RNA polymerase using the polyethyleneimine procedure through the phosphocellulose step, followed by chromatography on heparin-agarose and phosphocellulose. An alternative procedure for purification of T3 RNA polymerase (24) takes advantage of the fact that the enzyme in cell extracts is easily sedimented with cell debris, probably due to binding to ribosomes (9). It can be eluted from the pellet with salt solutions and subsequently fractionated by column chromatography. The procedure is reported to give reasonable recoveries (-20%) of enzyme of good specific activity 20. 21. 22. 23. 24.

Kassavetis, G., and Chamberlin, M. (1979). J . Virol. 29, 196. Oakley, J., Pascale, J., and Coleman, J . (1975). Biochemistry 14, 4684. Strothkamp, R., Oakley, J . , and Coleman, J. (1980). Biochemistry 19, 1074. Bailey, J., and McAllister, W. (1980). Nucleic Acids Res. 8, 5071. Chakraborty, P., Sarkar, P., Huang, H., and Maitra, U. (1973). JBC 248, 6637.

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(-600,000 unitshg), which gives a single band on SDS-polyacrylamide gels. Isolation of the RNA polymerase from Srrlrnonelki typliimuriiim infected with phage SP6 has proved to be somewhat easier than for the T7 enzyme, due in part to the greater stability of SP6 enzyme (12. 25). After removal of nucleic acids with streptomycin and ammonium sulfate precipitation, the enzyme is chromatographed on phosphocellulose, Blue DextranSepharose, and Bio-Gel P200, respectively, to give a homogeneous protein fraction. This gives yields of up to 30% overall of SP6 RNA polymerase activity and specific activities of 700,000 unitslmg (12, 25). 2. Enzyme Assoy

T7 RNA polymerase is usually assayed by following incorporation of radioactively labeled nucleotide into acid-insoluble material in the presence of T7 DNA as template. The reaction shows an absolute requirement for the four ribonucleoside triphosphates, Mi$+,and T7 DNA (8, 26). T7 DNA can be replaced by other duplex DNA templates that bear specific T7 polymerase promoter sites (see below) or by synthetic polynucleotides such as (dG);(dC),, (dI),.(dC)n, or poly(dC). The rate of synthesis is optimal between pH 7.7 and 8.3. The rate of synthesis is highly sensitive to reaction temperature (26) and falls off rapidly below 37"; there is about a twofold reduction at 30". This may be due, in part, to a requirement for DNA strand separation in a rate-controlling step (27), but the rate falls off nearly as rapidly with single-stranded poly(dC), as template (26 1, suggesting that other steps or temperature-dependent changes in enzyme conformation are also involved. The rate, and also the extent, of T7 RNA synthesis is affected by sulfhydryl reactive agents, such as p-chloromercuribenzoate; hence a thiol, such as dithiothreitol or P-mercaptoethanol, is included in the reaction solution. Similarly, the reaction, especially with early enzyme fractions, shows an enhancement by, or even complete dependence on, the addition of bovine serum albumin (8). This may be due to the high sensitivity of the T7 polymerase to inhibition by polyanionic compounds (28). Under optimal conditions the reaction continues at constant rate for about 20-30 min after a short (10-15 sec) lag. However, this extended period of synthesis involves many cycles of transcription initiation, elongation, and termination for each active polymerase. The longest of the 25. 26. 27. 28.

Butler, E . , and Chamberlin, M. (1982). JBC, in press. Chamberlin, M . , and Ring, J. (1973). JBC 248, 2235. Oakley, J . , Strothkamp, R . , Sarris, R . , and Coleman, J . (1979). Biochemistry IS, 528. Chamberlin, M., and Ring, J. (1973). JBC 248, 2245.

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transcription units controlled by a strong (class 111) T7 promoter is 12,000 bases ( 2 9 ) , corresponding to a transit time for T7 RNA polymerase of 60 sec at 200 nucleotides per sec (30), and the average transit time for class 111 transcriptional units is 20-25 sec. Hence each active T7 RNA polymerase must repeat the transcription cycle about 3 times each min. A consequence of this extensive recycling during transcription with T7 DNA is that the reproducibility of assays falls off rapidly at times over 5- 10 min (J. Ring and M. Chamberlin, unpublished studies). This is probably because T7 RNA polymerase is not highly stable at 3T, and hence small variations in reaction conditions, glassware, etc., affect the lifetime of enzyme released during the recycling process. Since loss of a small fraction of enzyme is multiplied exponentially in recycling, there is a disproportionate effect on the extent of incorporation at longer reaction times. For example, the presence of a factor or condition that slightly destabilizes free T7 RNA polymerase, so that 5% of the free enzyme is inactivated prior to each round of chain initiation, will reduce incorporation in a l0-min assay to 100 x (0.95)30= 20%, since there are about 30 rounds of transcription involved. Because of this, reaction times of no more than 5-10 min should be used. The definition of a unit of T7 RNA polymerase activity has generally been based on measurement of the rate of reaction in a 10-min incubation under specified reaction conditions with T7 DNA as template (8, 28). One unit is the amount of enzyme needed to give a rate of incorporation of 1 nmol of labeled substrate per hour under these conditions. Since T7 RNA polymerase elongates RNA chains at -200 nucleotides per sec at 37" (30),it can be calculated that a homogeneous protein of MW 100,000 involved solely m chain elongation at this rate would have a specific activity of-1.8 x 10' units/mg. However, T7 RNA polymerase requires -10-15 sec to initiate an RNA chain (26), and each active polymerase does so an average of three times per minute. Therefore, the maximum specific activity of fully active T7 RNA polymerase in these assays is probably about 1 x 10' unitdmg. This neglects any time required for release of enzyme at RNA chain termination, and assumes no inactivation of enzyme during the recycling reaction. This maximum value for the specific activity should be compared to the highest specific activities reported of -600,000. From the above considerations it is clear that the normal assay for T7 RNA polymerase, although useful for following the presence of active RNA polymerase during fractionation, does not give a quantitative assay 29. Carter,A., Morris, C., and McAllister, W. (1981). J . Virol. 37, 636. 30. Golomb, M., and Chamberlin, M. (1974). JBC 249, 2858.

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for the molar concentration of active RNA polymerase present. Changes in the activity do not necessarily reflect possible changes in the specificity of the enzyme, or in the rate or efficiency with which it carries out individual steps in the transcription cycle. This is even more true when reaction conditions are altered, inhibitors are present, etc., where it cannot be assumed that the rates or efficiencies of the different steps of the transcription cycle are equally affected. It would be highly desirable to have a quantitative RNA polymerase assay involving a single transcriptional cycle, similar to that developed for bacterial RNA polymerases (31). The requirements for such an assay are set forth elsewhere in this volume (32).It is clear that such an assay for T7 RNA polymerase cannot be devised with T7 phage DNA. It contains far too many transcription units that vary in transit time from about 3 to 60 sec. Since chain initiation, or establishment of a normal rate of chain elongation, requires about 10- 15 sec, it is clearly impossible to separate chain elongation from initiation and termination. Such a separation should be possible with cloned T7 polymerase promoters inserted in large cloning vectors. In principle, if there were no chain termination, and transcription were initiated on an intact circular DNA, RNA chain elongation would be continued more or less indefinitely and a quantitative measure of the concentration of active RNA polymerase could be obtained simply from the rate of incorporation and the elongation rate. Unfortunately, although cloned phage polymerase promoters are available (33),available vectors contain several partially effective terminator sites [Ref. (33),and D. Roulland, unpublished observations]. Hence, although these plasmid DNAs can generate extremely large amounts of RNA, in vitro, there is still the same problem of separating the different reaction steps. In view of the importance of obtaining a quantitative T7 RNA polymerase assay, it would be useful to attempt to develop better DNA templates for these assays by systematically attempting to remove in v i m termination sequences from the cloning vectors. The biochemical properties and assay procedures for other T7-like phage RNA polymerases are generally similar to those of the T7 polymerase, and have been studied for the T3 R N A polymerase (24, 33-37), Pseudomonas phage gh-1 (ZO), and Salmonella phage SP6 (Z2, 25). 31. 32. 33. 34. 35. 36.

Chamberlin, M., Nierrnan, W., Wiggs, J . , and Neff, N . (1979). JBC 254, 10061. Chamberlin, M . (1981). Chapter 3, this volume. McAllister, W., Morris, C . , Rosenberg, A . , and Studier, F. (1981). J M B , 153, 527. Dunn, J . , McAllister, W., and Bautz, E. (1972). EJE 29, 500. McAllister, W., Kupper, H . , and Bautz, E. (1973). EJB 34, 489. Salvo, R., Chakraborty, P., and Maitra, W. (1973). JEC 248, 6647.

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These different enzymes all show a requirement for their own specific class of promoter sites and generally will not use heterologous templates (see Section II,B), However all of the T7-like RNA polymerases studied thus far will use the synthetic polynucleotides (dG),*(dC), or poly(dC) as templates. These templates lack specific promoter sequences; transcription on such templates probably reflects simply the general catalytic activity of the polymerase in a reaction where specific promoter binding has been bypassed.

3. Physical and Chemical Properties Highly purified T7 RNA polymerase preparations contain a single polypeptide chain that is the protein product of T7 gene 1 (8). The mobility of the protein on SDS-polyacrylamide gels corresponds to a MW of 107,000-110,000 (8, 19). No other factors or components appear to be required for enzymatic activity. The active RNA polymerase has a sedimentation coefficient of 5.9 to 6.3 S (8, 1 9 ) , which is consistent with its existing as a monomer of MW -100,000. From these observations it is concluded that the active form of T7 RNA polymerase consists of a single subunit of MW -100,000 (8). The T7 gene coding for T7 polymerase has been cloned and sequenced, giving both the amino acid sequence and size of the T7 polymerase protein (38).T7 polymerase protein contains 883 amino acid residues, corresponding to a MW of 98,092. The amino acid composition is given in Table I. The true molecular weight is significantly lower than that estimated by SDS-polyacrylamide gel electrophoresis. This may be due to inaccuracies in the molecular weights of marker polypeptides in this molecular weight range (32); however, it is more likely that the T7 polymerase protein displays an abnormal mobility in SDS-polyacrylamide gels due to some feature of its structure. The sigma subunit of E. coli RNA polymerase displays a very abnormal mobility and gives an apparent MW of 80,00090,000 (39),although the true MW is close to 70,000 (40). Physical studies of T7 RNA polymerase have been handicapped by lack of availability of large amounts of homogeneous, fully active, protein (see Section II,A, 1). Several studies of physical and chemical properties have been carried out on purified T7 RNA polymerase preparations, however the preparations are likely to have contained significant amounts of con37. Chakraborty, P., Bandyopadhyay, P., Huang, H . , and Maitra, U. (1974) JBC 249, 6901. 38. Stahl, S., and Zinn, K. (1981). J M B 148, 481. 39. Lowe, P.,Hager, D . , and Burgess, R. R. (1979). Biochemistry 18, 1344. 40. Burton, Z., Burgess, R . , Lin, J . , Moore, D . , Holder, S . , and Gross, C. (1981). Nircleic Acids Rcs. 9, 2889.

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TABLE I AMINO ACIDCOMPOSITION OF T7 RNA POLYMERASE~ Amino acid Alanine Arginine Asparagine Aspartic acid Cysteine Glutamine Glutamic acid Glycine Histidine Isoleucine Leucine Lysine Methionine Phenylalanine Proline Serine Threonine Tryptophan Tyrosine Valine Total Molecular weight

Predicted 100

40 37 41 13 32 65 55 21 52 71 63 24 34 37 48 49 18 23 60 883 98,092

" Data are from Ref. (38).

taminating proteins, as well as unknown amounts of enzymatically inactive T7 RNA polymerase, and physical and chemical parameters that would be affected by this contamination are potentially in error. The amino acid composition of one such preparation does not agree well with that determined by DNA sequencing; this is especially true for the amino acids Tyr and Trp ( I Y ) . Values of U and Efgcalculated in these earlier studies (19) should probably be recalculated using the amino acid composition determined from the DNA sequence (38).Analysis of preparations of T3 RNA polymerase (41) and SP6 RNA polymerase (12, 25) using SDSgel electrophoresis shows that these proteins have mobilities slightly greater than that of the T7 enzyme. Hence the true molecular weights of these enzymes are probably slightly lower than those of the T7 enzyme, assuming that the mobilities of these proteins resemble the T7 polymerase. 41. Beier, H . , and Hausmann, R. (1974). NatitrP (London) 251, 538.

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T7 RNA polymerase contains tightly bound Zn", which appears to be required for catalytic activity (42). Other template-dependent nucleotidyltransferases also appear to contain Zn2+(43). It has been reported that addition of Zn2+to some preparations of T7 RNA polymerase enhances T7 RNA polymerase activity (42). The phage RNA polymerases coded by T3 and SP6 phages are cleaved by trypsin to give smaller subfragments that are catalytically active (25, 4 4 ) . In the case of the SP6 enzyme, the activity with SP6 DNA, which depends on specific promoter sites, is lost much more rapidly than the activity with (dI),-(dC),, which does not depend on such sites (25). Therefore, it appears that a much smaller subfragment of the phage RNA polymerase can carry out the catalytic functions of the enzyme. What parts of the phage RNA polymerase protein are involved in the biochemical reactions that specify each of the steps in synthesis of an RNA chain? In the case of the bacterial RNA polymerases, at least four kinds of subunits are involved, whereas for the phage enzyme only a single polypeptide is needed. This means that there must be multiple active sites and catalytic domains on the phage polymerase molecule. Although the bacterial RNA polymerases are more complex molecules, for functional studies this is an advantage of sorts since the subunits can be distinguished and the role of each one in the reaction probed separately. For the phage enzyme a more limited number of approaches is possible. Ideally one would like to have the three-dimensional structure of the phage RNA polymerase molecule, together with structures for the enzyme bound to a promoter site, and perhaps with substrates. This goal is certainly feasible in terms of current X-ray crystallographic techniques, but will depend on the development of procedures for isolation of large amounts of a homogeneous and fully active phage RNA polymerase. This has proved to be a difficult goal. An alternative would be to isolate mutant RNA polymerases and study their properties, possibly in conjunction with mutant promoter sites. Unfortunately, this classical approach has not been particularly useful in studying the phage RNA polymerase molecule. Another approach to probing the structural basis of promoter selectively for the phage RNA polymerases was initiated by Beier and Hausmann (41). They took advantage of the fact that viable, intergenic hybrids could be made between T7 and T3, and constructed a series of recombi42. Coleman, J. (1974). BBRC 60,641. 43. Mildvan, A., and Loeb, L. (1979). CRC Crir. Rev. 6, 219. 44. Bautz, E. (1976). In "RNA Polymerase (R. Losick and M. Charnberlin, eds.), p. 273. Cold Spring Harbor Laboratory, Cold Spring Harbor, New York.

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nant phage strains that contained hybrid gene 1 regions, using various pairs of T7 and T3 gene 1 amber mutants. The use of such RNA polymerase variants to investigate the RNA polymerase promoter interaction was made possible by the fact that the two parental gene 1 products differ significantly in promoter specificity (see Section II,B, 1). When the RNA polymerases of progeny from these crosses were tested for their ability to transcribe both phage templates, a range of template specificities was observed (41, 45). Each hybrid enzyme, in addition to having a preference for either T7 or T3 DNA, was found to be capable of transcribing the heterologous template to some extent. Since the map positions of the T7 and T3 umber mutations used in the construction of these hybrids were known, it was possible to predict the positions of the crossover events, and, in turn, correlate the template preference (i.e., promoter specificity) exhibited for these enzymes with the presence of a particular region of the gene 1 sequence. The region in question was identified as being between 0.7 and 0.78 gene 1 length, a distance corresponding to approximately 75 amino acids. However, recent studies on these hybrid gene 1 sequences have indicated a somewhat more complex situation concerning the functional anatomy of the enzyme (46). Using restriction sites to map the recombination events within the gene 1 region, it was discovered that in all 8 cases examined the genetic constitution of the hybrid gene 1 region differs significantly from that predicted based on the positions of the amber mutations in the parental phage. More specifically, this analysis revealed that the active hybrid gene 1 sequences were often the result of complex combinations of genetic rearrangements, including multiple crossovers, and presumably reversions and/or secondary mutations. This suggests that active hybrid T7/T3 gene 1 sequences are rarely formed by single genetic rearrangements, and that promoter selectivity is likely to be a function of more than one region of the polypeptide chain of the enzyme. It now appears that the region from approximately 23 to 5096, together with the carboxyl end of the molecule, are important in promoter recognition (46). A perfect correspondence was found for the level of heterologous activity with the origin of the DNA sequences between 23 and 50%. In conclusion, it no longer seems feasible to define a single small region as coding for a discrete functional domain in RNA polymerase that uniquely specifies template selectivity. At least two domains on the protein, separated from one another on the polypeptide chain, are involved. 45. Hausrnann, R . . and Tomkiewicz, C. (1976). f t r “RNA Polymerase (R.Losick and M. Charnberlin, eds.), p. 731. Cold Spring Harbor Laboratory, Cold Spring Harbor, New York. 46. Ryan, T., and McConnell, D. .I. (1982). J . Virol.. in press.

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It is not yet clear whether these two functional domains form a single active site, which in turn interacts with the promoter. The observed inadvertant selection of extra crossovers, in addition to those originally selected for, might support this idea. Alternatively, it is conceivable that the phage enzyme might behave in a somewhat analogous manner to that found for the E. coli enzyme, where enzyme promoter interaction seems to involve two separate specific interactions between the enzyme complex and the promoter sequence (47, 48). Evidence that there may be two separable DNA sequences involved in the recognition process at the phage RNA polymerase promoter is discussed in the next section.

B. CATALYTIC PROPERTIES 1. Trnnscriptbnal Mups nnd Trinplrrte Specificity

T7 RNA polymerase and related polymerases carry out DNA-directed synthesis of RNA from nucleoside triphosphate substrates. Synthesis of poly(rA) in a reaction dependent on single-stranded DNA and elevated substrate concentrations has also been reported (36). The template specificity of the T7-like phage RNA polymerases is quite striking. Unlike the bacterial RNA polymerases, the phage enzymes normally use only their homologous DNA templates at a substantial rate (8-10, 24). This suggested originally that the phage RNA polymerases were highly specific for particular promoter sites (8, 9, 26) this was confirmed originally by the mapping of these sites on the T7 and T3 genomes by the transcription of nuclease-digested templates (49, SO), of hybrid phage DNAs (SI),and by in )dtw translation (52).The identity, sequence, and position of what are probably all of the promoters for T7 RNA polymerase on the T7 genome has been determined by cloning and sequencing the regions involved (33, 53). When the entire nucleotide sequence of T7 phage is known, it should be possible to write the exact transcriptional map for the entire phage genome. The resulting transcriptional maps of the T7 and T3 genomes are very similar, although the two polymerases are quite different in their transcriptional specificity (9). All of the transcripts initiated by T7 RNA 47. 48. 49. 50. 51. 52. 53.

Siebenlist, U . , Simpson, R . , and Gilbert, W. (1980). Cell 20, 269. Rosenberg, M . , and Court, M. (1979). Anal. Re\,. Genet. 13, 319. Golomb, M . , and Chamberlin, M. (1974). P N A S 71, 760. Golomb, M . , and Chamberlin, M . (1977). J . Virol. 21, 743. Beier, H . , Golomb, M., and Chamberlin, M. (1977). J . Viuol. 21, 753. Niles, E. and Condit, R. (1975). J M B 98, 57. Dunn, J. and Studier, F. (1981). J M B 148, 303.

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polymerase on T7 DNA in ~ > i vand o in vifro are read from the r strand of the DNA ( 8 ) ,from left to right as the standard genetic and physical map is written. The T7 transcription units are arranged in two overlapping clusters, which share common terminator sites at -61 and loo%, respectively, on the T7 map (See footnote (53cr) and Refs. 29, 49, 50, 5 5 ) . There are three classes of T7 polymerase promoters on T7 DNA; class I1 and class I11 promoters govern transcription of genes in two different regulatory classes (33, 49, 56, 57), while the final class is probably involved in replication initiation. Class I11 promoters are used at a much greater rate than class I1 promoters in vitro ( 4 9 , 5 5 , 5 8 ) ;this may be related to the fact that genes in class 111 transcription units continue to be expressed throughout infection, whereas genes in class I1 transcription units cease to be expressed at late times (33, 56, 57, 59). There are five strong T7 class 111 promoters located at map positions 46.5, 55, 57.1, 70, and 87 (29). An additional class 111 promoter giving rise to a very large T7 RNA on polyacrylamide gels (T7 species I RNA) had been positioned near 62% (49-51); however no promoter has been found in this region ( 2 9 ) ,and it is likely that this RNA band is actually composed of a mixture of large RNA species initiated at class I1 and I11 promoter sites that result from readthrough of the 61% terminator (29). Another strong promoter originally designated a class 111 promoter (49) is located at 98.3% (T7 species VI RNA) and may play a role as a replication initiation site (60). The strong class I11 promoters account for over 90% of the in vitro transcripts by T7 polymerase from T7 DNA under normal conditions (SO). Transcripts initiated from the first three sites are terminated at a terminator at about 61% (29, 49, 52), while transcripts from the other two sites end at a terminator near 100% or run off the end of the DNA (29, 49, 55). The class I1 promoters are much weaker in vitro and are located between 14.6 and 44.4% on the T7 genome (29). Thirteen class I1 promoters have been identified; these all give transcripts that read into the T7 polymerase 61% terminator (29). Because these are weak promoters that 53a. The standard T7 physical map is measured from 0 (left end) to iOO% (right end) and contains 40,000 bp ( 5 4 ) . Positions are noted as 5? T7 unless stated otherwise. 54. Studier, F., Rosenberg, A . , Simon, M., and Dunn, J . (1979). J M E 135, 917. 55. Kassavetis, G . , and Charnberlin, M . (1979). J . Virol. 29, 196. 56. Studier, F. W. (1972). Scierice 176, 367. 57. McAllister, W., and Barrett, C. (1977). Virology 82, 275. 58. McAllister, W., and Carter, A . (1980). Ni!clcic Acids Res. 8, 4821. 59. McAllister, W., and Wu, H. (1978). P N A S 75, 804. 60. Studier, F., and Rosenberg, A. (1981). J M B 153, 503.

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give rather large transcripts, and because of the low resolution of early RNA gel systems, these promoters were not originally identified in the products from intact T7 DNA templates (30, 49). However transcription of restriction endonuclease fragments and cloned T7 segments, together with enhanced resolution of transcripts on RNA gels, has allowed detection and positioning of the promoters (29). This was facilitated by the finding that transcription from class I1 promoter sites is enhanced selectively at low Mg2+ concentrations (58). Exact positions have been assigned for most of these promoters by DNA sequencing, taking advantage of the characteristic T7 promoter sequence (53, 61-65). Tne transcriptional map for T3 RNA polymerase on T3 DNA is very similar to that found for T7 (23,50,51). Here again all transcripts are read from the r-strand of the DNA and there are two classes of promoter sites. Five strong class I11 promoters form two sets of overlapping transcription units from near 46% to a terminator near 59%, and from 67% to the end (100%). Eleven weaker class I1 promoters are spread throughout the left half of the genome from 1.5 to 44% and all read to a strong terminator near 59% (66). The sequences of T7 promoter sites show characteristic sequence homologies in the region of the RNA start site. All of the strong class I11 promoters share a common 23-base sequence (53, 61, 621, which begins 17 base-pairs (bp) prior to the nucleotide coding for the 5' terminus of the T7 polymerase transcript (- 17), and continues 6 bases past that site (+6). The class I1 promoters are quite similar (53, 63-66), but have slightly altered sequences in this region of homology (see Table 11). In analogy with the bacterial RNA polymerases it is likely that these sequences contain the DNA residues involved in recognition of the promoter site. The high degree of sequence identity among the phage RNA polymerase promoters is especially striking in view of the great diversity of bacterial RNA polymerase promoter sequences (47, 48). This probably reflects a greater demand for variation of promoter efficiency and interaction with regulatory factors for the bacterial promoter sites.

61. Rosa, M. (1979). Cell 16, 815. 62. Rosa, M. (1981). J M B 147, 199. 63. Oakley, J., and Coleman, J. (1977). PNAS 74, 4266. 64. Panayotatos, N . , and Wells, R. (1979). Narure (London) 280, 35. 65. Boothroyd, J . , and Hayward, R. (1979). Nucleic Acids Res. 7, 1931. 66. Carter, A., and McAllister, W. (1981). J M E 153, 825. 67. Adhya, S., Basu, S.,Sarkar, P., and Maitra, U. (1981). PNAS 78, 147. 68. Kassavetis, G., Butler, E., Roulland-Dussoix, D., and Chamberlin, M. (1982). JBC, in press.

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TABLE I1 DNA SEQUENCES FOR PROMOTER SITESUSEDB Y T7, T3, RNA POLYMERASE"

AND

SP6 P H G E

Nucleotide sequence Phage and promoter - 15

T7 class 111 consensus

Map location

- 10

-5

+I

+5

bp

TA AT ACGAC T CAC T ATAG GGAGA -

T7 class I1 4I.lA 4I.IB 41.3 41.5 61.6 42.5 43.8 b4.3 44.7 T3

SP6 class 111

" T7 class 111 sequence is from Rosa (61. 62). T7 class I1 promoter sequences are from the summary by Dunn and Studier (S3)and from Carter and McAllister (66). For sequences other than the consensus class I11 sequence, bases are shown only if there is a difference between that sequence and the class I11 sequence. The T3 sequence designated 1.2 is that of Adhya c't d.(67) and is for a T3 promoter near the left end of T3 DNA. Map positions and sequences for this and the other T3 promoters shown are from Ref. (23) and from unpublished studies kindly communicated by Dr. W. McAllister. For T3 it has not yet been clearly determined which promoters are class I1 or class 111. The SP6 sequence was determined by E. Butler (unpublished studies) using a Hi/idIII--Bgl I1 fragment spanning the region 39,450 bp to 41,100 bp from the left end of SP6 DNA (12). This fragment contains a strong SP6 promoter (68); the corresponding RNA sequence was determined by M. Gilman (unpublished studies). Map locations are distances from the left end of the DNA molecule, expressed in percentages of the total length. The underlined nucleotide in the first sequence is the transcriptional start site; this is designated + 1 by the standard nomenclature (48).

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The different T7-like RNA polymerases all seem to have evolved their own distinct promoter recognition specificity. T7 RNA polymerase uses T3 polymerase promoter sites on T3 DNA poorly or not at all, (68a) while T3 RNA polymerase uses T7 promoter sites on T7 DNA weakly but specifically (50). This suggested that there might be a partial homology between T7 and T3 specific promoter sites, and sequencing of a T3 promoter site confirmed this notion [Table 11, Ref. (67)l. Although there are differences between the T3 promoter sequence and the 23-base sequence of strong T7 class I11 promoters, there is a region of strong homology stretching from -9 to +4, and the region around - 15 also shows strong homology if it is assumed that the GA at - 11/- 10 on T7 is replaced by the single base, C, in T3 (W. McAllister, personal communication). Comparative studies of a number of T7-like bacteriophages that grow on different bacterial strains show that these phages all induce phagecoded RNA polymerases of MW about 100,000, and show patterns of growth and regulation similar to T7 (fI, f2). However these phages are generally not closely related to T7 or T3 in protein or nucleotide sequences ( I I, 12). The different phage RNA polymerases all show distinct promoter specificities in that they use their homologous phage DNA as template, but do not use heterologous templates such as T3 or T7 DNA (f042).Since these phages may well have evolved from a common ancestor ( 4 , II), this suggests that specific promoter sequences can evolve rapidly, along with other portions of the phage genome, despite the presence of about 20 such sites on the genome, which must change in concert with any alteration in the specificity of the polymerase. A promoter site for the SP6 phage RNA polymerase has been cloned and sequenced (E. Butler, personal communication). The SP6 RNA polymerase does not use T7 or T3 promoter sites and shows no DNA sequence homology by DNA hybridization (12, 25). However the SP6 specific promoter sequence is strikingly similar to the T7 and T3 sequences (Table 11) and bears identical sequences from -3 to -7 bp and at the start site! This suggests that, although SP6 appears unrelated to T7 at the level of gross nucleotide sequences, there may be significant 68a. While the T3 polymerase-specific promoter sites on T3 DNA are not used by T7 polymerase at an appreciable rate, T7 polymerase does use T3 DNA as an effective template ( 9 . 2 4 ) . This is due to the presence of a strong promoter site for T7 polymerase near 84% on the T3 genome (49.50), which has a sequence identical to the T7 class I11 promoter consensus sequence (69). This site is not used at an appreciable rate by T3 RNA polymerase and may be a vestige of the evolution of T3 from a T7-like ancestor. It is possible that T7 polymerase can use the true T3 promoter sites at a low rate, and in fact this is plausible in view of the homology they show with T7 promoter sites. However, the presence of the strong class 111-like T7 promoter site on T3 DNA may mask this low rate of use. 69. Rosa, M., and Andrews, N. (1981). J M B 147, 41.

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homologies at a finer level of analysis. Furthermore, although the T3, T7 and SP6 promoter sites are specific for their particular RNA polymerases, the differences are primarily due to changes in the part of the conserved promoter sequence from -8 to - 17 bp. The result is reminiscent of the situation with promoters for bacterial RNA polymerases, where there are three regions of DNA sequence homology centered about - 10 and -35 bp and at the start site that may play quite different roles in the promoter binding and RNA chain initiation process (47, 48). 2. Rates und Mechanism of Trunscription Cycle Steps As in the case of bacterial RNA polymerases, synthesis of a single RNA chain beginning with free T7 RNA polymerase and a template DNA involves a sequential series of steps-the transcription cycle (32).These are usually designated as template or promoter binding, RNA chain initiation, RNA chain elongation, and RNA chain termination and release. In the case of the bacterial enzymes, each step involves a complex series of reactions (32), and this is likely to be true for the phage polymerases as well (44. 70). However, much less is known about the individual transcription cycle steps carried out by the phage enzymes. This is due primarily to the fact that the phage polymerases do not form highly stable promoter complexes in the absence of substrates, as is found for the bacterial enzyme (21,26,28,35,70-72). In addition, because of the rapid RNA chain elongation rate (see Section II,A,2) the time needed to complete even a very long transcription unit is usually much less than 60 sec. These two properties of the phage RNA polymerases have made it difficult to separate the individual steps of the transcription cycle for detailed study, as has been possible with the bacterial enzyme. Despite these difficulties, some features of the individual steps are known. Specific promoter binding by the T3 or T7 RNA polymerases can be demonstrated in the absence of nucleoside triphosphates by the nitrocellulose filter binding procedure (21, 71). Rather large amounts of the enzyme are needed; a 50- to 100-fold molar excess of RNA polymerase protein gives about 50% retention of labeled phage DNA (21, 71). This is probably due to the fact that the binding constant for T7 RNA polymerase to its promoter sites is very low (-lO'M-'), rather than to a low efficiency of retention of complexes on filters (26), since efficient retention of T7 DNA fragments that bear only a single promoter has been observed (27). 70. Bautz, E. (1973). FEES Lett. 36, 123. 71. Chakraborty, P., Salvo, R., Majumder, H., and Maitra, U. (1977). JBC 252,6485 72. Salvo, R . , Chakraborty, P. R., and Maitra, U. (1973). FP 32, 645.

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The complexes formed with T3 polymerase dissociate quite rapidly 60 sec) when unlabeled DNA is added ( 7 / ) . Since about 20 promoter sites are involved on each DNA, the intrinsic dissociation rate for an individual polymerase-promoter complex can be no more than a few seconds (73).Binding of T7 RNA polymerase at its cognate promoter leads to opening of DNA in the region from - 5 to + 1, as shown by cleavage of the nontranscribed DNA strand (1 strand) with a single-strand-specific endonuclease (22).This gives direct evidence that T7 RNA polymerase, like the bacterial enzyme (47), directly opens base-pairs at the promoter to form an open promoter complex in which the DNA bases on the transcribed strand are available for base-pairing with an incoming substrate (74). Addition of nucleoside triphosphates to T3 polymerase-promoter complexes stabilizes these complexes to dissociation, as measured by acquisition of resistance to the inhibitor heparin (35). Addition of only GTP and ATP gives nearly full protection; this is not unexpected since T3 RNA chains start with the sequence pppGGGA and pppGGGG (75). The kinetics of T7 RNA synthesis with T7 RNA polymerase show a brief lag prior to achieving a maximal rate of incorporation (26).This lag is not abolished at elevated template or substrate concentrations and is probably due to the slow rate of forming stable, initiated transcriptional complexes, since chain elongation by these complexes is very rapid. It is not known what the true rate-limiting step is in this process. If T7 polymerase-promoter complexes are quite unstable it may require many encounters of polymerase with promoter before binding of substrates and chain initiation leads to trapping the polymerase. In support of this notion, addition of a competing template or an inhibitor of chain initiation, at any point during the lag, leads to blocking of transcription from the first template, hence there is no commitment of polymerase or tight binding at the promoter until transcription has begun (28). The rate of RNA chain elongation by both T7 and T3 RNA polymerases is between 200 and 300 nucleotides/sec at 37" (30, 52, 70, 7 1 ) . This is almost 10 times faster than the bacterial RNA polymerase under the same conditions (76). The K , values for T7 and T3 RNA synthesis have been determined by measuring the initial rate of transcription when three substrates are fixed at concentrations about 10 x K , and the fourth nucleotide is varied. Values in the range of 40-100 pLM are obtained for ATP, UTP, and CTP (26, 35, 37); GTP gives anomalous kinetics, probably due to its

-

73. Giacomoni, P. (1976). FEBS Lett. 72, 83. 74. Chamberlin, M. (1974). Annu. Rev. Biochem. 43, 721. 75. Maitra, U . , Jelinek, W., Yudelevich, A., Majumder, H . , and Guha, A . (1980). P N A S 77, 3908. 76. Kassavetis, G., and Chamberlin, M. (1981). JBC 256, 2777.

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role as a chain initiation nucleotide (26, 44, 70). These values probably reflect actual K , values for chain elongation by the phage RNA polymerases, although it cannot be ruled out that the substrates have an effect on some other step of the reaction, since the assay involves extensive recycling (see Section II,A,2). This kinetic treatment assumes a ping-pong type of reaction mechanism (77). It has been shown that chain elongation by bacterial RNA polymerases fits the general equation for such a mechanism, but the process of transcriptional pausing during the elongation reaction can alter the K , values considerably (76, 78). Thus the K , values obtained for E. coli RNA polymerase transcribing T7 DNA are much higher, in the range from 80 to 500 p M (79). Little is known about the chain termination-release phase of transcription by the phage RNA polymerases. The ability of these enzymes to efficiently recycle many times during the reaction (26, 79) testifies that chain termination and enzyme release are highly efficient and reasonably rapid. Utilization of the strong internal termination site for the T7 and T3 polymerases must involve some kind of DNA sequence recognition process similar to that found with bacterial RNA polymerases (48). Reading of this site is not completely efficient and generates a class of readthrough transcripts from both class 11 and class 111 promoter sites (29). It is interesting that at least two T7 genes (genes 11 and 12) appear to depend on this readthrough transcription for expression (29). Although T3 RNA polymerase uses T7 promoters poorly, it reads the T7 polymerase terminator site at 61% quite well (50). Thus for the phage RNA polymerases, as for the bacterial RNA polymerases (80),the recognition of termination signals may be evolutionarily conserved. This may be due to the requirement for a physical structure at the termination site rather than to a specific nucleotide sequence at which binding takes place, since the T7 phage polymerase terminator involves an inverted repeat sequence followed by a series of U residues, just as is found for bacterial RNA polymerases [Rosa and Dunn, cited in Ref. (331. 111.

Other Bacteriophage RNA Polymemses

A.

BACTERIOPHAGE PBS2 RNA POLYMERASE

Not all bacteriophage-coded RNA polymerases fit the mold of the T7 and T3 enzymes. In 1972 it was reported that the growth of the B. subrilis 77. 78. 79. 80.

Rhodes, G., and Chamberlin, M . (1974). JBC 249, 6675. Kingston, R., Nierman, W., and Chamberlin, M. (1981). JBC 256, 2787. Maitra, U . , and Huang, H. (1972). PNAS 69, 55. Wiggs, J., Bush, J., and Chamberlin, M. (1979). Cell 16, 97.

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phage PBS2 was unaffected by prior treatment of the host cells with rifampicin and related compounds (81). While T7 phage growth becomes resistant to treatment of infected cells with rifampicin after about 5 min (82), T7 growth absolutely depends on transcription of early genetic regions by the host RNA polymerase (56). This suggested that PBS2 phage might utilize a rifampicin-resistant RNA polymerase activity even in the initial stages of growth, possibly carried in the phage particle as for the enzymes in some eukaryotic viruses (6, 7). With this observation in mind, Clark, Losick, and Per0 (16) searched in extracts of PBSZinfected B. sirbtilis for an RNA polymerase activity that would transcribe PBS2 DNA in the presence of rifampicin. Such an activity was identified and purified by ammonium sulfate precipitation followed by repeated column chromatography. The final preparation contained five major polypeptide components, of MW 80,000, 76,000, 58,000, 53,000, and 48,000, respectively, as judged by SDS-polyacrylamide electrophoresis (I 7). These were present in roughly equimolar amounts, consistent with a molecule of MW 260,000 as shown by the sedimentation coefficient of 11 S. None of these peptides is present in the B. subrilis host RNA polymerase, and all appear to be synthesized after phage infection. Biochemical studies of the PBS2 RNA polymerase showed that it required the 4 ribonucleoside triphosphates and Mgz+for activity (Mn2+was 20% as active as Mg2+).The template specificity of the enzyme was notable, PBS2 DNA was the most effective template; poly(dAdT) was also active, but other, heterologous phage DNAs were used poorly or not at all. It should be added that PBS2 phage DNA contains uracil in place of thymine. Hybridization competition experiments using RNA transcribed from PBS2 DNA in vitro suggested that the PBS2 RNA polymerase gives transcripts from genetic regions used late in infection. This was consistent with the kinetics of appearance of the enzyme activity and of the enzyme subunits in infected cells; activity is first seen 10-15 min after infection. These results suggest that the PBS2 RNA polymerase purified by Clark is involved in late phage transcription. However, they leave unanswered the question of how early transcription is carried out. It was initially suggested that PBS2 might code for an RNA polymerase activity, carried in the phage particle that could account for early, rifampicin-resistant transcription. However, the PBS2 RNA polymerase components are not detected in the phage particle. PBS2 growth is sensitive to the drug lipiarmycin, which is an inhibitor 81. Price, A . , and Frabotta, M. (1972). BBRC 48, 1578. 82. Summers, W., and Siege], R. (1969). Nmrre (London) 223, 1111.

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of the B . srrhrilis host RNA polymerase (83). Furthermore, in certain lipiarmycin-resistant cells, PBS2 growth becomes sensitive to rifampicin. These results suggest that early PBS2 transcription probably does depend on the host RNA polymerase, but that some early modification may alter its rifampicin resistance characteristics. E. coli mutants have been reported that have such an effect and do not map in the known RNA polymerase subunits.

B . BACTERIOPHAGE N4 RNA POLYMERASE Shortly after the report that PBS2 phage transcription might be independent of the host RNA polymerase, a similar result was obtained with the E. coli bacteriophage N4 (13, 14). While early and middle classes of N4 transcription are resistant to rifampicin, late N4 transcription is sensitive and requires the host cell RNA polymerase. Thus N4 growth is rifampicin sensitive (85). Again, it seemed possible that an RNA polymerase activity carried in the phage particle might be involved. In this instance, that possibility was confirmed by the observation that disrupted N4 particles contain an endogenous RNA polymerase activity (14, 86). This activity was not affected by rifampicin and was dependent on the 4 ribonucleoside triphosphates, and Mg’+, and was highly specific for N4 phage DNA. The enzyme is coded for by an N4 phage gene, which is required for N4 early transcription as shown by isolation and study of temperature sensitive N4 mutants in the polymerase gene (14). Subsequent studies point to the existence of a second, NCspecific RNA polymerase activity induced in infected cells (87),which is responsible for synthesis of N4 middle transcripts. The latter enzyme has recently been isolated and is not the same as the early, virion enzyme, but biochemical studies are still in progress on its actual structure (W. Zehring, L. Rothman-Denes, personal communication). The N4 RNA polymerase from N4 particles was subsequently purified to homogeneity and shown to have extremely unusual properties (15). Preparations contain only a single polypeptide chain of MW 350,000 as measured by SDS-polyacrylamide gel electrophoresis. The sedimentation coefficient of the enzyme (9.5 S), taken with its other hydrodynamic prop83. 84. 85. 86. 87.

Osburne, M., and Sonenshein. A . (1980). J. Virol. 33, 945. Lathe, R., BUC, H . , Lecocq, J-P., and Bautz, E . (1980). PNAS 77, 3548. Zivin, R., Zehring, W., and Rothrnan-Denes, L. (1981).J M B 152, 335. Pesce, A . . Casoli, C., and Schito, G. (1976). Nrrtrrw ( h ~ 7 d ~ 262, J ~ ) 412. Falco, S., and Rothman-Denes, L. (1979). Virology 95, 466.

108

M. CHAMBERLIN AND T. RYAN

erties, confirms that the active enzyme consists of a single subunit. One or two copies are present in each phage particle. The transcriptional properties of the purified enzyme include an absolute dependence on denatured N4 DNA; native N4 DNA is not used (IS). However, the denatured template is transcribed asymmetrically, predominantly from one end of the genome. Further studies of the NCdirected transcription systems should be of considerable interest, especially since there would seem to be a requirement for additional components in the transcription of native N4 DNA.

Euk.aryotic RNA Poljmerases MARTIN K . LEWIS

RICHARD R . BURGESS

I . Introduction . . . . . . . . . . . . . . . . . . . . . . . . . I1. Purification . . . . . . . . . . . . . . . . . . . . . . . . . . A . Sources and Yields . . . . . . . . . . . . . . . . . . . . B . Activity Assays . . . . . . . . . . . . . . . . . . . . . . C . Purification Procedures . . . . . . . . . . . . . . . . . . . 111. Subunit Structures . . . . . . . . . . . . . . . . . . . . . . A . Problems of Determining Subunit Structure . . . . . . . . . . B . Subunit Quantitation . . . . . . . . . . . . . . . . . . . . C . Multiple Forms of the Largest Subunit in RNA Polymerase I1 . D . Subunit Structure of the Active Enzyme . . . . . . . . . . . E . Common Subunits in RNA Polymerases I, 11. and I l l . . . . . IV. Subunit Functions . . . . . . . . . . . . . . . . . . . . . . . A . Renaturation and Reconstitution Studies . . . . . . . . . . . B . The Role of Zinc . . . . . . . . . . . . . . . . . . . . . C . Affinity Labeling and Active Site Studies . . . . . . . . . . . D . Mutant RNA Polymerases . . . . . . . . . . . . . . . . . E . Modification by Phosphorylation . . . . . . . . . . . . . . . F. Activities Associated with Purified RNA Polymerases . . . . . V . Stimulatory Factors . . . . . . . . . . . . . . . . . . . . . . VI . DNA Binding and Catalytic Properties of Purified RNA Polymerases A . Nature of the Template . . . . . . . . . . . . . . . . . . . . B . Binding and Initiation at Nicks . . . . . . . . . . . . . . . C . Initiation on Unnicked DNA . . . . . . . . . . . . . . . . D . Attempts to Demonstrate Sequence-Specific Binding and Initiation E . Other Reactions . . . . . . . . . . . . . . . . . . . . . . VII . Inhibitors . . . . . . . . . . . . . . . . . . . . . . . . . . A . Amatoxins . . . . . . . . . . . . . . . . . . . . . . . . B . Rifamycin Derivatives . . . . . . . . . . . . . . . . . . . VIII . Eukaryotic Transcription Extract Systems . . . . . . . . . . . . A . Discovery of Transcription Extracts . . . . . . . . . . . . .

110 111 111

112 115 117 120 122 123 126 127 128 129 130 132 134 135 136 137 138 138 140 141 143 144 145

145 146 147 147

109 THE ENZYMES. VOL . XV Copyright 0 1982 by Academic Press. Inc . All rights of reproduction in any form reserved ISBN 0-12-122715-4

110

MARTIN K . LEWIS AND RICHARD R. BURGESS

B . Fractionation of Transcription Extracts . C. Do Extracts Mimic in Vivo Transcription? IX. Organelle- and Viral-Coded RNA Polymerases A. Mitochondria1 RNA Polymerase . . . . . B. Chloroplast RNA Polymerase . . . . . . C. Vaccinia Virus RNA Polymerase . . . .

1.

. . . .

. . . .

. . . .

. . . .

. . . .

. . . .

. . . .

. . . .

. . . .

. . . .

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

148 149 150 150 151 152

Introduction

DNA-dependent RNA polymerase (EC 2.7.7.6) transcribes DNA into RNA using the four ribonucleoside triphosphates as substrates. The reaction requires a divalent cation, such as magnesium. RNA is synthesized in the 5’ to 3’ direction and the resulting base sequence is complementary to the DNA template strand. One molecule of pyrophosphate is released for each nucleotide incorporated into the RNA chain. Eukaryotic nuclei contain three distinct RNA polymerases, which differ in function [see Ref. ( I ) ] . RNA polymerase I (or A) synthesizes ribosomal RNA precursors and is concentrated in the nucleolus. RNA polymerase I1 (or B) synthesizes mRNA precursors. RNA polymerase I11 (or C) transcribes 5 S and tRNA genes. In addition to the nuclear enzymes there are separate mitochondria1 and chloroplast RNA polymerases. This chapter concentrates on the nuclear enzymes. The organelle RNA polymerases are discussed briefly in Section IX. A major review of eukaryotic RNA polymerase appeared in 1976 (I). Comprehensive reviews had also been published in each of the three previous years ( 2 4 ) . Specialized reviews focusing on RNA polymerase subunit structure ( 5 ) and plant RNA polymerases (6) have subsequently appeared. By 1976 investigators were aware of the complex subunit structures of the eukaryotic enzymes. RNA polymerases I, 11, and 111 had been resolved chromatographically and distinguished in vitro by their template preferences and characteristic response to salt and divalent cation concentrations. The fungal toxin a-amanitin had been found to inhibit selectively RNA polymerase 11. For enzymes from most sources, the number of 1. Roeder, R. G. (1976).In “RNA Polymerase” (R. Losick and M. Chamberlin, eds.), pp. 285-329. Cold Spring Harbor Laboratory, Cold Spring Harbor, New York. 2. Chambon, P. (1975). Annrr. R e v . Biochem. 44, 613-638. 3. Chambon, P. (1974). “The Enzymes,” Vol. X, pp. 261-331. 4. Jacob, S . T. (1973). Propr. Nucleic Acid Res. Mol. B i d . 13, 93-126. 5. Paule, M. (1981). TIBS 6, 128. 6. Becker, W. M. (1979). I n “Nucleic Acids in Plants” (T. C. Hall and J. W. Davies, eds.), Vol. I, pp. 111-141. CRC Press, Boca Raton, Florida.

5. EUKARYOTIC RNA POLYMERASES

111

subunits had not been clearly defined. A number of areas have now been clarified. Milligram quantities of RNA polymerases I, 11, and 111, previously available only from yeast cells, are now routinely purified from Acunrlzamoeba and plant tissues (Section 11). New purification procedures have resulted in advances in the definition of enzyme subunit structure. Further studies have identified RNA polymerase subunits shared by enzymes I, 11, and I11 (Section 111). The isolation of mutants with altered RNA polymerase subunits promises to contribute to our understanding of subunit structure and function (Section IV). The selectivity of in vitro transcription has been examined. Using defined templates with pure enzyme, selectivity can be demonstrated only under highly artificial conditions, and the in v i m initiation sites are not restricted to those used in vivo (Section VI). It has been found that single-strand nicks in DNA are excellent binding sites for RNA polymerase 11. The 3'-DNA hydroxyl at a single-strand nick may serve in vitro as a primer for the extension of an RNA chain. Because of this, most of the RNA synthesized on nicked templates is covalently linked to DNA (Section VI). Since 1978 much excitement has been generated by the discovery of soluble extract systems that accurately transcribe purified eukaryotic genes (Section VIII). Much effort will now be directed to the fractionation of these systems and to the identification and characterization of the factors required for correct transcription. It may be discovered in the course of such fractionation studies that some of the necessary factors are the same as some of those that have already been found to stimulate the activity of purified RNA polymerase (Section V). II.

Purification

A. SOURCES A N D YIELDS RNA polymerases have been isolated from many eukaryotic tissues although only a few sources have yielded large quantities of pure enzymes. As seen in Table I (see 7-29), yeast, wheat germ, and calf thymus 7. Hager, G. J . , Holland, M . J., and Rutter, W. J. (1977). Biochrmistrv 16, 1. 8. Jendrisak, J. J., and Burgess, R. R . (1975). BiochPrnistry 14, 4639. 9. Hodo, J. G . , and Blatti, S. P. (1977). Biochemistry 16, 2334. 10. Teissere, M . , Penon, P., Azou, Y., and Ricard, J. (1977). FEBS L e f t . 82, 77. 1 1 . Jendrisak, J . J. (1981). P h n r Physiol. 67, 438. 12. Gissinger, F., and Chambon, P. (1972). EJB 28, 277. 13. Spindler, S. R . , Duester, G . L., D'Alessio, J . M . , and Paule, M. R . (1978). JBC 253, 4669. 14. D'Alessio, J. M . , Spindler, S. R . , and Paule, M. R. (1979). JBC 254, 4085.

112

MARTIN K. LEWIS AND RICHARD R. BURGESS

tissues provide the greatest enzyme yields. The enzymes from these sources have been very useful for studies of subunit structure and function. The discovery of new tissue sources has led to success in enzyme isolation and improved methods of solubilization and fractionation.

B. ACTIVITYASSAYS The assay of RNA polymerase activity is used to follow the enzyme over the course of purification. In order to compare relative levels of enzyme activity, assays must be performed in a range where activity is linearly proportional to the amount of enzyme assayed. It is important that the activity requires the presence of all four ribonucleoside triphosphates as well as exogenous DNA template. These characteristics define the activity as DNA-dependent RNA polymerase. RNA synthesis is absolutely dependent on the presence of a divalent cation. Magnesium (Mg’+) and manganese (Mn2+)are the most commonly used. The metal dependence of the reaction is presumably due to the requirement for binding the nucleotide substrates as metal chelates. In some instances the choice of metal cofactor is a practical decision made to obtain the most sensitive assay. Class I1 RNA polymerases are usually more active when assayed with Mn2+than with Mg2+( I ) .However, in vivo levels of Mn’’ are thought to be low, and use of this ion may alter the binding and site selection properties of the enzymes (30). For optimal activity, salts (usually NaCl, KCl, or ammonium sulfate) are included in the assay mixture at levels of 0.01 to 0.1 M. 15. Spindler, S. R., D’Alessio, J . M . , Duester, G. L., and Paule, M. R . (1978). JBC 253, 6242. 16. Witney, F. R., and Surzycki, S. J. (1981). Biochemistry, in press. 17. Greenleaf, A. L., and Bautz, E. K. F. (1975). EJB 60, 169. 18. Smith, S. S . , and Braun, R . (1978). EJB 82, 309. 19. Sridhara, S., and Gilbert, L. I. (1978). EJB 90, 161. 20. Stunnenberg, H. G . , Wennekes, L. M. J., and Van den Broek, J. W. J . (1979). EJB 98, 107. 21. Stunnenberg, H. G. (1981). Ph.D. Thesis, Landbouwhogeschool, Wageningen. 22. Guilfoyle, T. J . (1980). Biochemistry 19, 5966. 23. Goto, J., Sasaki, Y., and Kamikubo, T. (1978). BBA 517, 195. 24. Sasaki, Y., Ishiye, M., Goto, J . , and Karnikubo, T. (1979). BUA 564, 437. 25. Link, G., and Richter, G. (1975). UBA 395, 337. 26. Matsui, T., Onishi, T., and Muramatsu, M. (1976). EJB 71, 351. 27. Krebs, G . , and Chambon, P. (1976). EJB 61, 15. 28. Sugden, B . , and Keller, W. (1973). JBC 248, 3777. 29. Jaehning, J . A., Woods, P. S., and Roeder, R. G. (1977). JBC 252, 8762. 30. Anderson, J . A., Juntz, G. P. P., Evans, H . H., and Swift, T. J. (1971). Biochemistry 10. 4368.

113

5 . EUKARYOTIC RNA POLYMERASES

TABLE I TISSUESOURCES FOR T H E PURIFICATION OF EUKARYOTIC RNA POLYMERASES TYPES1.11, A N D 111” Yieldkg of RNA polymerase (mg) Source

I

Sacchnromyces cerevisiae (yeast) Wheat germ Calf thymus Acanthomoeba castellanii (soil amoeba) Human placenta Drosophila melanogasrer Physarum

poiycephaliim Mandirca sexta (tobacco hornworm) Aspergi1lu.T nidulans Cauliflower influoresence Pea Parsley cell culture Rat liver Hen oviduct KB cellsb

I1 20

I11 20

25 5 2.5

0.75

2 1

-

0.5

-

2

-

1

-

0.6

-

1 0.25

-

0.7 2

Refs,

-

1

0.5

“ This list contains enzymes purified

to near homogeneity. to limited tissue availability enzyme was purified from much less than 1 kg starting material.

* Due

3H- or 32P-labelednucleotides are used to label the product RNA. Only one labeled nucleotide is present in the typical assay mixture, and is generally CTP or UTP. This use derives from studies with E. coli RNA polymerase, which suggest that the concentration of the initiating nucleotide influences the rate of RNA synthesis much more than do the concentrations of the other three (31). Because the first base in an RNA chain is generally a purine, it is advantageous to assay in the presence of 31. Chamberlin, M. J. (1976). In “RNA Polymerase” (R. Losick and M. Chamberlin. eds.), pp. 17-67. Cold Spring Harbor Laboratory, Cold Spring Harbor, New York.

114

MARTIN K. LEWIS AND RICHARD R. BURGESS

high purine nucleotide levels, and deleterious to use purine label due to the resulting dilution of substrate specific activity. All four nucleoside triphosphates are included in an assay mix, even though most commercial substrates are sufficiently cross-contaminated for RNA chains several hundred nucleotides long to form with the nominal addition of only three of the four nucleotides. Polyhydric alcohols, such as glycerol and ethylene glycol (at concentrations up to 50%), are known to stimulate severalfold the in vitro activity of pure ( 3 2 ) and partially pure (33) RNA polymerases. The nature of the effect of these compounds is not understood, but could involve a stabilization of enzyme structure as well as a destabilization of DNA structure. Glycerol has, however, been reported to inhibit RNA polymerase activity (34) in a fashion dependent on the source and age of the glycerol. In some cases metal ions in less pure glycerol cause inhibition that can be prevented by increasing the EDTA concentration. Inhibitory activities also contaminate commercial preparations of calf thymus DNA, a popular template for RNA polymerase assays. RNase is found in this DNA and may be removed by repeated phenol extraction or by heating (35). Contaminating protease is also present. Incubation of purified wheat germ RNA polymerase I1 with commercial native calf thymus DNA results in the cleavage of the large enzyme subunits to smaller molecular weight polypeptides (35a). After the assay has been run for the desired time, labeled RNA is separated from unincorporated substrate by precipitation with trichloroacetic acid (TCA) and filtered onto glass fiber or nitrocellulose filters. Alternatively, the reaction mixtures are spotted onto DEAEcellulose paper disks and unincorporated nucleotides are washed from the paper with 5% sodium phosphate (36). TCA precipitates RNA larger than about 15 nucleotides, whereas DEAE filters retain RNA molecules of trinucleotide size or greater. The DEAE filter assay is the more reproducible of the two assays. TCA precipitation of RNA in assays that contain native DNA converts the nucleic acid to a sticky clump, which can be lost on the walls of the reaction vessel. Also, basic proteins (such as histones) appear to stimulate transcription in the TCA assay, but not the DEAE filter assay, by increasing the precipitation efficiency of the RNA (M. K. Lewis, unpublished). 32. Dynan, W. S., and Burgess, R. R. (1979). Biochemistry 18, 4581. 33. Buss, W. C., and Stalter, K. (1978). Biochemistry 17, 4825. 34. Blair, D. G. R. (1977). Can. J . Biochem. 55, 1117. 35. Dynan, W. S . , Jendrisak, J. J., and Burgess, R. R. (1977). A n d . Biochem. 79, 181. 35a. Jendrisak, J . J. (1978). Unpublished. 36. Lowe, P. A . , Hager, D. A., and Burgess, R. R. (1979). Biochemistry 18, 1344.

5. EUKARYOTIC RNA POLYMERASES

115

However, the TCA precipitation assay has a lower background and allows a larger sample volume to be assayed. To minimize liquid scintillation artifacts, filters containing ‘H-labeled RNA should be solubilized before they are counted (35).This is especially important when varying the DNA concentration or the type of DNA in the assay. It has been shown (35)that the apparent preference of wheat germ RNA polymerase I1 for transcription of heat-denatured template over native template is reversed when assay artifacts are overcome: among these is the greater self-absorption of unsolubilized filters that contain native DNA. Quantitation of the number of initiated RNA chains is achieved by using yJ2P-labeled nucleotides in the reaction mixtures. The above assay methods seriously underestimate the number of chains initiated if the events often produce RNA chains too short to be retained on the filter. The average RNA chain length is determined by calculating the ratio of end to internal label. These data should be interpreted with care. Changes in reaction conditions that decrease the actual average length of some of the RNA below the assay cutoff may decrease the end to internal incorporation ratio, and hence appear to increase the average length of the RNA. A more reliable method of determining RNA chain length is to analyze the RNA by electrophoresis on polyacrylamide or agarose gels with appropriate molecular weight markers (37, 38). C . PURIFICATION PROCEDURES

1. S o h biliztitiori Eukaryotic RNA polymerase activity was originally detected in 1959 in the insoluble cell-free aggregate that sedirnented at low speed with rat liver chromatin (39). Difficulty was experienced in solubilizing a stable activity that was dependent on the addition of exogenous DNA. In 1969 Roeder and Rutter reported the technique of sonication of isolated nuclei or whole tissue homogenates in the presence of 0.3 M ammonium sulfate (40). This released chromatin-bound RNA polymerase activity, and soon found widespread use. A drawback was the length of time required to sonicate large amounts of tissue. It was subsequently found that homogenization in buffers of low ionic strength released much soluble 37. Dynan, W. S . , and Burgess, R. R. (1981). JBC 256, 5866. 38. Wed, P. A . , Luse, D. S . , Segall, J., andpoeder, R. G . (1979). Cell 18, 469. 39. Weiss, S . B. (1976). In “RNA Polymerase” (R. Losick and M . Chamberlin, eds.), pp. 3-13. Cold Spring Harbor Laboratory, Cold Spring Harbor, New York. 40. Roeder, R. G . , and Rutter, W. J . (1969). Nrrrrtre (London) 224, 234.

116

MARTIN K . LEWIS AND RICHARD R. BURGESS

RNA polymerase activity from many tissues (8-10, 13-16, 18, 20, 23, 24, 41, 42). It now appears that simple homogenization of whole tissue in a buffer that contains low concentrations of salt is the most effective way of obtaining soluble RNA polymerase activity from most sources. Isolation of nuclei generally should be avoided as a first step in purification because yield is reduced and only a small purification advantage is achieved.

2. Nucleic Acid Removal Sonication in buffers that contain high concentrations of salt according to the early procedures stripped chromatin proteins from DNA. Lengthy ultracentrifugation was then required to remove the solubilized template. The advantage of the procedure using low salt concentrations, described in the preceding section, is that DNA remains as chromatin and can be mostly removed by low speed centrifugation. Residual nucleic acid can be conveniently removed by precipitation with polyethyleneimine (Polymin P) (8). Following enzyme solubilization and separation from nucleic acid, the activity can be concentrated by ammonium sulfate precipitation. Difficulty experienced in dissolving ammonium sulfate precipitates from crude cell extracts treated with polyethyleneimine can be overcome by keeping the salt concentration of the sample sufficiently high. The use of detergents is not recommended at this point. 3. Column Chromatography The next step in purification is generally chromatography on a DEAEsubstituted resin. DEAE-Sephadex is the only resin that separates all three forms, and the RNA polymerases generally elute sequentially with increasing salt concentrations in the order I, 11, and I11 (40, 43). DEAESepharose CL-6B separates RNA polymerase I from I1 and I11 ( I I ) , whereas DEAE-cellulose separates RNA polymerase I1 from I and I11 (43). The polymerases bind more tightly to DEAE-Sepharose CL-6B than to the other DEAE-substituted resins (18, 43). Chromatography on this resin can therefore be performed in buffers containing higher salt concentrations, alleviating problems of aggregation at low ionic strength. Other commonly used methods for purification are chromatography on phosphocellulose and on affinity resins that contain heparin (10, I / , f8)or immobilized DNA (7, 20, 44). 41. Gupta, K . C . , and Taylor, M. W. (1977). Anal. Biochem. 82, 396. 42. DeLorbe, W. J . (1977). Ph.D. Thesis. University of Iowa, Ames, Iowa. 43. Jendrisak, J. J. (1980). In “Genome Organization and Expression in Plants” (C. J. Leaver, ed.), pp. 77-92. Plenum, New York. 44. Guilfoyle, T. J., and Jendrisak, J. J . (1978). Biochemistry 17, 1860.

5. EUKARYOTIC RNA POLYMERASES

I17

Batch column procedures have sometimes been used to advantage (13-15, 42). Sample aggregation in solutions of low ionic strength is mini-

mized in the presence of the ion exchanger. The kinetics of protein binding to both anion (13. 15) and cation exchange resins (14) have been exploited for polymerase purification by varying either column flow rate or the time of batch adsorption. After elution of RNA polymerase from a column it is usually necessary to adjust the salt concentration of the sample prior to the next step. This is usually accomplished by dilution or by ammonium sulfate precipitation ( 8 ) . Dialysis can often cause large activity losses, but may be the preferred technique for dilute protein solutions that precipitate poorly in ammonium sulfate and should not be further diluted. 4. Piirity

It is important to realize that the measurements of total RNA polymerase activity listed for each step of a purification may not be very reliable because stimulatory or inhibitory contaminants are apt to be present in the earlier stages. Depending on the tissue source, a 5000to 30,000-fold purification is required to achieve a homogeneous RNA polymerase preparation. An enzyme is somewhat arbitrarily defined as pure if its apparent subunit composition is unchanged by further purification steps and the specific activity is constant across an activity peak. Purified eukaryotic RNA polymerases have been shown in many cases to sediment or chromatograph as single activity peaks across which the polypeptide composition is identical (7, 10, 13-15,20, 27). Electrophoresis of purified enzyme samples on polyacrylamide gels under nondenaturing conditions has revealed a single band that is coincident with enzyme activity (43), or several active bands that, as demonstrated by subsequent electrophoresis under denaturing conditions, have related polypeptide compositions (19, 45-47).

111.

Subunit Structures

The subunit structures of eukaryotic RNA polymerases are complex. Electrophoresis on SDS-polyacrylamide gels reveals between nine and fourteen subunits. The polypeptide compositions of RNA polymerases I, 11, and 111 from wheat germ, yeast, and Accrrirhrrmoebrr are shown in Table 11. 45. Kedinger, C . , Gissinger, F., and Charnbon, P. (1974). EJB 44, 421. 46. Link, G.,Kidd, G. H., Richter, G.,and Bogorad, L. (1978). EJB 91, 363. 47. Dezelee, S., Wyers, F., Sentenac, A . , and Fromageot, P. (1976). EJB 65, 543.

118

MARTIN K. LEWIS AND RICHARD R. BURGESS TABLE I1

COMPARATIVE S U B U N ISTRUCTURES T OF EUKARYOTIC RNA P O L Y M E R A S E P ~ Wheat germ

Yeast I

I1

I

111

I1

Acanthnrnoehn

111

I

185

I1

I11

193 (178)

169 160

150

150

152

140 135

138 130

128

133

125 94

82

82 52

55

49 44.5 43

42 40

40

41.5

40

40

37 34.5

35 32

34 30 28.5

30 28

27

27

27

'

27 '

24 23

23

23

24.5 22.5

22.5

22.5

18

16.0

15.5

15.5

15.5

I19

5. EUKARYOTIC RNA POLYMERASES TABLE I1 (Continued) Yeast I 14.5

Wheat germ

I1

I11

14.5

14.5

I

I1

Accinrhnmoebri

I11

I

12.6

I1

I11

12.5 12.0

12.2 11 10 "

10

10

10

10

Subunits in common between enzyme classes are boxed. Parentheses indicate subunit is a degradation product. Subunits that are basic proteins.

As discussed by Paule (3,the subunits may be classified as large (greater than 100,000 daltons) or small (less than 50,000 daltons). A subunit of intermediate size (80,000-90,000 daltons) is found only in RNA polymerase 111. Each enzyme possesses two nonidentical large subunits whose molecular weights are similar but not identical between organisms. The largest polypeptide (200,000-220,000 daltons) is found in RNA polymerase 11. As discussed in Section III,C, this subunit is often present in a degraded form of somewhat smaller molecular weight (indicated by parentheses in Table 11). Within a given organism several polypeptides have identical molecular weights in RNA polymerases I, 11, and 111 while other subunits are unique to each enzyme class. The common subunits are discussed in Section III,E. Sedimentation studies of purified RNA polymerases I, 11, and I11 in sucrose or glycerol gradients have indicated molecular weights of 500,000 to 600,000 for each of the native enzymes (13-15, 23, 26, 45, 46, 48-54). These large molecular weights are consistent with the complex subunit patterns observed, but do not prove that all of the polypeptides are required for enzymatic function. 48. 49. 50. 51. 52. 53. 54.

Buhler, J . M., Sentenac, A . , and Fromageot, P. (1974). JBC 249, 5963. Coupar, B . E. H . , and Chesterton, C. J. (1975). EJB 59, 25. Weaver, R . F. (1976). ABB 172, 470. Valenzuela, P., Weinberg, F., Bell, G . , and Rutter, W. J. (1976). JBC 251, 1464. Van Keulen, H . , Planta, R. J . , and Retel, J. (1975). BBA 395, 179. Sklar, V. E. F., Jaehning, J . A., Gage, L . P., and Roeder, R. G . (1976).JBC 251,3794. Sklar, V. E. F., and Roeder, R . G . (1976). JBC 251, 1064.

120

MARTIN K . LEWIS AND RICHARD R. BURGESS

A. PROBLEMS OF DETERMINING S U B U N ISTRUCTURE T A polypeptide is generally considered a subunit of an enzyme if it copurifies with the activity through multiple chromatographic steps and is present in approximately stoichiometric amounts. While an enzyme may, during the course of purification, be separated from polypeptides that influence its activity, these polypeptides should not be considered to be subunits of the enzyme until functionally relevant binding is demonstrated. Complexity of subunit structure will be underestimated if subunits are lost during isolation or analysis, or if they are not all resolved or detected during gel analysis. The complexity will be overestimated if persistent impurities are present or if some polypeptides are partially degraded or modified. Some examples are discussed below. Purification by ion exchange chromatography can be a harsh step. Ion exchange resins may displace from the enzyme polypeptides that under similar buffer conditions would cosediment with the activity. Nondenaturing gel electrophoresis of enzyme samples may reveal contaminants that have cochromatographed with, but do not bind to, RNA polymerase. Because the conditions of electrophoresis under nondenaturing conditions might dissociate from the enzyme components that are required for activity, it is important to assay (47) the protein bands appearing on the gels. A further problem is that contaminating basic proteins not bound to the enzyme do not migrate into the gel and are not visualized. The subunit composition of most preparations of purified RNA polymerase has not been adequately assessed due to the analysis of insufficiently large amounts of protein, and to the use of a limited range of acrylamide gel concentrations. Later reports show RNA polymerase subunit structures to be more complex than previously reported [compare Refs. (9 and 45; 55 and 50, 56, 57; 58 and .59)]. The importance of acrylamide gel concentration and of applying sufficient amounts of protein to the gels is illustrated in Fig. 1. At acrylamide concentrations below 12%, small molecular weight subunits of wheat germ polymerase I1 comigrate with the tracking dye front and are not resolved. Sufficient protein must be applied to the gel for all the small subunits to be visualized (at least 25 kg on cylindrical gels stained with Coomassie brilliant blue). In preparation for SDS-polyacrylamide gel electrophoresis, it is impor55. 56. 57. 58. 59.

Smith, S. S., and Braun, R. (1981). FEBS Lett. 125, 107. Burgess, A. B . , and Burgess, R. R. (1974). PNAS 71, 1174. Hildebrandt, A., and Sauer, H . W. (1973). FEBS Lett. 35, 41. Jendrisak, J . , and Guilfoyle, T. J. (1978). Biochemistry 17, 1322. Guilfoyle, T. J . , and Key, J. L. (1977). BBRC 74, 308.

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FIG. 1. Polyacrylamide gel electrophoresis of wheat germ RNA polymerase 11. (Adapted from Ref. 6 1 ) . (A) Effect of acrylamide gel concentration on appearance of RNA polymerase subunit structure. Fifteen micrograms of wheat germ RNA polymerase I1 was loaded onto cylindrical gels of polyacrylamide concentrations as indicated. Gels were stained with Coomassie brilliant blue and destained. (B)Effect of amount of protein loaded. Various amounts of wheat germ RNA polymerase I1 (as indicated) were loaded onto 12.5% polyacrylamide gels, stained with Coomassie brilliant blue, and destained. Numbers on the right refer to subunit molecular weights in kilodaltons. [Reprinted with permission from Biochemistry 16, 1959. Copyright (1977) American Chemical Society.]

tant to boil the sample immediately after the addition of sodium dodecyl sulfate (SDS) (60). Proteases (which may contaminate the sample) are inactivated by boiling in the presence of SDS and reducing agent. Many proteases are active in SDS solutions at room temperature. SDS dramatically increases rates of proteolytic degradation of most proteins, presumably because this detergent disrupts tertiary structure, rendering potential cleavage sites more accessible to the protease. Protein bands, which appear as singlets upon electrophoresis in sodium phosphate buffer, may appear as doublets in SDS-polyacrylamide gels 60. Pringle, J. R. (1970). BBRC 39, 46.

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upon electrophoresis in the discontinuous Tris-glycine buffer of Laemmli (61). The latter gel system does not fractionate strictly according to mo-

lecular weight, but may also respond to protein modifications. Related peptides, differing only by covalent modification, may thus appear as doublet bands. The two polypeptides of the doublet at 40,000 daltons in wheat germ RNA polymerase I1 (Fig. 1A) have been shown to be highly related, as revealed by analysis of the polypeptides produced by partial proteolysis (M. K. Lewis, unpublished). These two polypeptides do not separate on SDS gels run in sodium phosphate buffer (61). They may differ solely by modification. It is not ruled out, however, that the two subunits are coded by separate but similar genes. The presence of contaminants common to enzymes from different tissue sources has been considered (62). It has been suggested that actin (about 40,000 daltons) may be a contaminant in some purified RNA polymerase preparations (62). Because eukaryotic organisms are diploid or greater, related nonidentical gene products of homologous alleles may complicate enzyme subunit structure. In wheat, however, the subunit structure of RNA polymerase I1 from hexaploid tissue is identical to that from tetraploid tissue (58). B. S U B U N IQUANTITATION T Polypeptide molar ratios are determined by polyacrylamide gel analysis from measurements of molecular weight and protein mass, as inferred by dye binding. Both measurements are subject to significant errors, a fact that demands a careful interpretation of the presence of subunits in apparent nonstoichiometric amounts. For quantitation of relative subunit mass it is important to carefully equilibrate the gels with protein stain prior to densitometric analysis (45, 6 1 ) . The problem of differential dye binding should also be considered (45, 61). Molecular weight determinations are not absolute and are especially inaccurate for small proteins (63). The presence of subunits in nonintegral molar ratios could be due to the alteration of some fraction of a subunit population by covalent modification. In this case, the sum of the ratios of the modified forms should be integral. The two related 40,000 dalton polypeptides in wheat germ RNA polymerase I1 have molar ratios of 0.4 and 0.6 (61). 61. Jendrisak, J. J . , and Burgess, R. R. (1977). Biochernistrf 16, 1959. 62. Smith, S. S., Kelly, K. H . , and Jockusch, B. M. (1979). EBRC 86, 161. 63. Maurer, H . R. (1971). “Disc Electrophoresis and Related Techniques of Polyacrylamide Gel Electrophoresis,” pp. 1-17. Walter de Gruyter, New York.

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C. MULTIPLE FORMSOF POLYMERASE I1

THE

123

LARGEST S U B U N I ITN RNA

Electrophoresis of calf thymus or rat liver RNA polymerase I1 under native conditions revealed several bands (45). Electrophoresis of each of the bands in the presence of SDS revealed that the bands contained identical polypeptides except for the largest polypeptide, which varied in size from 180,000 to 220,000 daltons (45). Neither deliberate aging of the crude extract nor use of the serine protease in hibitor phenylmethylsulfonyl fluoride (PMSF) had any effect on the relative amounts of the different enzyme forms. Thus it was felt that the heterogeneity was not due to limited proteolysis during enzyme isolation (45). Conversely, in yeast the use of PMSF and the careful control of extract pH to minimize activation of proteases were judged necessary for the detection of an undegraded species of the largest subunit of RNA polymerase XI (47). These precautions allowed the isolation of an enzyme that contained a 220,000 dalton polypeptide, whereas previously only a 180,000 dalton band had been observed. In nondenaturing gels, the new enzyme preparation showed two protein bands, each of which was enzymatically active. Resolution of the subunit structures of these bands on SDS-polyacrylamide gels revealed that each contained a 150,000 dalton subunit and the usual spectrum of smaller polypeptides. However, the molecular weight of the largest subunit was 220,000, in one band, and 180,000, in the other. The occurrence of proteolysis during enzyme isolation has been a matter of some concern ( 9 , 14, 18, 19, 22, 25, 27,47,55,59,64-68). As pointed out by Dezelee et ul. (47), proteases present in crude extracts may be complexed with inhibitors that are not removed until later stages of the purification. Thus, incubation of crude extracts may not increase in vitro proteolysis. Furthermore, it should be recalled that of the five mechanistic classes of proteases (69), only one is sensitive to PMSF. Evidence for sequence homology among the various large subunit forms was provided by tryptic peptide mapping of these proteins from cultured parsley cells (46). RNA polymerase I1 from this source contains polypeptides of 220,000, 200,000, 180,000, and 140,000 daltons. This pattern of 64. Coupar, B. E. H., and Chesterton, C. J . (1977). FEES Lett. 77, 273. 65. Pflugfelder, C . , and Sonnenbichler, J. (1978). FEES L e f t . 93, 361. 66. Greenleaf, A . L., Haars, R., and Bautz, E. K . F. (1976). FEES L e f t . 71, 205. 67. Osuna, C . , Renart, J . , and Sebastian, J . (1977). EBRC 78, 1390. 68. Guilfoyle, T. J., and Malcolm, S. (1980). Develop. B i d . 78, 113. 69. Walsh, K. E. (1975). In “Proteases and Biological Control’’ (E. Reich, D. B . Rifkin, and E. Shaw, eds.), pp. 1-11. Cold Spring Harbor Laboratory, Cold Spring Harbor, New

York .

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MARTIN K. LEWIS AND RICHARD R. BURGESS

subunits is strikingly similar to that reported for purified rat liver RNA polymerase I1 (45). The molar ratio of the 140,000 dalton subunit is approximately equal to the sum of the molar ratios of the three higher molecular weight peptides. This relationship had previously been observed with the rat liver enzyme (45). Electrophoresis of the cultured cell polymerase under nondenaturing conditions separated enzyme forms that each contained the 140,000 dalton subunit in association with a different higher molecular weight subunit. The tryptic maps of the 220,000, 200,000, and 180,000 dalton subunits appeared identical, whereas the map of the 140,000 dalton subunit was entirely distinct. Recent evidence indicates that the molecular weight of the largest subunit of RNA polymerase I1 purified from germinating plant tissue differs from that of the enzyme purified from quiescent, embryonic tissue (43, 70). RNA polymerase I1 from soybean embryo has, for example, a 215,000 dalton subunit, whereas enzyme purified from the metabolically active soybean hypocotyl tissue contains a 180,000 dalton polypeptide. It is possible that the 180,000 dalton protein is a degradation product of the 215,000 dalton form since germinating tissue contains protease capable (in vituo) of such degradation, but embryonic tissue does not. Peptide mapping by partial proteolysis of the two forms of soybean large subunit shows nearly identical patterns (70), whereas no homology was detected between these peptides and the 138,000 dalton soybean subunit. Enzymes from embryonic tissue are designated form IIA; those from germinated tissue form IIB. Both forms are active and there is no evidence for marked differences in catalytic properties. RNA polymerase I1 from wheat germ is a IIA enzyme with large molecular weight subunits of 220,000 and 140,000 daltons (61). Purified enzyme preparations may, however, also contain minor bands that migrate to positions between that of the 140,000 and 220,000 dalton subunits. Electrophoresis of wheat germ RNA polymerase IIA on a 5% polyacrylamide-SDS gel is shown in Fig. 2A. Two minor bands (labeled D1 and D2) are seen to migrate to positions corresponding to molecular weights of 210,000 and 180,000, respectively. Except for the small amount of these polypeptides, relative to the 220,000 dalton subunit, the pattern closely resembles that of the parsley (46), rat liver ( 4 9 , and murine plasmacytoma (71) enzymes. Each of the four bands was excised from the gel and analyzed by partial proteolysis with a-chymotrypsin (72). As shown in Fig. 2 the pattern of partial degradation products generated from the 70. Guilfoyle, T. J., and Jendrisak, J. J. (1978). Biochemistry 17, 1860. 71. Sklar, V. E. F., Schwartz, L. B . , and Roeder, R. G. (1975). PNAS 72, 348. 72. Cleveland, D. W., Fischer, S. G., Kirschner, M. W., and Laemmli, U. K. (1977). JBC 252, 1102.

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EUKARYOTIC RNA POLYMERASES

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FIG.2. Relatedness of the large subunits of wheat germ RNA polymerase 11. (A) Large subunits of wheat germ RNA polymerase 11. Five micrograms wheat germ RNA polymerase I1 was 3H-labeled by reductive methylation and electrophoresed on a 5% polyacrylamideSDS gel. The gel was stained with Coomassie brilliant blue and destained. Numbers refer to the molecular weights of the large subunits. D1 and D2 indicate minor bands migrating between the large subunits. (B) Partial proteolysis of the large enzyme subunits. Each of the top four of the designated bands from (A) was excised from the gel, subjected to partial proteolysis with a-chymotrypsin, and analyzed by electrophoresis on 10% polyacrylamideSDS gels. Shown is a fluorogram of the resulting partial degradation patterns exposed for either 8 days (bands 220 and 140) or 32 days (bands D1 and D2). Numbers on the right indicate molecular weights in kilodaltons.

220,000 dalton subunit is identical to those deriving from the D1 and D2 polypeptides. The cleavage pattern of the 140,000 dalton subunit is clearly different. D1 and D2 may be generated by limited proteolysis of the 220,000 dalton subunit in vivo or during enzyme purification. The similarity of the patterns of the small molecular weight subunits from IIA and IIB

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enzymes suggests that the small peptides of enzyme I1 do not derive from proteolytic cleavage of the largest subunit. Multiple forms of the largest subunit have been observed only with the class I1 enzymes. T OF T H E ACTIVEENZYME D. S U B U N ISTRUCTURE Polypeptides present in enzyme preparations in mole ratios less than unity may be contaminants. On the other hand, they may be subunits that modulate, or are rate-limiting for enzyme activity. It is obvious that a polypeptide present overall in trace amounts is not contained in every enzyme molecule. It is possible that only molecules possessing that polypeptide are enzymatically active. Since it is not known what fraction of the molecules in an RNA polymerase preparation are active, it is possible that the subunit structure of the active enzyme might be somewhat different than that of the bulk enzyme preparation. This would especially be possible if the fraction of active enzyme were as low as the 5% reported in one case (73). So far no one has determined the subunit structure of the RNA polymerase in a transcribing complex. Some evidence has been obtained that implicates some and not other subunits with enzymatic function or structural subassemblies. Electrophoresis of yeast RNA polymerase I under nondenaturing conditions shows two protein bands (74. 51). Resolution of the structures on SDS-polyacrylamide gels reveals that one form is missing subunits of 34,500 and 49,000 daltons (74, 51). These polypeptides are loosely associated with the enzyme, as judged by their release during phosphocellulose chromatography ( 7 3 , and by the ability to wash them from the polymerase bound to Sephadex columns that contain covalently bound antibody directed against the largest enzyme subunit (74). Removal of these polypeptides by phosphocellulose chromatography does not affect the ability of RNA polymerase I to transcribe poly(dAdT) but does decrease the ability of the enzyme to transcribe native calf thymus DNA. When calf thymus RNA polymerase I was chromatographed on CMSephadex resin (76)the 51,000 dalton subunit was observed to fractionate away from the peak of enzyme activity. This subunit is present in a mole ratio of one in the bulk enzyme (45). However, column fractions depleted 73. Seidman, S . , Surzycki, S. S. , Delorbe, W., and Gussin, G. N . (1979). Biochemistry 18, 3363.

74. Huet, J . , Dezelee, S . , Iborra, F., Buhler, J. M . , Sentenac, A . , and Fromageot, P. (1976). Biochimir 58, 71. 75. Huet, J . , Buhler, J. M., Sentenac, A.. and Fromageot, P. (1975). P N A S 72, 3034. 76. Gissinger, F., and Chambon, P. (1975). FEBS Lett. 58, 53.

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in this subunit were found to be as active on poly(dAdT) and calf thymus DNA as fractions enriched for the peptide. Two polypeptides of 32,000 and 16,500 daltons are dissociated from yeast RNA polymerase I1 upon native gel electrophoresis (47) or chromatography on DEAE-Sephadex A25 in 1.2M urea (77).The enzyme that lacks these two proteins is almost normally active in in vitro assays (77). When wheat germ RNA polymerase I1 is eluted from DEAE-Sephadex A25, with a shallow salt gradient the ratio of subunit mass to enzyme activity across the peak is constant for all subunits except for the subunits of 27,000 and 25,000 daltons (M. K. Lewis, unpublished). The latter is found primarily in the first half of the peak, the former in the second. Individually these peptides do not fractionate with the enzyme activity, but their sum does. It is possible that the two are functionally homologous and required for enzyme activity. Or both may be inert contaminants that influence somewhat the chromatographic behavior of the enzyme. Although the detection of chromatographically distinct activities within a given enzyme class has led to the purification of polymerase forms differing in the composition of small molecular weight polypeptides (54, 78), or in charge (71), fractionation of a crude activity into multiple chromatographic peaks can not be taken as evidence for true heterogeneity. RNA polymerase complexed unequally with a diverse spectrum of contaminants would be expected to exhibit chromatographic heterogeneity due to the interaction of these peptides with the resin. E.

COMMON SUBUNITS I N RNA POLYMERASES I, 11, A N D 111

One of the more interesting developments in the analysis of RNA polymerase subunit structures was the discovery of subunits common to all three enzyme classes. Electrophoresis of calf thymus RNA polymerases on SDS-polyacrylamide gels showed that some subunits of RNA polymerase I had mobilities identical to subunits of RNA polymerase I1 (45). Similar observations were reported for the murine plasmacytoma enzymes I, 11, and 111 (71). Direct evidence for the existence of such common subunits was provided by two-dimensional gel electrophoresis and tryptic peptide mapping of the 27,000, 23,000, and 14,500 dalton polypeptides found both in yeast RNA polymerases I and I1 77. Ruet, A.. Sentenac, A . , Fromageot, P., Winsor, B., and Lacroute, F. (1980)JBC 255, 6450. 78. Matsui, T., Onishi, T., and Muramatsu, M. (1976). EJB 71, 351.

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(79) (see Table 11). Each of these subunits is also present in the yeast class I11 polymerase, as determined by immunological (80) and two-dimensional gel analysis (a]), though in this enzyme the 27,000 and 23,000 dalton poljpeptides are modified with respect to charge (81). In addition, polypeptides of 40,000 and 19,000 daltons, were found in both yeast polymerases I and 111, but not in polymerase I1 (80, 81). For identifying common subunits, tryptic peptide mapping must be considered more reliable than immunological cross-reactivity of certain subunits in one enzyme with antibodies directed against all the subunits of another. It is not certain that the cross-reacting antibody is directed against subunits of common mobility. Recently the antibodies were prepared to each of the purified subunits of yeast RNA polymerase I(A) and II(B) and used to confirm the common yeast subunits boxed in Table I1 (XOtr). These antibodies also showed significant cross reaction with RNA polymerase I1 purified from wheat germ, Artemia salina, Drosophila melanognster, and calf thymus. Subunit structure analysis of the A . castellanii RNA polymerases has also revealed common subunits in the different enzyme classes (82). The subunit patterns of RNA polymerase I, 11, and I11 from this organism are shown in Table I1 and Fig. 3. All three enzymes contain subunits of 22,500, 15,500, and 13,300 daltons, while polymerases I and I11 in addition share subunits of 39,000, 37,000, and 17,500 daltons. A variety of twodimensional gel systems was used to confirm the identity of the common mobility interclass polypeptides. One-dimensional gel analysis of RNA polymerases I, 11, and I11 from wheat germ suggests that the three enzymes share common subunits of 20,000, 17,800, and 17,000 daltons (11). In addition, a subunit of 38,000 daltons appears to be in common between enzymes I and 111, and one of 25,000 daltons in common between enzymes I1 and 111.

IV.

Subunit Functions

The complex subunit structures of eukaryotic RNA polymerases suggest a correspondingly complex regulation of enzyme function. The view has been offered ( 5 ) that the enzymes consist of a “basic functional core” 79. Buhler, J. M., Iborra, F., Sentenac, A., and Fromageot, P. (1976). JBC 251, 1712. 80. Buhler, J. M., Huet, J . , Davies, K. E . , Sentenac, A . , and Fromageot, P. (1980). JBC 255, 9949. 80a. Huet, J., Sentenac, A., and Fromageot, P. (1981). JBC, in press. 81. Valenzuela, P., Bell, G. I . , Weinberg, F., and Rutter, W. J. (1976). BBRC 71, 1319. 82. D’Alessio, J. M . , Perna, P. J . , and Paule, M. R. (1979). JBC 254, 11282.

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FIG.3. Subunit structures ofA. cnstellanii RNA polymerases I, 11, and 111. Protein was

run on 12% polyacrylamide gels in the presence of SDS and stained with Coomassie brilliant blue. Numbers on the left refer to the molecular weights of common subunits (see Table 11). (Reproduced from Ref. 82).

composed of the two large subunits and the three small polypeptides that are common to each enzyme class. So far, there is no reason to consider this core to be an active assembly. Each class of RNA polymerase is assembled from ten to twelve different polypeptides, presumably encoded by an equal number of distinct genes. Theoretically, enzyme regulation could be achieved by modulating the synthesis or availability of a single polypeptide that is required for enzyme activity. The concentration of some set of essential subunits could limit enzyme assembly, much like the a subunit of E. coli RNA polymerase seems to govern the assembly of p and p’, which do not associate in its absence (83). A.

RENATURATION A N D

RECONSTITUTION STUDIES

If the subunits of RNA polymerase could be separated and then remixed to allow reconstitution of enzyme activity, each subunit could be 83. Ishihama, A . , and Ito, K. (1972). J M B 72, 111 .

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tested to determine if it were a necessary part of the enzyme. Unfortunately, no successful reconstitution of activity has yet been reported with eukaryotic RNA polymerases. Escherichiu roli RNA polymerase whose subunits have been completely unfolded in 6 M guanidine hydrochloride (GuHC1) under reducing conditions can be renatured by simple dialysis into buffer that contains no denaturant (84). Attempts to renature wheat germ RNA polymerase I1 exposed to such conditions have not been successful (M. K. Lewis, unpublished). Exposure of the wheat germ enzyme to GuHCl concentrations of 1 M (or above) results in the apparently irreversible loss of catalytic activity. Under these conditions the half-life of enzyme activity is about one minute (M. K. Lewis, unpublished). Gel filtration of wheat germ RNA polymerase I1 on Sephadex GlOO in 1 M GuHCl generated two protein peaks. The voided peak contained the two large subunits (220,000 and 140,000 daltons) as well as the 16,500 and 14,000 dalton'enzymeassociated polypeptides (see Fig. 1 and Table 11).The other nine subunits were eluted in the included volume (M. K. Lewis, unpublished). The inactive void peak could not be reactivated by dialyzing away the denaturant, either in the presence or absence of the included volume polypeptides. In analogous experiments, wheat germ RNA polymerase I1 was exposed to LiCl. Concentrations of LiCl of 4 M or greater caused irreversible loss of enzyme activity. Gel filtration in 4 M LiCl revealed that only the 27,000 and 25,000 dalton subunits, both basic in charge, eluted in the included volume, and thus were the only two subunits released from the enzyme under these conditions (M. K. Lewis, unpublished). At denaturant concentrations insufficient to destroy catalytic activity, no dissociation of subunits was observed. It is not clear why activity is irreversibly lost under conditions where most subunits remain aggregated. It is possible that the conditions facilitate removal of zinc from the enzyme and cause irreversible structural alterations in essential metal-containing polypeptides.

B. THEROLEOF ZINC Tightly bound zinc is found associated with highly purified eukaryotic RNA polymerases. The reported stoichiometries are not consistent with a simple picture of the functional role of the bound metal. Thus RNA polymerase I1 from yeast was reported to contain 1 g-atom of ZnZ+ (85), polymerase I1 from wheat germ 7 g-atoms Zn2+(86), and polymerase 84. Harding, J . D . , and Beychok, S. (1974). P N A S 71, 3395. 85. Lattke, H . , and Weser, U. (1976). FEBS Left. 65, 288. 86. Petranyi, P., Jendrisak, J. J., and Burgess, R. R. (1977). BBRC 74, 1031.

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I1 from Ei&wrr grrrcilis 2.2 moles bound Zn2+per mole of enzyme (87). RNA polymerases I and 111 from yeast have also been reported to contain tightly bound zinc in respective stoichiometries of 2.4 and 4.3 g-atoms per enzyme mole of (88, 89). These metal determinations have rested on the use of atomic absorption spectroscopy, neutron activation analysis, and microwave-induced emission spectroscopy. The different zinc stoichiometries reported for RNA polymerase I1 from various organisms may simply reflect the analysis of impure enzyme preparations in some cases. Furthermore, some fraction of the enzyme may have released zinc during the course of purification; the procedures for purification are different for each enzyme source. The bound metal may be required for enzyme activity. However, there is no indication of the fraction of polymerase molecules that is active in the final purified preparations. Due to both the intrinsic differences in enzyme catalytic properties and the use of dissimilar assay conditions, relative estimates of this fraction for enzymes from different sources can not be obtained from the reported enzyme specific activities. A correlation exists between the presence of bound zinc and enzyme inhibition by the metal chelator 1,lO-phenanthroline. Inhibition of RNA polymerase activity by this compound, but not by its nonchelating isomers, has been considered diagnostic of the presence of essential bound metal (85,87, 90,91). This suggests that RNA polymerase I from rat liver (91), sea urchin (90, 91), andE. gracilis (87); RNA polymerase I1 from rat liver (88); and RNA polymerase I1 and 111 from sea urchin (91) are also zinc-containing metalloproteins. The interpretation of such inhibition studies should be approached with caution, however. While enzyme activity is more sensitive to the chelating phenanthroline inhibitors than to the nonchelating analogs, the latter compounds can also inhibit when present at somewhat higher, but still relatively low, concentrations (85. 91). These observations suggest that inhibition can occur for reasons other than chelation of enzyme-bound metal. It has been proposed that Zn2+functions at the catalytic site by promoting the acidity of the RNA 3’-hydroxyl, thus facilitating nucleophilic attack of the phosphate ester linkage in the nucleotide substrate (92, 93). 87. 4468. 88. 89. 90. 91. BBRC 92.

Falchuk, K . H . , Mazus, B., Ulpino, L., and Vallee, B. L. (1976). Biochemistry 15, Auld, D. S . , Atsuya, I., Campino, C., and Valenzuela, P. (1976). BBRC 69, 548. Wandzilak, T. M., and Benson, R. W. (1977). BBRC 76, 247. Slater, J. P., Mildvan, A. S., and Loeb, L. A. (1971). BBRC 44, 37. Valenzuela, P., Morris, R. W.,Faras, A., Levinson, W., and Rutter, W. J. (1973). 53, 1036. Slater, J. P., Tarnire, A., Loeb, L. A., and Mildvan, A . S . (1972). JBC 247, 6784.

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The finding that Znz+ is associated with many nucleotidyltransferases (9/), including reverse transcriptase (94) and the RNA polymerases from E. cofi (95) and B. suhrilis (96), suggest mechanistic similarities among these enzymes. In addition to serving a catalytic role, enzyme-bound zinc may also contribute to the stabilization of tertiary structure. The inhibition of the activity of wheat germ RNA polymerase 11that accompanies the removal of Zn2+by 1,lO-phenanthroline is not reversed in the absence of the inhibitor, even when exogenous zinc is supplied (86). This result is somewhat at variance with the report (87) that E. gracilis RNA polymerase I1 recovers full activity upon dilution from 1,lO-phenanthroline. It is possible, however, that real differences exist between the two enzymes with respect to the function as well as the stoichiometry of enzyme-bound zinc. In this context, it is interesting to note not only that the reported Zn2+complement is greater in the wheat germ enzyme, but that the inhibition by 1,10-phenanthroline is a much slower process for this polymerase than for the Euglena enzyme. The greater zinc content of the wheat germ RNA polymerase I1 may contribute to the greater relative stability of its catalytic activity. Irreversible structural changes accompanying the removal of zinc from the wheat germ enzyme may account for the failure to renature enzyme activity from the unfolded state. The reversibility of 1,lOphenanthroline inhibition of the Euglena polymerase makes this enzyme an attractive candidate for renaturation-reconstitution studies. It is not known which subunits of the eukaryotic RNA polymerases contain bound zinc. It is likely, however, that some such subunits will also contain a substrate binding site. In this connection it has been shown that ZnZ+is bound exclusively to the p’ subunit of Bacillus subtilis RNA polymerase (96). This subunit is retained on columns of Blue-Dextran Sepharose, an affinity resin specific for nucleotide binding domains (96, 97).

c.

AFFINITY LABELING AND ACTIVE SITE STUDIES Attempts to localize functional domains of RNA polymerase to particular enzyme subunits have focused on determining the sites of interaction 93. Springgate, C. F., Mildvan, A. S. , Abramson, R., Engle, J. L . , and Loeb, L. A. (1973). JBC 248, 5987. 94. Auld, D. S., Kawaguchi, H., Livingston, D. M., and Vallee, B. L. (1975). BERC 62, 296. 95. Scrutton, M . C . , Wu, C. W., and Goldthwait, D. A. (1971). PNAS 68, 2497. %. Halling, S. M . , Sanchez-Anzaldo, F. J., Fukuda, R., Doi, R. H., and Meares, C. F. (1977). Biochemistry 16, 2880. 97. Thompson, S. T., Cass, K. H., and Stellwagen, E. (1975). PNAS 72, 669.

5. EUKARYOTIC RNA POLYMERASES

I33

of various inhibitors of enzyme activity. Amanin is a cyclic peptide belonging to the amatoxin family (of which a-amanitin is a member) and is a potent inhibitor of RNA polymerase I1 activity (see Section VII). L3H1Amanin has been covalently coupled to calf thymus RNA polymerase I1 by reaction of the amanin-enzyme complex with a water soluble carbodiimide (98). It was suggested that the reagent couples the aspartyl carboxyl group of amanin to a lysyl amino group on the enzyme (98). Resolution of the enzyme subunit pattern on polyacrylamide gels containing sodium dodecyl sulfate showed that the radioactivity migrated at the position of the second largest polymerase subunit (140,000 daltons). This suggests that the amanin binding site is located on this subunit, but it is also possible that the inhibitor binds to an adjacent subunit in a manner such that cross-linking occurs to the neighboring 140,000 dalton polypept ide . Yeast RNA polymerase I is inhibited by low concentrations of pyridoxal 5’-phosphate (PLP) (99). The aldehyde function of this compound can form a Schiff base with the €-amino group of enzyme lysyl residues. Inhibition by PLP is reversed by dilution or by addition of amines, such as Tris, ethylenediamine, or free lysine (99). Although tests of the possible stimulatory effect of these compounds on the uninhibited enzyme were not reported, the results are consistent with a reversible dissociation of the enzyme-inhibitor complex. This view is strengthened by the inability to reverse inhibition after reduction by NaBH, (99); NaBH4inactivates free PLP by reducing the aldehyde function, and fixes bound PLP by reducing the enzyme-inhibitor S c h 8 base complex. An estimate of the number of bound PLP molecules was obtained by reduction in the presence of [3H]NaBH4. The number of hits required to inactivate the enzyme was then determined by comparing the amount of bound inhibitor to the residual enzyme activity. This number was found to be in the range of three to four (99). Assuming that the inhibitor reacts at random, this result could be interpreted to mean that one in four of the lysines available for reaction is a catalytically essential residue. There is no a priori reason to believe that PLP binds selectively to a particular functional domain. Lysines are expected to be involved in DNA, RNA, and substrate binding. Since the unionized form of the lysine amino group is the reactive species, and the catalytically functional form is likely cationic, it is not expected that essential lysines will be more 98. Brodner, 0. G . , and Wieland, T. (1976). B i n c / ~ e m i s t rIS, ~ 3480. 99. Martial, J . , Zaldivar, J . , Bull, P., Venegas, A., and Valenzuela, P. (1975). Biochemistry 14, 4907.

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reactive. Studies ofE. coli RNA polymerase have implicated ten to twenty cationic residues in DNA binding alone (fUU-fU2). Display of the labeled subunit pattern following reaction of yeast polymerase I with PLP and reduction with [3H]NaBH4does not show labeling of all subunits in proportion to their mass (f0.3). The observed distribution may simply reflect the dissimilar lysine contents of the different enzyme subunits. Furthermore, it is not clear that selective labeling is not achieved through differential rates of reduction enforced by steric and microenvironmental constraints. However it is surprising that an extremely basic subunit of 28,000 daltons (79) apparently labels very little, if at all (103). Reduction with [3H]NaBH4in the presence of PLP and nucleoside triphosphates selectively inhibits tritium incorporation into the largest enzyme subunit (185,000 daltons) (103). This is suggestive evidence for the presence of a nucleotide binding site on this subunit. Earlier studies (99) examined the protection provided by nucleoside triphosphates against inhibition by PLP, and noted that a single nucleotide gave less protection than a mixture of three when present at the same total concentration. This suggests that the different nucleotide substrates may bind the polymerase at nonidentical sites. Modification studies of yeast RNA polymerase I have used dinitrothiobenzoate (DTNB) and p-chloromercuribenzoate (pCMB) to quantitate reactive sulfhydryl groups in both the native and urea-denatured enzyme (104). Forty-five sulfhydryl residues reacted in the unfolded enzyme, whereas only twenty were available in the native state. Both reagents inactivated the polymerase. This inhibition could be reversed in the presence of reducing agents. The binding of two moles of jp-mercuribenzoate per mole of enzyme was sufficient to completely inhibit enzyme activity. It is not known if the modifying reagents react selectively with particular cysteine residues, or bind more or less at random to a sulfhydry1 reaction surface in which about one-half of the residues are essential for catalysis. D. MUTANT RNA POLYMERASES The characterization of mutant RNA polymerases offers both an assurance of functional relevance in vfvo and a means to discover the catalytic 100. deHaseth, P. L., Lohman, T. M., Burgess, R. R., and Record, M. T.,Jr. (1978). Biochemistry 17, 1612. 101. Lohman, T. M., Wensiey, C. G . , Cina, J., Burgess, R. R . , and Record, M. T., Jr. (1980). Biuchemisfry 19, 3516. 102. Siebenlist, U., and Gilbert, W. (1980). P N A S 77, 122. 103. Valenzuela, P., Bull, P., Zaldivar, J., Venegas, A., and Martial, J . (1978). BBRC 81, 662. 104. Bull, P., Wyneken, U., and Valenzuela, P. (1981). Riorhemisrry, in press.

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role of individual enzyme subunits. Temperature-sensitive mutants are particularly useful in this regard. Though mutant cells in culture, including Chinese hamster ovary (CHO), rat myoblast (105-107), and mutant Drosophila (108, 109) have been shown to contain a-amanitin resistant RNA polymerase 11, biochemical characterization of the enzymes has not yet assigned the mutations to particular subunit genes. If the amanitin binding domain is formed by more than one subunit, or if the structural integrity of this domain depends on subunit-subunit interactions, it is likely that mutations in various subunit genes will confer an amanitin-resistant phenotype. A temperature-sensitive phenotype does not strongly correlate with a-amanitin resistance, since of 168 amanitin-resistant CHO cell clones only 9 were found to be temperature-sensitive for growth (107). Crude extracts of yeast mutants temperature-sensitive for RNA synthesis were screened using an assay that specifically detects RNA polymerase I1 (110, I f I). A strain exhibiting depressedin vitro activity was found to contain an alteration in the largest subunit of RNA polymerase 11 (77). Partial proteolysis in the presence of SDS produced degradation patterns of the 220,000 dalton polypeptide; the patterns were distinct in the mutant and wild-type enzymes. Significantly, the patterns of the 180,000 dalton polypeptide showed the same alteration, providing strong evidence that the two species of largest subunit are encoded by the same gene.

E. MODIFICATION BY PHOSPHORYLATION Purification of RNA polymerases from yeast cells grown in the presence of 32P04 identified phosphorylated subunits of 190,000, 43,000, 34,500, 23,000, and 19,000 daltons in RNA polymerase I (112, 113); of 220,000, 44,000, and 23,000 daltons in RNA polymerase I1 (113); and of 24,000 and 20,000 daltons in RNA polymerase 111 (112) [note that the subunit called 105. Crerar, M. M., Andrews, S. J . , David, E. S . , Somers, D . G., Mandel, J . L., and Pearson, M. L. (1977). J M E 112, 317. 106. Gupta, K . C., and Taylor, M. W. (1978). Mutcrrion Res. 49, 95. 107. Ingles, C . J. (1978). P N A S 75, 405. 108. Greenleaf, A. L . , Borsett, L. M., Jiamachello, P. F., and Coulter, D. E. (1979). Cell 18, 613. 109. Greenleaf, A . L., Weeks, J . R . , Voelker, R. A , , Ohnishi, S . , and Dickson, B. (1980). Cell 21, 785. 110. Winsor, B . , Lacroute, F., Ruet, A , , and Sentenac, A . (1979). Molec. Gen. Genet. 173, 145. 1 1 1 . Ruet, A . , Sentenac. A . . and Fromageot, P. (1978). EJE 90, 325. 112. Bell, G . I . , Valenzuela, P., and Rutter, W. J. (1977). JBC 252. 3082. 113. Buhler, J . M . , Iborra, F., Sentenac, A . , and Fromageot, P. (1976). FEES Lett. 71, 37.

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MARTIN K . LEWIS AND RICHARD R. BURGESS

24,000 daltons in Ref. (112) is the same as that called 23,000 daltons in Ref. (113)]. Phosphoserine and phosphothreonine were identified in RNA polymerase I ( 1 12). The 23,000 dalton polypeptide, a subunit shared by enzymes I, 11, and 111, is strongly labeled. The degraded form of the large subunit (180,OoO daltons) in RNA polymerase I1 was not phosphorylated ( 1 13). In addition, no 32Plabel was found associated with the 44,000 dalton polypeptide in enzyme that contained a degraded large subunit. There is disagreement ( 1 12) as to whether the undegraded large subunit (220,000 daltons) is phosphorylated, or whether labeling is due to a contaminant migrating in close proximity to this polypeptide. It has been reported that the 220,000 dalton subunit of wheat RNA polymerase I1 is phosphorylated at early times in germination (114). RNA polymerases I and I1 have been phosphorylated in vitro by protein kinases isolated from the homologous tissue source (112, 115-118). A yeast protein kinase which initially copurified with RNA polymerase I phosphorylates the 49,000 dalton subunit of polymerase I and the 34,500 dalton subunit of polymerase IT, in addition to the subunits that are also phosphorylated in vivo ( 1 12). Protein kinases have been reported to stimulate the activity of purified RNA polymerases I and I1 (115-118). Since RNA polymerase phosphorylated in vitvo by a protein kinase has not been purified away from the kinase before polymerase activity was determined, it is not clear whether the stimulation observed is due directly to phosphorylation of the polymerase or to some other effect.

F. ACTIVITIES ASSOCIATED WITH PURIFIED RNA POLYMERASES Polypeptides of 42,000 and 24,600 daltons are associated with RNA polymerase I from Morris hepatoma ( 1 17). Immunological and other studies suggest that these polypeptides are the same as those that compose a homologous nuclear protein kinase (I 17, I 19). Enzyme depleted of these polypeptides was poorly active, but could be stimulated by addition of the purified kinase (117, 119). However, polypeptides of 44,000 and 26,000 daltons composing calf thymus casein kinase I1 were shown by tryptic fingerprinting to be unrelated to the 44,000 and 25,000 dalton subunits of calf thymus RNA polymerase I ( 1 190). The kinase is a persistent con114. Mazus, B.,Szurmak, B., and Buchowicz, J. (1980).Acra Biochim. Polon. 27, 9. 115. Kranias, E. G., Schweppe, J. S., and Jungmann, R. A. (1977).JBC 252, 6750. 116. Dahmus, M. E. (1976).Biochemistry 15, 1821. 117. Duceman, B. W.,Rose, K. M., and Jacob, S. T. (1981).JBC 256, 10755. 118. Martelo, 0.J. and Hirsch, J. (1974).BBRC 58, 1008. 119. Rose, K. M., Stetler, D. A., and Jacob, S. T. (1981).P N A S 78, 2833. 119a. Dahmus, M.E . (1981).JEC 256, 11239.

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taminant in preparations of calf thymus RNA polymerase I, but may be removed by rechromatography on DEAE-Sephadex A25 ( I 1%). Polypeptides of 49,000 and 40,000 daltons in purified yeast RNA polymerase I have been shown to have RNase H activity (120-122). The 49,000 dalton polypeptide has a peptide map identical to that of a 49,000 dalton chromatin-associated RNase H (120). The 40,000 dalton polypeptide is also found in yeast RNA polymerase I11 (see Table 11).

V.

Stimulatory Factors

There is no shortage of reports concerning stimulatory factors of eukaryotic RNA polymerases (123-130, and reviewed in 1 , 6). As a rule these factors have been reported to be small (20,000-40,000 daltons) basic proteins that appear to stimulate severalfold transcriptional elongation rates of either RNA polymerase I or I1 on native, but not denatured, DNA-especially under conditions of very low ionic strength. In many cases the factors are heat-stable. There is no evidence that these proteins alter transcriptional site selection in vitro, or function by binding RNA polymerase or specific DNA sequences. Such stimulatory factors might act at the levels of binding, initiation, or elongation. They could increase the rate of enzyme cycling or alter the fraction of molecules that is catalytically active. Too often, reports do not include controls for the wide variety of technical artifacts that can lead to apparent increased RNA polymerase activity. Studies with sheared, naked-cell DNA leave unclear the role of the stimulatory factors in transcription of the intact in vivo template that is already complexed with a group of small basic proteins. A review discusses the structure of transcriptionally active chromatin (131). 120. Iborra, F., Huet, J., Breant, B . , Sentenac, A., and Fromageot, P. (1979). JBC 254, 10920. 121. Huet, J . , Buhler, J. M., Sentenac, A., and Fromageot, P. (1977). JBC 252, 8848. 122. Huet, J . , Wyers, F., Buhler, J. M., Sentenac, A., and Fromageot, P. (1976). Nntrrre (London) 261, 431. 123. Nagamine, Y., Natori, S., and Mizuno, D. (1976). FEBS Len. 67, 198. 124. Sekimizu, K . , Kobayashi, N., Mizuno, D., and Natori, S. (1976). Biocheniistry 15, 5064. 125. Link, G. and Richter, G . (1977). EJB 76, 119. 126. Kuroiwa, A., Mizuno, D., and Natori, S. (1977). Biockemistry 16, 5687. 127. Nakanishi, Y . , Sekimizu, K . , Mizuno, D., and Natori, S . (1978).FEBS Lett. 93, 357. 128. Ueno, K . , Sekimizu, K., Mizuno, D., and Natori, S. (1979). Nnrirre (London) 277, 145. 129. Spindler, S. R. (1979). Biockemistrv 18, 4042. 130. Sawadogo, M . , Sentenac, A . , and Fromageot, P. (1980). JEC 255, 12.

138 VI.

MARTIN K . LEWIS A N D RICHARD R. BURGESS

DNA Binding and Catalytic Properties of Purified RNA Polymerases

Eukaryotic RNA polymerases transcribe both denatured singlestranded and native double-stranded DNA templates. The in vitro interactions with native templates are expected to bear more closely on the physiological functioning of the enzymes. An RNA polymerase is expected to sense DNA sequence, selecting from a large population of potential sites only particular ones at which to bind and initiate RNA synthesis. It is not clear that the purified eukaryotic enzymes exhibit any sequence-specific binding of double-stranded DNA. And though the observed preference for initiation in vitro with purines, and in particular with GTP (/32-/34),implies that transcription does not initiate entirely at random, the catalytic action of the pure eukaryotic enzymes resembles in certain ways the action of prokaryotic core polymerase. Since the eukaryotic RNA polymerases can be isolated on columns that contain immobilized DNA, most polymerase molecules in a fresh preparation are capable of template binding. The presence of a small fraction of enzyme molecules capable of specific binding and initiation might be obscured in biochemical assays that measure bulk properties. Nonetheless, it has become apparent that the average catalytically active species responds not to nucleotide sequence-dependent initiation signals, but to structural features of the DNA template, such as single-strand nicks and gaps.

A. NATURE OF

THE

TEMPLATE

DNA from eukaryotic tissue is isolated in a sheared form, with a double-stranded molecular weight of 10-50 x lo6 daltons (52, 135-f37). Each DNA fragment contains about ten nicks or gaps (52, 137). Since the unique sequence complexity of eukaryotic DNA is on the order of loL2 daltons (138), a given gene will be found on only about one fragment in 10". Studies of the binding and in vitro transcription of less complex 131. Mathis, D . J . , Oudet, P., and Chambon, P. (1980). Progr. Nircleic Acid Rrs. M i d . Biol. 24, 1-55. 132. Mandel, J. L . , and Chambon, P. (1974). EJB 41, 379. 133. Lewis, M. K . , and Burgess, R. R. (1980). JBC 255, 4928. 134. Dezelee, S., and Sentenac, A. (1973). EJB 34, 41. 135. Long, E . , and Crippa, M. (1976). FEES Lett. 72, 67. 136. Gissinger, F., Kedinger, C., and Chambon, P. (1974). Bioclzimie 56, 319. 137. Van Keulen, H . , and Retal, J . (1977). EJB 79, 579. 138. Lewin, B. (1974). "Gene Expression," Vol. 2. Wiley, New York.

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templates have used viral DNAs extracted from SV40 (132, 133, 139-143), polyoma (144, / 4 5 ) , adenovirus (73, 146-1423), and recombinant plasmids containing eukaryotic gene sequences (149, 150). SV40, polyoma, and plasmid DNAs are isolated in a supercoiled form. When the supercoils are removed either by the introduction of a singlestrand nick or by the nicking-resealing activity of DNA topoisomerase I, the molecules are referred to as relaxed. The superhelical density of a circular covalently closed template is influenced by the ionic conditions and temperature (151). When studying the effects of supercoiling on transcription it7 vitro, it is important to consider that covalently closed DNA that is relaxed under one set of conditions may become supercoiled under another set of conditions. The digestion of DNA with restriction enzymes has been used to produce templates for in vitro transcription (see Section VIII). These enzymes are frequently contaminated with RNase. For this reason it is advisable to treat restricted DNA with diethylpyrocarbonate prior to its use for in vitro transcription (133). This chemical reacts negligibly with double-stranded DNA. It should also be noted that restriction enzymes have the ability to introduce single-strand nicks at specific secondary recognition sites, and that these nicks may serve as efficient initiation sites in t*itro (133). Native calf thymus DNA was observed to be a much better template for calf thymus RNA polymerases I and I1 than the intact DNAs from either bacteriophage lambda or T4. Denaturation of the phage DNAs increased their template activities, as did treatment with pancreatic DNase I (152). These results suggest that disruption of double-stranded DNA structure creates new sites for RNA synthesis. 139. Mandel, J . L., and Chambon, P. (1974). EJE 41, 367. 140. Hossenlopp, P., Oudet, P., and Chambon, P. (1974). EJB 41, 397. 141. Saragosti, S . . Lescure, B . , and Yaniv, M. (1979). BERC 88, 1077. 142. Saragosti, S., Croissant, O., and Yaniv, M. (1980). EJB 106, 25. 143. Chandler, D . W., and Gralla, J. (1980). Eiorliemisiry 19, 1604. 144. Lescure, B . , Chestier, A., and Yaniv, M. (1978). JME 124, 73. 145. Lescure, B . , Dauguet, C . , and Yaniv, M. (1978). JME 124, 87. 146. Witney, F. R., Surzycki, J. A., Seidman, S . , Dodds. J. R . , Gussin, G. N . , and Surzycki, S. J. (1980). M o / e c . Geri. Gtwet. 179, 627. 147. Seidman, S. L., Witney, F. R . , and Surzycki, S. J. (1980). Molec. Geri. Genet. 179, 647. 148. Hossenlopp, P., Sumegi, J., and Chambon, P. (1978). EJB 90, 615. 149. Lescure, B . , Williamson, V., and Sentenac, A. (1981). N/ir/eic Acids Res. 9, 31. 150. Lescure. B . , Bennetzen, J . , and Sentenac, A . (1981).JEC 256, 11018. 151. Bauer, W., and Vinograd, J . (1968). J M B 33, 141. 152. Gniazdowski, M., Mandel, J . L . , Gissinger, F., Kedinger, C . , and Chambon, P. (1970). EERC 38. 1033.

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MARTIN K. LEWIS AND RICHARD R. BURGESS

The template activity of rat liver DNA toward the homologous RNA polymerases I and I1 was seen to vary according to the protocol followed for DNA isolation (153). The template was activated severalfold by lowlevel DNase treatment, suggesting that single-strand nicks were serving to promote RNA synthesis. The template activity of bacteriophage T7 DNA toward yeast RNA polymerases I and I1 was increased up to threefold by treatment of the template with pancreatic DNase (154). The introduction of short singlestranded gaps into the DNA by successive treatment with DNase and exonuclease I1 provided even greater stimulation (154). Treatment of native calf thymus DNA with S1 nuclease decreased its template activity for yeast RNA polymerases I and 11. The observed inhibition was not necessarily due to the removal of single-stranded regions in the template. The nuclease digestions were performed in low salt (154), and under those conditions S1 nuclease cleaves opposite singlestranded nicks, destroying these sites and creating double-stranded DNA ends (155, 156). It is not clear whether DNA ends are used as initiation sites, are inhibitory to transcription, or have any effect at all. Treatment of native lymphocyte DNA with pancreatic DNase I greatly stimulated transcription of this template by calf thymus RNA polymerase I1 (156). Treatment with micrococcal nuclease however was without effect (156). DNase I generates nicks containing 3’-hydroxyls, whereas micrococcal nuclease generates nicks containing 3’-phosphates (156). Micrococcal nuclease-treated DNA can be activated as a template by removal of the 3‘-phosphates with alkaline phosphatase (156). Addition of 5’phosphates with polynucleotide kinase has no effect on the transcription of micrococcal nuclease-treated DNA. It was concluded that a nick must possess a free 3’-hydroxyl in order to serve as an initiation site for calf thymus RNA polymerase 11. AT NICKS B . BINDINGA N D INITIATION

There is evidence for strong binding interactions at single-strand nicks. A specific nick was introduced into supercoiled SV40 DNA, and the DNA was then treated with a restriction enzyme (143). Wheat germ RNA polymerase I1 selectively retained on nitrocellulose filters the DNA fragment that contained the nick (14.3). Binding of this nick by the enzyme 153. Hint, S. J., Pomerai, D. I., Chesterton, C. J., and Butterworth, P. H. W. (1974). EJE 42, 567.

154. Dezelee, S., Sentenac, A . , and Fromageot, P. (1974). JEC 249, 5971. 155. Germond, J. E., Vogt, V. M., and Hirt, B. (1974). EJE 43, 591. 156. Dreyer, C., and Hausen, P. (1976). EJE 70,63.

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resulted in protection against DNase digestion of a 34-38 base pair double-stranded DNA fragment (143). Resolution of the structure of the protected fragment on denaturing gels showed precise placement of the nick, implying precise positioning of the nick within the enzyme-DNA complex. Limited DNase I digestion of bacteriophage lambda DNA increases its activity as a template for wheat germ RNA polymerase 11. The increased levels of transcription are resistant to the polyanionic inhibitor heparin (32). Heparin inactivates free (but not template-bound) polymerase, suggesting that it is tight binding to nicks that protects the enzyme from inactivation. It is impressive that the introduction of only a few singlestrand nicks per lambda DNA molecule results in a severalfold increase in transcription (32). The ratio of the number of the nicked sites to the number of potential random initiation sites on intact DNA is on the order of 10-4.

It has been shown (133) that much of the RNA synthesized in v i m by wheat germ and calf thymus RNA polymerase I1 is covalently linked to template DNA. This situation results from the use of single-strand nicks as primers for the extension of an RNA chain. Wheat germ RNA polymerase I11 also catalyzes this primed reaction (M. K. Lewis, unpublished). Priming of RNA synthesis by the 3’ DNA hydroxyl at singlestrand nicks was first demonstrated for E. coli core polymerase (157, 158). Two modes of nick-dependent initiation were distinguished for wheat germ RNA polymerase I1 (133). In addition to the priming mode, the enzyme also catalyzes the nick-dependent de novo initiation of RNA synthesis. In this case most RNA chains are begun with GTP (133). Priming is inhibited by MnZ+(133). The use of this ion by earlier workers (156) may explain their failure to detect the reaction. It is not clear whether MnZ+allows more efficient transcription from intact DNA, or whether in its presence nicks are used just as efficiently but in the unprimed mode. The efficiency with which eukaryotic RNA polymerase I1 binds to and transcribes from nicks approaches that of the interactions of E. coli RNA polymerase holoenzyme with double-stranded promoter regions (133).

C. INITIATIONOF UNNICKED DNA The low template activity of relatively intact phage DNAs compared to that of extensively degraded whole-cell DNA preparations raised the issue 157. Wickner, S . , Hurwitz, J . , Nath, K., and Yarbrough, L. (1972). BBRC 48, 619. 158. Nath, K . , and Hunvitz, J. (1974). JBC 244, 2605.

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MARTIN K . LEWIS AND RICHARD R. BURGESS

of whether all transcription was dependent on the presence of structural discontinuities in the template. An analysis of the increase in transcription resulting from the introduction of a known number of nicks into linear SV40 DNA suggested that initial transcriptional levels were consistent with the number of nicked molecules present in the predominantly supercoiled population prior to linearization (133).However, RNA synthesis by wheat germ RNA polymerase I1 on unnicked SV40 DNA was stimulated more at low salt and divalent cation concentrations than was synthesis from a deliberately nicked template (133).It was suggested (133)that this could reflect a differential effect of salt on initiation at nicks versus initiation on intact DNA. Transcription of unnicked bacteriophage lambda DNA by wheat germ RNA polymerase I1 was much more sensitive to inhibition by heparin than was transcription of deliberately nicked DNA (32). This suggested, but did not rigorously show, that unnicked DNA could be transcribed. Subsequently it was rigorously demonstrated that transcription can occur on unnicked relaxed DNA (159). In this study wheat germ RNA polymerase I1 was allowed to bind and initiate transcription on plasmid DNA (containing the entire SV40 genome), which had been relaxed under the same conditions as used in the transcription reaction. Transcription complexes associated with unnicked DNA molecules were fractionated, by agarose gel electrophoresis in the presence of ethidium bromide, away from the complexes initiated on contaminating nicked molecules. Although a few percent of the template that was nicked was transcribed very efficiently (about one-half of the total RNA) it was clear that relaxed unnicked DNA could be utilized as a template by the RNA polymerase. Wheat germ RNA polymerase I1 transcribed supercoiled plasmid DNA one hundred times better than the unnicked relaxed DNA (159). This is presumably because of a negative free energy contribution from the reiaxation of superhelical tension that accompanies strand separation of a negatively supertwisted DNA (160). Facilitated strand separation is expected to enhance RNA polymerase binding and initiation, but may also affect the elongation reaction. Negatively supercoiled DNA had also been reported previously to be a much better template for prokaryotic (161) and eukaryotic (133, 140, 162) RNA polymerases than either linear or unnicked relaxed DNA. 159. Dynan, W. S., and Burgess, R. R. (1981). JBC 256, 5866. 160. Vinograd, J., Lebowitz, J., and Watson, R . (1968). J M B 33, 173. 161. Wang, J . C. (1974). J M B 87, 797. 162. Lilley, D. M . J . , and Houghton, M. (1979). Nucleic Acids Res. 6, 507.

5.

D.

EUKARYOTIC RNA POLYMERASES

I43

TO DEMONSTRATE SEQUENCE-SPECIFIC BINDING ATTEMPTS INITIAT~ON

AND

Studies of the binding of wheat germ RNA polymerase I1 to restriction fragments of lambda, adenovirus, and SV40 DNAs have revealed no sequence specificity, as measured by the technique of filter binding (R. S. Boston and M. K. Lewis, unpublished). The ability of several restriction enzymes to introduce single-strand nicks at specific secondary sites in DNA templates (133) can lead to an artifactual selectivity in studies of this kind. Thus it is important to confirm apparent specificity by using different restriction enzyme digests. Reports of the specific binding of wheat germ (7.3, 146) and human placental RNA polymerase I1 (146, 147) on adenovirus DNA, and of the apparent selective binding of calf thymus and wheat germ RNA polymerase I1 on polyoma (144, 145) and SV40 DNA (142, 144, MS), should be approached with some caution. It is possible that the fixation techniques used in these electron microscopic analyses selectively crosslinked randomly bound enzyme to easily denatured template regions. The use of Mn2+in these studies calls into question the physiological relevance of the results. Mn2+ions serve to further destabilize the double-stranded structure of supercoiled DNA. While Mg'+ stabilizes the double helix by binding to phosphate groups, Mn2+binds the N-7 of guanine and is thus expected to shift the helix coil equilibrium toward the denatured form (140).

Small amounts of specific transcription could be obscured by a large background of random transcription from nicks. The isolation of ternary transcription complexes formed by wheat germ RNA polymerase I1 on unnicked relaxed pBR322ISV40 DNA has been described (159) (see Section V1,C). When the RNA associated with these complexes was purified and run on polyacrylamide gels, many minor bands were seen (159). This suggests that transcription was not entirely at random. Supplementation of transcription reactions with high levels of GTP (0.4 instead of 0.04 mM) gave rise to different RNA bands than did supplementation with high levels of ATP (159). It was concluded, however, that wheat germ RNA polymerase I1 was transcribing from many sites not under the control of the in viva SV40 promoter, since the bands were not altered when mutant DNA that contained a deletion (0.67-0.72 map units) of the SV40 control region was transcribed. Recently an analysis of the transcription of supercoiled DNA has provided a convincing demonstration of in vitm selectivity by purified yeast RNA polymerase I1 (149, 150). Selective transcripts were synthesized by

144

MARTIN K . LEWIS AND RICHARD R. BURGESS

this enzyme on a supercoiled plasmid DNA that contained the yeast alcohol dehydrogenase gene (149, 150). A minor transcript starting at or very near the correct in vivo initiation site was detected but the major transcript started elsewhere. This selective transcription was dependent on the DNA being in a supercoiled form and, since it also required Mn2+ and high levels of the initiating dinucleotide primer UpA, it is not clear that the observed restriction of enzyme site selection has physiological relevance. Results showing apparent selective transcription of ribosomal gene sequence in total yeast DNA by yeast RNA polymerase I have been presented (163, 164). However, the complexity of the template being transcribed and the fact that the in vitro RNAs were analyzed only by indirect methods make these studies difficult to interpret. Later studies using cloned yeast ribosomal DNA suggest that yeast polymerase I does not initiate transcription in vitro at the presumptive in vivo initiation site (165). Analysis of RNA synthesized by purified HeLa cell RNA polymerase I11 on adenovirus 2 DNA indicated random transcription of this template (148). This lack of specificity by purified RNA polymerase I11 has been confirmed (166).

E. OTHERREACTIONS In addition to catalyzing the synthesis of free RNA that contains a 5’-triphosphate terminus, and synthesizing RNA covalently linked to template DNA by extending the 3‘-hydroxyl primer at a single-strand nick (133), wheat germ RNA polymerase I1 has also been shown (167) to incorporate the dinucleotide ApU into the 5‘ ends of RNA chains. Under low substrate conditions, various other dinucleotides have been shown to stimulate the enzyme (168, 169). It is not clear whether the eukaryotic RNA polymerases catalyze the “abortive initiation” reaction observed with prokaryotic RNA polymerase (1 70), characterized by the synthesis and release of many 163. Van Keulen, H . V., and Retel, J. (1977). EJB 79, 579. 164. Holland, M. J . , Hager, G . L . , and Rutter, W. J . (1977). Biochemistry 16, 16. 165. Sawadogo, M . , Sentenac, A . , and Fromageot, P. (1981). BBRC 101, 250. 166. Weil, P. A., Segall, J., Harris, B . , Ng, S . Y . and Roeder, R. G. (1979). JBC 254, 6163. 167. Yarbrough, L. R. (1981). J B C , in press. 168. Shaw, P. A., and Saunders, G. R. (1979). FEBS Lett. 1106, 104. 169. Shaw, P. A., Marshall, M. V., and Saunders, G . F. (1980). Cytogenet. Cell Gener. 26, 211. 170. Johnston, D. E., and McClure, W. (1976). In “RNA Polymerase” (M. Chamberlin and R. Losick, eds.), pp. 413-428. Cold Spring Harbor Laboratory, Cold Spring Harbor, New York.

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molecules of dinucleotide or trinucleotide product per enzyme molecule. One report is in the affirmative for yeast RNA polymerase I1 (149), whereas another is in the negative for wheat germ RNA polymerase I1 (167). The use of different templates and assay conditions may explain this discrepancy. In addition to catalyzing DNA-dependent RNA synthesis, RNA polymerase 11from tomato and wheat germ also catalyze RNA-dependent RNA synthesis in the presence of Mn2+ (171). This reaction was first described for prokaryotic RNA polymerase (172). Although RNAdependent RNA synthesis is catalyzed by the eukaryotic enzymes two orders of magnitude less efficiently than DNA-dependent synthesis, evidence has been presented for specific initiation on circular viroid RNA templates (171 ). VII.

Inhibitors

A. AMATOXINS Though eukaryotic RNA polymerase is inhibited by substrate analogs and by agents that act on template DNA we discuss here only inhibitors acting on the polymerase itself. A comprehensive volume has been published dealing with inhibitors of DNA and RNA polymerases, and includes sections on the eukaryotic enzymes (173). a-Amanitin, a member of the amatoxin family, is the most commonly used inhibitor of eukaryotic RNA polymerase. In addition, derivatives of the prokaryotic RNA polymerase inhibitor, rifamycin, have been used to inhibit the eukaryotic enzymes. Amatoxins (a-, ,B-, and y-amanitin, amanin, and amanullin) are potent inhibitors of eukaryotic RNA polymerase 11, 50% inhibition occurring at concentrations at low as 0.05 micrograms/ml (58). The class I11 enzymes are also sensitive (11) but require higher inhibitor levels (5- 10 pglml) to effect 50% activity inhibition. In general, class I enzymes are insensitive. Yeast is exceptional in having a class I enzyme that is slightly amanitinsensitive, a class I11 enzyme that is completely resistant, and a class I1 enzyme that is only as sensitive as most class I11 enzymes (I). In addition, both class I and I11 enzymes fromBombyx mori are completely resistant to high levels of toxin (1). I n vivo, low concentrations of a-amanitin (0.1 to 1.O pglml) selectively inhibit synthesis of poly(A)+ messenger RNA in 171. Rackwitz, H . R., Rohde, W., and Sanger, H. L. (1981). Nntitre (London) 291, 297. 172. Fox, C. F., Robinson,W. S., Haselkom, R . , and Weiss, S. B. (1964).JBC 239, 186. 173. Sarin, P. S. and Gallo, R. C. (eds). (1980). “Inhibitors of DNA and RNA Polymerases” Pergamon, New York.

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germinating wheat embryos (174). Higher concentrations were found to inhibit 5 S and tRNA synthesis (174) in a manner consistent with the in v i m results. The amatoxins are bicyclic octapeptides found in the poisonous mushroom Arncinira phrilloides ( 1 75). Their molecular weights are about 1000 daltons. a-Amanitin is the most readily available and frequently used derivative. P-Amanitin and amanin contain a free carboxyl group (175) and are thus suited for carbodiimide coupling to proteins or column supports. A uniquely amanitin-resistant RNA polymerase I1 is found in the fungus Aspergillus nidulnns ( 1 7 5 ~ )It. has been reported (176, 177) that the amanitin sensitivity of RNA polymerase I1 isolated from Amanira species is not as great as that of enzymes isolated from plant or mammalian tissue. Amanitin concentration is measured spectrophotometrically (molar extinction coefficient = 15,400 at 305 nm) (178). Its ultraviolet absorbance is due to the presence of a sulfoxytryptophan residue that bridges the molecule ( / 75). Scission of a peptide bond or destruction of a bridge bond will destroy the toxicity (179) but not the absorbance. It has been shown that a-amanitin inhibits transcription by blocking the elongation reaction (178). a-Amanitin binds extremely tightly to RNA polymerase 11. Only one inhibitor molecule binds per enzyme molecule (178). The dissociation half-time of complexes between a radioactive amanitin derivative and calf thymus RNA polymerase I1 was found to be about 100 hr at 0" (178). The ability to titrate RNA polymerase I1 with labeled amanitin has allowed an estimate of the number of enzyme molecules in crude cell extracts (180).

B. RIFAMYCIN DERIVATIVES Rifampicin inhibits bacterial RNA polymerase by blocking elongation of the initiated dinucleotide but has no effect if elongation has proceeded past this point (170). Although eukaryotic RNA polymerases were found 174. Jendrisak, J. (1980). JBC 255, 8529. 175. Fiume, L., and Wieland, T. (1970). FEBS Lett. 18, 1 . 175a. Stunnenberg, H. G . , Wennekes, L., Spierings, T., and vanden Broek, H. (1981). EJB 117, 121. 176. Johnson, B. C., and Preston, J . F. (1979). Arch. Mirrobiol. 122, 161. 177. Johnson, B. C . , and Preston, J. F. (1980). EBA 607, 102. 178. Cochet-Meilhac, M . , and Chambon, P. (1974). BBA 353, 160. 179. Wieland, T., and Wieland, 0. (1959). Plrnrmcicol. Rev. 11, 187. 180. Cochet-Meilhac, M., Nuret, P., Courvalin, J. C., and Chambon, P. (1974). BEA 353, 185.

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to be refractory to inhibition by rifampicin, their activity was affected by several semisynthetic rifamycin derivatives (181). The structure and action of these derivatives has been discussed extensively (173). Though it seemed that these compounds could be acting nonspecifically as detergents due to their hydrophobic substituents (181, 182), one derivative, rifamycin AF/013, was shown to specifically inhibit the initiation reaction when present at low concentration (182). As pointed out, however the mode of action of rifamycin AFiO13 resembles that of heparin (181). It has been shown (183) that this compound destabilizes DNA-bound histone, while rifampicin does not. VIII.

A.

Eukaryotic Transcription Extract Systems

DISCOVERY O F TRANSCRIPTION EXTRACTS

The discovery by Wu (184) that the virus-associated (VA) RNAl gene of adenovirus 2 DNA was correctly transcribed by the RNA polymerase I11 in translation extracts derived from human KB cells is a landmark in the field of in i t t m eukaryotic transcription. This finding was the first demonstration of the correct transcription of exogenous DNA by a eukaryotic RNA polymerase in a soluble cell-free system. This circumstance was accompanied by the exciting possibility of purifying the factors necessary for the faithful transcription of a eukaryotic gene. Since the original announcement, cloned 5 S and tRNA genes of various kinds have been shown to be correctly transcribed in crude extracts of Xenopus oocytes and cultured mammalian cells (166, 185-187). It was found (38)that supplementation of the Wu extract as modified by Weil et crl. (166), with purified RNA polymerase 11, resulted in specific transcription by the exogenous enzyme from the adenovirus 2 late promoter. This was demonstrated by several independent techniques. In v i m transcripts were found to be capped at the same sites seen on in vivo-made mRNAs (166). Subsequently, Manley et d.(188)showed that a whole-cell 181. Meilhac, M., Tysper, A,, and Charnbon, P. (1972). EJB 28, 291. 182. Riva, S.,Fietta, A . , and Silvestri, L. G . (1972). BBRC 49, 1263. 183. Lilley, D. M. J . , Jacobs, M. F., and Houghton, M . (1979). Nucleic Acids Re.\. 7,377. 184. Wu, G. J . (1978). P N A S 75, 2175.

185. Mattoccia, E., Baldi, M. I . , Carrara, G., Fruscoloni, P., Benedetti, P., and Tocchini-Valentini, G. P. (1979). C P / /18, 643. 186. Garber, R . L. and Gage, P. L. (1979). Cell 18, 817. 187. Hagenbuchle, O., Larson, D., Hall, G . I . , and Sprague, K . U. (1979). Cell 18, 1217. 188. Manley, J . L . , Fire, A , , Cano, A , , Sharp, P. A., and Gefter, M. L. (1980). !“AS 77, 3855.

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HeLa extract, prepared by modification of the method of Sugden and Keller (189) for isolation of HeLa RNA polymerase 11, was in itself sufficient to accurately transcribe the adenovirus DNA. In both cases, specific transcription was demonstrated in “run-off’’ assays (190) that utilized DNAs cut with restriction enzymes to generate transcripts terminating at a restriction fragment end. These systems have been used to show specific in vitro transcription from the presumptive in vivo promoters of SV40 ( 1 91, 192), cloned P-globin (193, 194), conalbumin ( 1 9 9 , and ovalbumin (195) genes. Recently Weingartner and Keller (1950) have prepared HeLa cell extracts which can utilize as a template a circular plasmid carrying a cloned fragment of early region I11 adenovirus 2 DNA to produce transcripts which are apparently initiated and spliced correctly. An RNA polymerase I extract system is also now available (I 96). Crude extracts of Ehrlich ascites tumor cells have been reported to accurately transcribe cloned mouse ribosomal DNA in a reaction that is insensitive to a-amanitin ( I 96).

B. FRACTIONATION OF TRANSCRIPTION EXTRACTS Factors responsible for the specific transcription of Xenopus 5 S rDNA genes by RNA polymerase 111 have been fractionated from oocyte extracts (173). One of these factors has been purified to homogeneity and is a basic protein of molecular weight 40,000 (197). This protein was shown to bind specifically to an internal region in 5 S DNA (I 97) that had been shown by transcriptional analysis of in vitro deletion mutants to be essential for accurate transcription initiation (198). Interestingly, the same protein was found to be a component of oocyte 7 S ribonucleoprotein parti189. Sugden, B . , and Keller, W. (1973). JBC 248, 3777. 190. Wickens, M. P., and Laskey, R. A. (1981). In “Genetic Engineering” (R. Williamson, ed.), Vol. I , pp. 103- 167. Academic Press, New York. 191. Rio, D., Robbins, A,, Myers, R . , and Tjian, R. (1980). PNAS 77, 5706. 192. Handa, H . , Kaufman, R. J . , Manley, J., Gefter, M., and Sharp, P. A. (1981). JBC 256, 478. 193. Luse, D. S . , and Roeder, G . C. (1980). Cell 20, 691. 194. Talkington, C. A., Nishioka, Y., and Leder, P. (1980). PNAS 77, 7132. 195. Waslyk, B., Kedinger, C., Corden, J., Bison, O., and Chambon, P. (1980). Nature (London) 285, 367. 195a. Weingartner, B., and Keller, W. (1981). PNAS 78, 4092. 196. Grummt, I. (1981). PNAS 78, 727. 197. Engelke, D. R., Ng, S. Y., Shastry, B. S. , and Roeder, R. G. (1980). Cell 19, 717. 198. Bogenhagen, D. F., Sakonju, S . , and Brown, D. D. (1980). Cell 19, 27.

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I49

cles, and to bind to purified 5 S RNA (199, 200). This binding has been proposed as a possible autoregulatory mechanism, since 5 S RNA inhibited its own synthesis when added to thein vitro transcription system ( I99, 200). In addition to this factor and RNA polymerase 111, at least two other separable components have been judged essential for accurate transcription of 5 S genes ( I97). The purified oocyte factor is apparently not needed for transcription of tRNA genes ( I97). The 40,000 dalton specific DNA-binding RNA polymerase I11 factor constitutes about 15% of the soluble protein in oocytes (200, 201), a circumstance contributing to the ease of its complete purification. Purifications of the accessory factors and those involved in transcription by RNA polymerase I1 have proved more difficult. These latter factors have been partially purified from KB and HeLa cells (202, 203), an undertaking of some magnitude given that 500 liters of culture are needed to obtain 1 Kg of starting material. The studies indicate that at least 3-4 different components are needed in addition to RNA polymerase I1 for the selective transcription of adenovirus late genes. Fractionation efforts are currently hampered by the lack of large quantities of starting tissue. Suitable RNA polymerase I1 extracts are now available only from some cultured mammalian cells. Development of a soluble transcription extract system from sources such as yeast, wheat germ, or calf thymus would greatly expedite purification studies.

C. DO EXTRACTS MIMIC in Vivo TRANSCRIPTION? The ability of eukaryotic transcription extracts to mimic in vivo events is a matter of some concern. In particular, the question arose as to whether the capped in vifro transcripts resulted from specific transcription initiation or from specific RNA processing events. It was not clear, furthermore, that a mRNA cap site represented a start point for in vivo RNA synthesis. The demonstration of an activity in vaccinia virus cores that phosphorylated RNA 5'-monophosphate termini (204), strengthened the possibility that new 5' ends could be generated in vivo by phosphorylation and capping of RNA species arising from the cleavage of larger precursor molecules. That cap sites do, however, represent sites of transcription initiation in vivo for at least some transcripts has been strongly suggested 199. 200. 201. 202. 203. 204.

Honda, B . M., and Roeder, R. G. (1980). Cell 22, 119. Pelham, H. R., and Brown, D. D. (1980). PNAS 77, 4170. Picard, B . , and Wegnez, M . (1979). PNAS 76, 241. Matsui, T., Segall, J., Weil, P. A . , and Roeder, R. G. (1980). JBC 255, 11911. Dynan, W. S . and Tjian, R . (1981). ICN-UCLA S y m p . , in press. Spencer, E., Loring, D., Hurwitz, J . , and Monroy, G. (1978). PNAS 75, 4793.

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by studies of late SV40 RNAs synthesized in permeabilized CV-1 cells in the presence of P-32P-labelednucleotides (205). Analogous studies have shown that the capped 5' ends of transcripts synthesized in vitro on adenovirus and P-globin DNA template do represent sites of transcription initiation (206). Several studies (207-209) indicate that RNA polymerase I1 extract systems recognize the so-called Goldberg-Hogness or TATA box, a DNA sequence of the form TATAAAT, which is found 25-30 bp upstream of the mRNA cap site in many eukaryotic genes. Deletion or mutation of this sequence abolishes in vitro transcription, In vivo however, the situation is more complex since a sequence very similar to this is dispensable for the synthesis of SV40 early RNA in vivo, and another sequence exists 70- 155 bp upstream from the mRNA cap site that is required for in vivo but not for in vitro expression (209, 210). Eukaryotic transcription extract systems do not generally exhibit the regulation that is seen in v i v a Uninfected cell extracts recognize the adenovirus late promoter and extracts of Xenopus oocytes transcribe somatic 5 S RNA genes that are not normally expressed at high levels in this developmental stage. However, SV40 T antigen has been shown to block early SV40 transcription in vilro, mimicking its in vivo effect (191). IX.

Organelle- and Viral-Coded

RNA

Polymemses

A. MITOCHONDRIAL RNA POLYMERASE The mitochondria1 RNA polymerases have been partially purified from yeast (211-216), Nnirospcm c r m w (217), Xenopris laevis (218), and rat liver (219). The topic is treated only briefly here. For more detailed stud205. Contreras, R . , and Fiers, W. (1981). Nircleic Acids Res. 9, 215. 206. Hagenbuchle, O . , and Schibler, U. (1981). P N A S . in press. 207. Corden, J . , Wasylyk, B . , Buchwalder, A., Sassone-Corsi, P., Kedinger, C . , and Chambon, P. (1980). Science 209, 1406. 208. Wasylyk, B., Derbyshire, R . , Guy A,, Molko, D . , Roget, A., Teoule, R . , and Chambon, P. (1980). PNAS 77, 7024. 209. Mathis, D. J . , and Chambon, P. (1981). NrJture (London) 290, 310. 210. Benoist, C., and Chambon, P. (1981). Nrrtitrr (London) 290, 304. 211. Wattiaux, R., and Waittiaux-De Coninck, S. (1970). BBRC 40, 1185. 212. Scragg, A. H . (1971). BBRC 45, 701. 213. Tsai, M. J., Michaels, G ., and Criddle, R. S. (1971). PNAS 68, 473. 214. Wintersberger, E. (1972). BBRC 48, 1287. 215. Rogall, G., and Wintersberger, E. (1974). FEBS Lett. 46, 333. 216. Levens, D., Lustig, A., and Rabinowitz, M. (1981). JBC 256, 1474. 217. Kuntzel, H . , and Schafer, K. P. (1971). Nntrire Neb$,B i d . 231, 265. 218. Wu, G . J . , and Dawid, I. B . (1972). Biorhemistry 11, 3589. 219. Mukerjee, H . , and Goldfeder, A. (1973). Biochemistry 12, 5096.

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ies, the reader should consult the review article (220) dealing with this subject alone. Solubilization of mitochondrial RNA polymerases has proved somewhat ditficult, requiring such techniques as high-salt mechanical disruption or treatment with detergents (see above Refs.). Once solubilized, however, the enzyme is amenable to purification by standard chromatographic techniques. The difficulty in solubilization suggests that the enzyme may be tightly bound within the organelle, possibly associated with membrane structures. Conflicting reports have emerged as to whether the yeast enzyme is sensitive to rifampicin (211-216). It is possible that bacterial contamination of yeast cultures is responsible for this state of affairs. Insensitivity to a-amanitin has been uniformly reported. The purest mitochondrial RNA polymerase preparations have been reported to exhibit only one major band upon electrophoresis on polyacrylamide gels in the presence of SDS. This putative enzyme band had a molecular weight of 46,000 in Xenoprrs kievis (218), 450,000 in yeast (216), 64,000 in Nrtiiosporui (217), and 66,000 in rat liver (219). In no case was it shown that the major protein band in these preparations had enzymatic activity. However, antibody directed against the gel-purified 45,000 dalton polypeptide from yeast inhibited purified mitochondrial RNA polymerase activity, but not the activity ofE. coli RNA polymerase (216). Workers should be wary of possible contamination with the nuclear RNA polymerases, which can give rise to multiple peaks of activity upon chromatography of extracts from isolated mitochondria (217). Because of aggregation of mitochondrial polymerase induced by low-salt conditions, multiple sedimenting forms may also be observed (217, 218). It is not clear whether the mitochondrial RNA polymerases are coded by the organelle genome or by nuclear chromosomal DNA. Little is known of the template properties of these enzymes or of their possible ability to selectively transcribe mitochondrial DNA. Preliminary work suggests, however, that purified mitochondrial RNA polymerase from yeast may transcribe yeast mitochondrial DNA in a nonrandom fashion (216).

B. CHLOROPLAST RNA POLYMERASE RNA polymerase has been purified to apparent homogeneity from Zea mays chloroplasts (221). The enzyme has a subunit structure consisting of 220. Wintersberger, E. (1973). I n “Regulation of Transcription and Translation in Eukaryotes,” 24. Mosbacher Colloquim (E. K . F. Bautz, ed.), pp. 179-193. Springer Verlag Berlin-Heidelberg, New York. 221. Smith, H . J . , and Bogorad, L. (1974). P N A S 71, 4839.

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two large subunits of 200,000 and 180,000 daltons as well as several smaller polypeptides (221). Peptide mapping studies (222) have shown that the 180,000 dalton subunit from the chloroplast enzyme is different from the 180,000 dalton subunit of maize nuclear RNA polymerase 11. It was not demonstrated, however, that the complex subunit structure of the chloroplast enzyme was not due to contaminating nuclear RNA polymerase I or 111. A stimulatory protein of about 30,000 daltons has been purified from chloroplasts (223).This peptide, called S factor, elutes from DE52 resin at 0.5 M KCI, whereas the chloroplast RNA polymerase elutes at 0.2M KCI. S factor stimulates transcription of supercoiled DNA by the chloroplast polymerase, but apparently has no effect on the level of transcription from a relaxed template (223). A transcriptionally active chromosome has been isolated from chloroplasts of Eirglena gracilis (224, 225) and of spinach (226). The major transcript in vitra was shown to be ribosomal RNA and preliminary evidence for selective initiation was presented (227). A transcriptionally active complex, purified by gel filtration in high salt, had a specific activity comparable to E. coli holoenzyme and exhibited a relatively simple protein band pattern on SDS-polyacrylamide gels (major bands of 125,000 and 50,000 daltons and a minor band of 47,000 daltons) (228).

c.

VACCINIA

VIRUS

RNA POLYMERASE

DNA-dependent RNA polymerase from vaccinia virus has been purified from the cytoplasm of virus-infected HeLa cells (229) and from virions (230). The purified enzyme has seven subunits of 135,000, 130,000, 77,000,34,000,19,500,16,500, and 13,500 daltons, each present in approximately equal stoichiometry (229). The enzyme is insensitive to both a-amanitin (229-230) and rifampicin (229, 230) and much prefers denatured over double-stranded DNA templates (229, 230). 222. Kidd, G. H., and Bogorad, L. (1979). P N A S 76, 4890. 223. Jolly, S. O . , and Bogorad, L. (1980). P N A S 77, 822. 224. Hallick, R. B. and Lipper, C. (1976). I n “Molecular Mechanisms in the Control of Gene Expression” (Donald P. Nierlich, W. J. Rutter, C. Fred Fox, eds.). Academic Press, New York. 225. Hallick, R. B . , Lipper, C . , Richards, 0. C., and Rutter, W. .I. (1976). Biochemistry 15, 3039. 226. Briat, J . F., Laulhere, J. P., and Mache, R . (1979). EJB 98, 285. 227. Rushlow, K . , Orozco, E . , Lipper, C . , and Hallick, R. (1980). JBC 255, 3786. 228. Rushlow, K. (1980). Ph.D. Thesis, University of Colorado, Boulder, Colorado. 229. Nevins, J. R., and Joklik, W. K. (1977). JBC 252, 6930. 230. Spencer, E., Shuman, S . , and Hurwitz, J. (1980). JBC 255,5388.

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ACKNOWLEDGMENTS This work was supported by NIH Program Project Grant CA-23076, by NIH Core Grant CA-07175, and NIH Training Grant CA-09135. We thank numerous colleagues for providing unpublished research results and Ann B. Burgess, William S . Dynan, and Jerry J. Jendrisak for many helpful suggestions.

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Priming Enzymes EDMUND W. BENZ, JR. JERARD HURWITZ

DANNY REINBERG

I. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . 11. dnuG Gene Product from Esclrrrichicr coli . . . . . . . . . . . .

111. IV. V. VI. VII.

VIII.

1.

A. Assay and Purification . . . . . . . . . . . . . . . . . . B. Catalytic Properties . . . . . . . . . . . . . . . . . . . . Multiple Pathways for dncrG Priming in SS to RF DNA Replication Studies on the Specificity of dnrrG-Template Interactions . . . . . Priming by RNA Polyrnerases . . . . . . . . . . . . . . . . . Priming on Double-Stranded DNA . . . . . . . . . . . . . . . Phage-Encoded Priming Enzymes . . . . . . . . . . . . . . . A. Bacteriophage T7 . . . . . . . . . . . . . . . . . . . . . B. Bacteriophage T4 . . . . . . . . . . . . . . . . . . . . . Priming in Eukaryotic Systems . . . . . . . . . . . . . . . . .

155 156 156 157 160 165 166 169 174 174 176 178

Introduction

Unlike RNA polymerases, DNA polymerases cannot begin synthesis de n o w ; these polymerases must have, in addition to a template upon which to generate a progeny strand, availability of a free 3’-OH to serve as acceptor for the transfer of the first nucleotide and formation of the first phosphodiester bond. The molecules that provide the required 3’-OH have been termed primers. Without primers DNA replication cannot initiate, thus priming becomes an important regulatory control point. Recent studies on the replication of small, circular single-stranded DNA bacteriophages have demonstrated that the complexity of DNA replication resides primarily in the complexity of the events governing the 155 THE ENZYMES. VOL. XV Copyright @ 1982 by Academic Press. lnc. All rights of reproduction in any form reserved. ISBN 0-12- I227 15-4

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E. W. BENZ, JR., D. REINBERG, AND J. HURWITZ

specificity of initiation. The small DNA chromosomes from phages that infect E. coli (phages a3, st-I, 4K, G4, 4x174, fd, and M13) can be replicated in vitro with purified or partially purified proteins; the first step is the conversion of the positive strand of circular DNA to its duplex replicative form ( 1 -6). This event requires the de novo initiation of negative strand synthesis and in many respects is analogous to Okazaki fragment formation during bacterial DNA replication, particularly since only host proteins are involved (7). Using these phage DNAs and complementation of thermolabile E. coli extracts, the E. coli gene product dnaG was identified as the protein that synthesizes the primer molecule during initiation of DNA synthesis in RNA polymerase-independent pathways (8-11). Unlike RNA polymerase, dnaG protein activity is resistant to the antibiotic rifampicin (12). In this chapter we discuss the enzymology of the dnaG protein from E. coli (also called primase) since this protein has been well-studied; moreover, we compare the properties and modes of action of this protein with other proteins thought to have a dnaG-like role in other systems. II.

dnaG Gene Product from Ercherichia coli

A. ASSAYA N D

PURIFICATION

1. Methods ($Assay

Catalysis of reaction can be determined by measuring either the formation of primer or the elongation of primer-DNA complexes into RFII 1. Wickner, S. (1978). In “The Single-Stranded DNA Phages” (D. T. Denhardtl, D. Dressler, D. S. Ray, eds.), pp. 255-271. Cold Spring Harbor Laboratory, Cold Spring Harbor, New York. 2. Wickner, S. (1978). Annu. Rev. Eiochem. 47, 1163-1191. 3. Sumida-Yasumoto, C., Ikeda, J.-E., Benz, E., Marians, K. J., Vicuna, R., Sugrue, S., Zipursky, S. L., and Hunvitz, J. (1978). CSHSQB 43, 311-329. 4. McMacken, R., Rowen, L., Ueda, K., and Kornberg, A. (1978). In “The SingleStranded DNA Phages” (D. T. Denhardt, D. Dressler, and D. s. Ray, eds.), pp. 273-285. Cold Spring Harbor Laboratory, Cold Spring Harbor, New York. 5. Schekman, R., Weiner, A., and Kornberg, A. (1974). Science 186, 987-993. 6. Kornberg, A. (1978). C S H S Q B 43, 1-9. 7. Ogawa, T. and Okazaki, T. (1980). Annu. Rev. Bichem. 49, 421-457. 8. Wickner, S. (1977). PNAS 74, 2815-2819. 9. Rowen, L., and Kornberg, A. (1978). JBC 253, 758-64. 10. Rowen, L. and Kornberg, A. (1978). JBC 253, 770-774. 11. Benz, E. W., Jr., Reinberg, D., Vicuna, R., and Hurwitz, J. (1980). JBC 255, 10961106. 12. Bouche, J.-P., Zechel, K., and Kornberg, A. (1975). JEC 250, 5995-6001.

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157

structures. The first assay measures the amount of oligonucleotide synthesized by the dnaG protein in the presence of circular, single-stranded phage DNA that is coated with DNA-binding protein (DBP). The second measures the rate of DNA synthesis due to extension by the DNA polymerase I11 elongation proteins of primer-DNA complexes formed by dnaG protein during the reaction. This elongation reaction can be performed in incubations containing either purified proteins or crude extracts of E. coli. Complementation assays are variations on the elongation assay using E. coli extracts that are thermolabile for DNA synthesis and rendered dependent on exogenous DNA and the addition of the wild-type complementing protein. Whether the product is an oligonucleotide or a long chain molecule, it is hybridized to the input circular DNA and can be separated from the precursor molecules by gel filtration chromatography. In the case of the elongation assay, the product can be recovered by acid precipitation. These two assays have been called the uncoupled and coi4ppled assays ( I I ) . In the coupled assay, one unit ofdnaG activity corresponds to one nmol of dTTP incorporated into acid-insoluble product in 20 min at 37".

2 . Pitrijiccition and PI1ysical Properties The dnaG protein has been purified from extracts of gently lysed E. coli HMS-83 cells (9, 11). A 24,000-fold purification from an extract of 400 g of net cells gave 320 pg of purified enzyme, representing 8% of the original activity of the extract ( I f ) . The purified preparation contained a single major polypeptide of 64,000 MW (SDS gel electrophoresis) and was free of contaminating enzymatic activities. As determined by a high-salt glycerol-gradient centrifugation, the dnuG protein has a native MW of about 60,000, suggesting it is composed of a single subunit (8, 1 1 ) . The enzyme has an isoelectric constant of 5.9 at 8". The activity is resistant to N-ethylmaleimide treatment and binds weakly to single-stranded DNA. B. CATALYTIC PROPERTIES 1. Synthesis de Novo of cin Oligoribonrrcleotide Primer When the DBP-coated template molecule is single-stranded circular DNA from phages a3, 4 K , st-1, or G4 and only the four ribonucleoside triphosphates are present, dnaG protein synthesizes an 11-28 base-long oligoribonucleotide that remains base-paired to template DNA (9, 11). In a subsequent step, the primer hybridized to the template can be elongated by DNA polymerase I11 (and its elongation system consisting of DNA EFI, DNA EFIII, and dnaZ protein) forming the gapped, duplex DNA

158

E. W. BENZ, JR., D. REINBERG, AND J. HURWITZ

molecule, RFII (8, 9, I I ) . On these template DNAs the formation of a single primer molecule is sufficient to support the synthesis of nearly full-length minus strand DNA. Elongation does not depend on the continued availability of either dna G protein or DBP since deproteinized primer DNA molecules can be readily elongated ( / I ) . No other proteins are required for in vitro priming when these phage DNAs are used as templates. The sequences of these primer oligonucleotides are complementary to an 11 to 28 base-long region at the origin of minus strand replication. 2. Deoxyriircleosicie Triphosphcites Alter Primer Formation Addition of deoxynucleoside triphosphates to the priming reaction mixture containing the ribonucleoside triphosphates changes the length, yield, and composition of primers formed ( 8 4 1 , IS). Although the protein requirement and the stoichiometry of protein to template do not change, the presence of deoxynucleotides in the reaction reduces the rate of primer formation by as much as fivefold, and the length of primer twofold (from 26 to 13 bases long). Only after prolonged incubation (60 min) at 37” do full-length primer molecules appear. This effect occurs even at a 10: 1 ratio of ribo- to deoxyribonucleotides ( II). Deoxynucleotide precursors are incorporated into these primer molecules, forming copolymers of riboand deoxyribonucleotide moieties ( / I , 14). These chains can be labeled with both radioactive ribo- and deoxyribonucleotide precursors ; alkaline hydrolysis of such primers show ribonucleotides covalently linked to deoxyribonucleotides via a phosphodiester bond. The effect of deoxyribonucleotide precursors is to substitute, base for base, for ribonucleotides, with subsequent extension or completion of that chain being limited but not completely prevented. The same dominance and chain-shortening efEect is seen in the coupled reaction, where the rib0 primers are predominantly di- and trinucleotides, but whether this is completely due to deoxy dominance or protein-protein interactions between dnciG and the elongation proteins is not known ( 1 1 , 14). The initial nucleotide could be a deoxynucleotide (dATP), but the rate is manyfold greater if the first base is a ribonucleotide (ATP) (9, I I). Ribo-, deoxy-, or ribodeoxy copolymer primers serve equally well to prime DNA polymerase elongation of deproteinized primer DNA complexes. 13. McMacken, R . , Bouche, J.-P., Rowen, S. L., Weiner, J. H., Ueda, K., Thelander, L., McHenry, C., and Kornberg, A. (1977). I n ”Nucleic Acid-Protein Recognition” (H. J. Vogel, ed.), pp. 15-29. Academic Press, New York. 14. Bouche, J.-P., Rowen, L., and Kornberg, A. (1978). JBC 253, 765-769.

159

6. PRIMING ENZYMES TABLE I

Y I E L DO F P R I M EFORMED R BY r h G "

drmG protein added (fmol)

Active origin DNA DBP-coated (fmol)

0 13 27 80 165

60 60 60 60 60

Primer formed (fmol) ~~

~

0 16 32 45 50 ~

~

~

~~~~~

" Reactions are as described in the text. Each primer molecule was composed of 10 GMP residues as dictated by the sequence at the origin of 0 3 minus strand DNA synthesis.

3. Stoicliiometty of dnuG-Dependent Primer Formation

The dnuG enzyme is present in the wild-type bacterial cell at very low concentrations (about 5 to 25 molecules per cell), whereas replication of chromosomal DNA involves the formation of thousands of Okazaki fragments (7). However, in v i t w the molar yield of primer never exceeds the molar amount ofdnaG protein added (11, 15). (See Table I.) Thus, in vitro dnaG does not behave catalytically. Similar observations have been made for both the uncoupled and coupled reactions (11). 4. ADP Cun Replace ATP

When ADP is added to the priming reaction in place of ATP, no change in the rate or the extent of synthesis is observed, nor is the length of the primer formed changed (8, 11). [cx-~'P]ADPas radioactive precursor serves equally well as ATP. Since there are six adenine residues dictated by the sequence, labeling at the 5' terminus, as opposed to incorporation internally, would be reflected by a decrease in overall incorporation of label. This is not observed. Primers labeled in reactions with [cx-~'P]ADP have the same specific activity as those labeled with [cx-~'P]ATP,are 65% resistant to bacterial alkaline phosphatase treatment, and have a diphosphate 5' terminus; alkaline hydrolysis of these primer molecules yield "P-labeled AMP, GMP, and ppAp, indicating internal phosphate transfers (11). It is likely then that ADP serves as the donor of the first adenine as 15. Benz, E. W., Jr., Sims, J . , Dressler, D . , and Hurwitz, J. (1980). I n "ICN-UCLA Winter Symposium on Mechanistic Studies on DNA Replication and Recombination'' (B. Alberts, ed.), pp. 279-291. Academic Press, New York.

160

E. W. BENZ, JR., D. REINBERG, AND J. HURWITZ

well as internal adenines during primer formation. ADP is the only diphosphate that can substitute for its homologous triphosphate (8, 11). Although the activity of dnaG protein is inhibited by inorganic pyrophosphate (and by inorganic phosphate), no exchange reaction of ["2P]PPiinto nucleoside triphosphates was demonstrated (1 I). 111.

Multiple Pathways for dnaG Priming in SS to RF DNA Replication

During the replication of the single-stranded (SS) DNA phages (a3,st-1,

4K,G4, and 41x174) to replicating form (RF) DNA, after a primed circle has been formed the DNA polymerase 111elongation machinery (consisting of DNA elongation factor I, the dnaZ gene product, DNA elongation factor 111, and DNA polymerase 111) can extend the primer to yield gapped, duplex circular DNA that can be filled in by DNA polymerase I and sealed by E. coli DNA ligase to form the relaxed, covalently closed, circular duplex molecule RFI' (2, 16-19). This product can be converted to superhelical RFI DNA by the DNA gyrase system (20,21).Evidence has further suggested that the complex of proteins assembled during priming is conserved throughout RF replication and that this complex can introduce one or more supercoils in the resultant RF product (22, 23, 24). The replication of the phage DNAs described above differs in the pathway each employs to form a primed circle. In each case saturating amounts of the single-stranded DNA binding protein (DBP) from E. coli are required. Whereas the DBP is sufficient to promote primer synthesis by the dnaG gene product on a3, st-1, $K,and G4 DNAs, other proteins are necessary for proper priming in the case of +X174 DNA. With 4x174 DNA an RNA polymerase-independent pathway is followed that requires the action of at least six different proteins (replication factors X, Y, and Z, E. coli single-stranded DBP, dnaB protein and dnaC protein) to act on the 16. Tomizawa, J., and Selzer, G. (1979). Annu. Rev. Biorhern. 48, 999-1034. 17. Hurwitz, J . (1979). CRC Rev. Microbiol. 18. Gefter, M. L. (1975). Annu. Rev. Eiochem. 44, 45-78. 19. Geider, K. (1976). C u n . Topics Microbiol. lrnrnrtnol. 74, 55-112. 20. Gellert, M., Mizuuchi, K., O'Dea, M. H., and Nash, H. A. (1976). PNAS 73, 38723876. 21. Marians, K. J., Ikeda, J.-E., Schlagman, S., Hunvitz, J. (1977). PNAS 74, 19651969. 22. Arai, K., Low, R., and Kornberg, A. (1981). PNAS 78, 707-711. 23. Low, R. L., Arai, K., and Kornberg, A. (1981). PNAS 78, 1436-1440. 24. Arai, K., Low, R., Kobori, J., Shlomai, J., and Kornberg, A. (1981). JBC 256, 5273-5280.

6.

PRIMING ENZYMES

161

DNA template in order for subsequent primer formation to occur (25-33). The primer molecule itself is synthesized by the dnuG protein (8, 30). Since all of these proteins are host-encoded, they can act by a similar mechanism during bacterial DNA replication. The dna G-dependent pathways for the formation of primer molecules on single-stranded templates are distinguished by three different mechanisms. The simplest and least specific priming pathway involves a general mechanism that cannot operate in the presence of DBP but has a very simple protein and template requirement. This general mechanism requires only dnaB and dnaG proteins and can operate on virtually any DNA template that is not covered by DBP (34,35).In an ATP-dependent reaction a stable dnn B protein-DNA complex is formed that can be recognized by the dnaG protein, which can then form primers at many nearly random sites. Recent evidence suggests that ATP (and its nonhydrolyzable analogs) serve as an allosteric effector in forming a stable ternary dncr B-ATP-DNA complex; hydrolysis of ATP leads to destabilization of the complex (22,36-38). In this pathway the dnaG protein can apparently interact at many sites, synthesizing oligoribonucleotides from 10 to 60 bases in length. This pathway is nonprocessive and is precluded by the presence of DBP (34). When DBP is present two additional, more specific, pathways function. Specificity in these pathways apparently depends on a complex set of interactions between the template and additional host-encoded proteins (15,39). These interactions presumably result in a higher-order modification of the template DNA molecule, forming either unique or multiple nucleoprotein complexes that can in turn serve as sites for dnaG action. These sites may also affect the nature of the enzymatic reaction catalyzed 25. 26. 27. 28. 29. 30. 31. 32. 33. 34. 35. 36. 37. 38. 39.

Wickner, S., and Hurwitz, J. (1974). PNAS 73, 4120-4214. Shlomai, J . , and Kornberg, A. (1979). PNAS 77, 799-803. Shlomai, J., and Kornberg, A. (1980). JBC 255, 6794-6798. Shlomai, J., and Kornberg, A. (1980). JEC 255, 6789-6793. Meyer, R. R., Glassberg, J . , and Kornberg, A. (1979). PNAS 70, 1702-1705. McMacken, R., Ueda, K., and Kornberg, A. (1977). PNAS 74, 4190-4194. Weiner, J . H., McMacken, R . , and Kornberg, A. (1976). PNAS 73, 752-756. Reha-Krantz, Hunvitz, J. (1978). JBC 253, 4043-4050. Reha-Krantz, Hurwitz, J. (1978). JBC 253, 4051-4057. Arai, K . , and Kornberg, A. (1979). PNAS 76, 4308-4312. Arai, K . , and Kornberg, A. (1981). JBC 256, 5267-5272. Arai, K . , Yasuda, S., and Kornberg, A. (1981). JBC 256, 5247-5252. Arai, K., and Kornberg, A. (1981). JEC 256, 5253-5259. Arai, K . , and Kornberg, A. (1981). JBC 256, 5260-5266. Sims, J., and Benz, E. W., Jr. (1980). PNAS 77, 900-904.

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E. W. BENZ, JR., D. REINBERG AND J. HURWITZ

by dnaG, such as turnover, length of primer formed, and preferred substrate (ribonucleotide versus deoxyribonucleotide). Priming replication with DBP-coated phage a3 DNA (or 4K, st- 1, or G4 DNAs) is an example of the formation of a single unique nucleoprotein complex (only one site per DNA molecule) (39). The protein requirements for this pathway are simple: Only DBP and dnaG protein are necessary. However, the DNA requirement is extensive and intricate. The formation of the nucleoprotein complex involves a rather large and specific region of DNA located at the origin of replication (40-42); two distant hairpin structures are brought close together in an ATP-independent reaction by DBP. Only in the presence of DBP can a single molecule ofdmG protein bind to both structures and synthesize a unique primer (39). By contrast, primer synthesis on DBP-coated 4x174 DNA requires many of the additional host-encoded proteins presumably required for initiations in discontinuous E. coli chromosome replication (7). Like the much simpler general pathway described above, the 4x174 pathway requires thednaB protein as well as the dnaG protein, and ultimately many different loci on the chromosome can support primer formation (30). Unlike the general pathway, DBP is required for proper priming and the formation of the nucleoprotein complex intermediate involves the participation of at least four additional proteins, replication factors X, Y, and Z, and the gene product of dnaC (see Fig. 1 and Table 11). The nature of the DNA structure required is not fully understood but appears to involve the ATP-dependent assembly of these proteins and drzu B protein at a specific structural site on the 4 X chromosome (called the replication factor Y site), possibly involving a large hairpin structure (25-28, 43). To reconcile the multiplicity of the transcripts with the assumption that the dnnG protein binds and initiates primer synthesis only where dnci B protein is present, it has been proposed that dnaB protein, after it is bound, migrates on the template (30). Movement of the complex of proteins is processive, appears to occur in the antielongation direction (5' -+ 3' direction of the template) (22, 4 3 ) , and can occur without necessarily forming primers. Hydrolysis of dATP (and probably ATP) by the n' protein (replication factor Y) is linked to the movement of the protein complex with the accompanying local displacement of DBP (22, 23,36-38,43). The special properties of this protein complex imply that, although there is a unique origin on the 4x174 chromosome for minus strand replication, this site may not necessarily be the site of primer formation. The mechanism of 40. 41. 42. 43.

Sims, J . , Capon, D., and Dressier, D. (1979). JBC 254, 12615-12628. Sims, J., Koths, K . , and Dressier, D. (1978). C S H S Q B 43, 349-365. Sims, J . , and Dressler, D. (1978). P N A S 75, 3094-3098. Arai, K . , and Kornberg, A . (1981). P N A S 78, 69-73.

163

6. PRIMING ENZYMES

*

+

DNA Binding protein

4dNTPs

FIG.1. Pathways ofdnuG-dependent priming on two bacteriophage DNAs. The roles of dnaG protein are described in Section 111. (*) In the absence of the elongation proteins, many primers are formed at many sites on the 6x174 DNA.

action of this mobile promoter during actual primer synthesis is unknown, but it may involve local displacement of DBP in the proximity of dnaB protein, and subsequent utilization of the general pathway mechanism for primer formation. The 4x174 SS to RF DNA replication pathway thus involves fiveE. coli

164

E. W.BENZ. JR., D. REINBERG, AND J. HURWITZ TABLE I1 PRIMING PROTEINS I N DIFFERENT SYSTEMS Protein

Replication factor X (protein i) Replication factor Y (protein n')

System

Gene

E . cnli

Replication factor Z (protein n+n") dncC protein

dnaC

dnc.B protein

dna B

ssb-1

DBP (SSB) dnaG protein (primase) DNA polymerase I11 elongation system (DNA polymerase I11 holoenzyme) DNA polymerase 111 (4

dnnG

Function Prepriming Prepriming ssDNA-dep- ATPase Prepriming Prepriming dnaB-dnaC complex DN A-dep-rNTPase prepriming priming mobile promoter Binding to ssDNA Primer synthesis DNA elongation

dna E

pole

Elongation factor I ( p ) Elongation factor 111 (6) dnaZ protein ( y ) 25 K protein (4 10 K protein (0) 80 K protein (7) T7 gene 4 protein (T7 primase)

dnaZ

T7

5 rrxA ( E . coli)

T7 DNA polymerase T4 gene 41 protein

4

T4

41

T4 gene 61 protein

61

T4 gene 32 protein (helix-destabilizing protein) T4 DNA polymerase T4 genes 44/62 proteins (polymerase accessory proteins)

32

T4 gene 45 protein (polymerase accessory proteins)

43 44/62 45

Processivity of elongation Processivity of elongation Processivity of elongation Rimer synthesis DNA-dep-NTPase unwinding DNA elongation Primer synthesis (+gene 61 protein) DN A-dep-NTPase fork movement? Primer synthesis (+gene 41 protein) fork movement? Binding to ssDNA DNA elongation Processivity of elongation DNA-dep-ATPase Processivity of elongation

6. PRIMING ENZYMES

165

coded proteins, replication factors X, Y , and Z, dnaC, and dnaB, which must act on the DBP-coated 4X chromosome, presumably forming a nucleoprotein complex that dnu G protein can recognize. The DNAdependent ATPase activity of replication factor Y is specific for the 4 X chromosome when compared to other single-stranded phage DNAs. Furthermore, replication factor Y DNA-dependent ATPase activity can be localized to a single region on the 4X chromosome. Using a combination of restriction enzyme digests and digestion with E. coli exonuclease VII, this activity can be supported by an exoVII-resistant fragment, 55 bases long, which resides within the 107-nucleotide untranslated region between phage coat protein genes F and G (26). The location of the minus strand origin of phage G4 (but not a3, +K, or st-1) also lies between genes F and G (40). This fragment has the potential of forming a single large hairpin that does not appear to have any special properties. There is little sequence homology between this intercistronic origin region and those of 4 K , a3, st-I, and G4. IV. Studies on the Specificity of dna G-Template Interactions

The three dnu G-dependent priming pathways previously described are clearly differentiated by their protein requirements (see Fig. 1). The differentiation of these pathways on the basis of template requirements is not so clearly defined. However, the template requirement plays a critical, interactive role in determining the specificity of priming. Since the general pathway (dnuB + dnuG in the absence of DBP) forms many primers on many different templates, the physical characterization of the template site for initiations is incomplete. The formation of families of primers of discrete size and class in this pathway does at least suggest some local template preference for initiation sites (34). What features these families of sites may have in common are unknown. The fact that even deoxyhomopolymers, such as poly(dT), can act as a template for this pathway adds to the puzzle and suggests little specificity. The second pathway, that previously described for phage DNAs from a 3 , 4 K , st-1, and G4, involves a simple protein requirement but a complex DNA template requirement. The origins of minus strand synthesis in these four phages have been located, sequenced, and their interactions with dnuG protein investigated (15.39-42,44). These studies have provided the following observations: (a) In all four phages the initiation site for minus

44. Fiddes, J. C . , Barrell, B. G . , and Godson, B. N . (1978). P N A S 75, 1081-1085.

166

E. W.BENZ, JR., D. REINBERG, AND J. HURWITZ

strand synthesis resides within an intercistronic region of approximately 135 bases of DNA. However, with phage G4 the origin occurs between genes coding for the viral coat proteins F and G, but with a3, st-1, and 4 K the origin is shifted to a position between coat protein genes G and H. (b) Extensive nucleotide conservation exists at the minus strand origins but does not extend into the adjacent coding regions. The conserved DNA consists of two regions, 42 and 45 bases long, that are separated by 13 bases of divergent sequence. (c) Correlated with the two regions of conserved sequence are two regions of potential hairpins (see Fig. 2). The start point of minus strand synthesis is located just upstream from the smaller of these two hairpins. Similarities in both primary sequence and secondary structure can be found in the general origin regions of bacteriophage lambda and E . coli (40). (d) Single-stranded DNA fragments containing these structures support the dna G binding reaction only when DBP is present in the reaction, and only if both hairpin structures are present on the same molecule (15, 39). Only the origin-containing strand of DNA that acts as a template for minus strand synthesis supports the dnaG reaction; its complement is totally inactive. (e) ThediiuG reaction is inactivated and reactivated by reagents or procedures that cause changes in the higher-order structure of the DNA without changing its primary structure (11). (f) The stoichiometry of these reactions is unchanged; that is, one molecule of dntrG protein forms one molecule of primer on one molecule of template. (g) Under these conditions the binding of dnaG to origin-containing DNA fragments is shown to protect nucleotides at the base of both hairpins; removing the larger, downstream hairpin from the fragment that contains the smaller, primer hairpin prevents binding to the primer hairpin (see Fig. 2). (h) The dnaG protein molecule is too small to span a linear distance of about 115 bases, the distance between protected regions of the DNA. From the sequencing, enzymological, and footprinting experiments, the conclusion was drawn that the structure the dnciG must require for proper binding to occur is a higher-order structure that has both hairpins approximated in space, presumably in a reaction stabilized by DBP. These studies suggest that structure, whether dictated by nucleotide sequence or resulting from protein interaction with the nucleotide chain is as important as the sequence per se. V.

Priming by RNA Polymerares

Like the systems described for the isometric phages a3, st-1, +K,and G4, and for 4x174, the single-stranded DNAs from the filamentous phages fd and MI3 can be replicated in vitro in both purified protein and

v; 'G 'C

vvvvvvv

GG C G G C

vvv

A

G

c

C C G

E F'.

v &Err

t

ttt

tttt

T T A A A T A A A A G C G A G C ~ A T A C G G A G A T A C C C G A T A A A C T A G GA A C G T G C C T C C T G C T A A G C C C A A A . A A G G 5' 4 4 3' I + , I 0

+so

*,o

+

+L

FIG.2. Secondary structure of the @K intercistronic region, showing the effects ofdfurG protein on the micrococcal nuclease cleavage pattern. Protections are indicated by downward-pointing arrowheads: upward-pointing arrows indicate enhancements. In all cases, the effect is on cleavage to the 3' side of the indicated nucleotide [from Ref. (39)l. The first nucleotide of the primer is indicated as + 1 .

168

E. W. BENZ, JR., D. REINBERG, AND J . HURWITZ

crude extract systems ( 5 , 45, 4 6 ) . Minus strand DNA synthesis on plus strand template DNAs begins at a unique origin on the chromosome (47). These reactions require all four ribonucleoside triphosphates as well as the four deoxyribonucleoside triphosphates, and are inhibited by rifampicin, an antibiotic that specifically inhibits E. coli DNA-dependent RNA polymerase. The products of these reactions are minus strands of nearly full genomic length in a duplex RFII structure that contains a gap in a specific region. These products are analogous to those described for the isometric phages (48, 49). In vitro reconstitution of the protein requirements show that in addition to RNA polymerase other proteins are required in the SS to RF reaction. The other proteins do not include the dnaG protein. As described for the isometric phages, elongation of the primer molecule proceeds via the DNA polymerase I11 elongation system pathway. A single oligoribonucleotide, about 30 bases long, is formed in a rifampicin-sensitive reaction that requires the combined action of DBP and RNA polymerase (50). This oligoribonucleotide is complementary to sequences located at the origin of replication, between genes I1 and IV in phage fd. The DNA requirement in this reaction has been studied by digesting fd DNA, complexed with DBP and RNA polymerase, with DNase I ( 4 9 ) . After the digestion reached completion, the resultant DNase I resistant fragment was isolated and analyzed. This fragment is about 120 bases long, contains sequences located between genes I1 and IV and the sequences used as template for the synthesis of the oligoribonucleotide primer molecule. Potentially, this fragment could form two large hairpin structures separated by a short (12 base) single-stranded stretch (51). Unlike thednaG site in phages +K, a3, st-1, and G4, only the hairpin that actually serves as template for primer formation is thought to be essential, since insertion of foreign DNA into the loop of the second hairpin does not inactivate origin function. However, binding studies have not been done to show that RNA polymerase binds to the same site. The origin region in these phages does not show significant sequence homologies with promoter regions for RNA polymerase (52). 45. Vicuna, R., Hunvitz, J . , Wallace, S . , and Girard, M. (1977). JBC 252, 2524-2533. 46. Vicuna, R . , Ikeda, J.-E., and Hurwitz, J. (1977). JBC 252, 2534-2544. 47. Tabak, H . F., Griffith, J., Geider, K . , Schaller, H., and Kornberg, A. (1974). JBC 249, 3049-3054. 48. Shaller, H . , Uhlmann, A., and Geider, K. (1976). PNAS 73, 49-53. 49. Gray, C. P., Sommer, R., Polke, C . , Beck, E., and Schaller, H. (1978). PNAS 75, 50-53. 50. Geider, K., Beck, E., and Schaller, H. (1978). PNAS 75, 645-649. 51. Schaller, H. (1978). CSHSQB 43, 401-408. 52. Schaller, H., Gray, C., and Hermann, K . (1975). PNAS 72, 737-741.

6. PRIMING ENZYMES

169

As is the case for the replication factor Y site of 4x174, the characteristics that make the fd origin DNA site unique are unknown. Primer synthesis by RNA polymerase begins with an adenine residue dictated by the single-stranded DNA region upstream of the primer hairpin. RNA polymerase then moves into the hairpin, presumably melting it and stopping at a point just short of the postulated loop. The signals that cause RNA polymerase to stop are no better understood than the signals that cause it to start. It is conceivable that destabilization of the duplex structure of the hairpin by RNA polymerase might allow DBP to bind to the now single-stranded downstream stem of the hairpin and block further translocation of RNA polymerase. There is little sequence homology between the fd origin DNA, the factor Y site DNA in $X, and the dnaG binding region at the origin of phages 4K,a3, st-1, and G4. As expected, extensive sequence homology exists among the origin regions of fd, M13, and fl phages, which are all related filamentous phages (53). Purified RNA polymerase can prime SS to RF DNA replication in 4x174 DNA as well as fd DNA; however, the predominant pathway for priming 4x174 DNA replication is rifampicin-resistant and involves dna G protein. Thus, additional proteins must be present in crude extracts that allow discrimination between the two pathways (45, 46). These proteins have been isolated and involve RNase H and two proteins called discrimination factors alpha and beta. The role of these proteins appears to be that of precluding priming by RNA polymerase on +X174 templates, presumably by degradative action. More detailed knowledge of the role of alpha and beta proteins is not yet known. VI.

Priming on Double-Stranded DNA

In the previous sections we described multiple pathways for priming on single-stranded DNA templates. The degree of complexity and specificity of each pathway is determined by a number of different enzymes that are necessary to deliver the dnaG protein or RNA polymerase to the origin or priming site. Presumably, the DNA sequence or structure of the origin region dictates the proteins needed. Irrespective of the pathway, after a primed template is formed elongation occurs by a common mechanism. The special requirements of duplex DNA replication present new problems for initiation. All of the initiation sites that have been studied occur only on single-stranded templates; thus, there is an apparent requirement for additional proteins to create single-stranded regions from duplex 53. Horiuchi, K . , Ravetch, J. V., and Zinder, N. D. (1978). CSHSQB 43, 389-399.

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DNAs. Furthermore, the antiparallel orientation of each strand in the duplex and the requirement that replication proceed in a 5’ + 3‘ direction on each template strand imply that multiple initiations must occur on the strand that grows from the 5’ end (lagging strand). The duplex must be unwound in order for replication to proceed on both strands, and the strain that accumulates during this unwinding must be relieved by some mechanism. I n vitw and in v i m data suggest that in some replicons both dnnG protein and RNA polymerase can play important roles in priming DNA synthesis on the same template. Thus, methods devised to study initiation of DNA synthesis of double-stranded templates must be able to resolve the two or more mechanisms that may be operating on different strands of the duplex. Studies on 4X174-RFI DNA replication ir? vitro, in which the requirements for the synthesis of each strand were resolved separately, have provided the clearest picture of duplex DNA replication. Leading strand synthesis (the replication of plus or viral strand DNA) requires the 4X gene A protein, the rep protein, DBP, and the DNA polymerase I11 elongation system (54-58). Net synthesis of viral (plus) strand DNA can be obtained in excess of input DNA (58,59).In the case of RF to R F replication with qbX DNA, the 3’-OH necessary for elongation by DNA polymerase I11 is provided by the action of a specific phage-encoded endonuclease, the 4XA protein, which remains covalently attached to the 5’ terminus of the cleaved DNA molecule (60, 61). This cleavage also relaxes the RFI structure thereby allowing unwinding to occur without the accumulation of strain. Unwinding of the duplex is catalyzed by the rep protein and requires the presence of DBP (62-64). The displaced plus (viral) strand then acts as template for minus strand synthesis as described above for S S to RF DNA replication (22, 43, 65). This reaction requires thednoB, C, and G proteins, DBP, and replication factors X, Y,and Z,as 54. Sumida-Yasumoto, C., Yudelevich, A., and Hurwitz, J. (1976). P N A S 73, 18871891. 5 5 . Sumida-Yasumoto, C., and Hurwitz, J. (1977). PNAS 74, 4195-4199. 56. Eisenberg, S., Scott, J., and Kornberg, A. (1976). PNAS 73, 3151-3155. 57. Scott, J. F., Eisenberg, S., Bertsch, L. L., and Kornberg, A. (1977). P N A S 74, 193- 197. 58. Eisenberg, S., Scott, J. F., and Kornberg, A. (1976). PNAS 73, 1594-1597. 59. Eisenberg, S., Griflith, J., and Kornberg, A. (1977). P N A S 74, 3198-3202. 60. Ikeda, J.-E., Yudelevich, A., and Hurwitz, J. (1976). P N A S 73, 2669-2673. 61. Eisenberg, S . , and Kornberg, A. (1979). JEC 254, 5328-5332. 62. Duguet, M., Yarranton, G., and Gefter, M. (1978). CSHSQB 43, 335-343. 63. Scott, J. F., and Kornberg, A. (1978). JBC 253, 3292-3297. 64. Yarranton, G . T., and Gefter, M.(1979). P N A S 76, 1658-1662. 65. Arai, K.-I., Arai, N., Shlomai, J., and Kornberg, A. P N A S 77, 3322-3326.

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well as the complete DNA pol I11 elongation system, and gives rise to RFII molecules. RFII DNA can be converted to RFI‘ molecules by the actions of DNA polymerase I and DNA ligase. The RFI’ structures are supercoiled by DNA gyrase to form RFI molecules (21). In order for minus strand synthesis to start on the displaced plus strand, DNA replication of the viral strand must proceed for a distance equivalent to at least 65% of the minus strand template; that is, until the replication factor Y site is encountered in a single-stranded form. After this is achieved interaction of the replication factor Y protein, dnriB protein, and d m G protein takes place at the origin of minus strand synthesis (22, 23, 66, 671, probably occurring in a sequence of reactions analogous to those depicted in Fig. 1. This assemblage of proteins consists of at least the replication factor Y protein, the dnuB protein, and the dnaG protein, and can move in an antielongation direction (5’ to 3’ with respect to the displaced strand) (22, 43); thus, this complex has been termed a mobile promoter for primer synthesis. The reconstitution of 4X174-RF to RF DNA replication system has been accomplished in Kornberg’s laboratory with purified proteins (23, 6-5-67). These results are confirmed by studies showing that extracts prepared from E. coli mutants temperature-sensitive in driaB, C , or G protein, when incubated at nonpermissive temperatures and supplemented with (bX-RFJ DNA and purified $JXgene A protein, will catalyze only the synthesis of viral (plus) strands. This product can be chased to RF molecules by adding purified wild-typednaB, C, or G proteins to their respective inactivated extracts (670). Similar experiments have been carried out using G6RFI DNA as template. The accumulation of plus strand single-stranded circles is observed at nonpermissive temperatures in extracts prepared from E. coli cells temperature-sensitive for the dncrG protein (strain NY73) but not in extracts prepared from dnci B and dncr C temperature-sensitive mutants. These results indicate that plus (leading) and minus (lagging) strand synthesis during 4X174-RF DNA replication have different requirements and that the mechanism of initiation on each template strand is different. Since the start of minus strand synthesis in 4X174-RF DNA replication is located some 2500 bases distant form the initiation of plus strand synthesis, substantial synthesis of plus strand DNA must occur before displacement of the parental replication factor Y site occurs (see Fig. 3). Prior to displacement, the replication factor Y site, residing within the intergenic region between genes F and G, is inactive in its duplex form. The conse66. Shlomai, J . . Polder, L., Arai, K . , and Kornberg, A. (1981). JEC 256, 5233-5238. 67. Arai, N . , Polder, L., Arai, K . , and Kornberg, A. (1981). JBC 256, 5239-5246. 67a. Reinberg, D., Zipursky, S . L., and Hurwitz, J . (1981). JBC. in press.

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F I G . 3. Pathways of 4X174-RFI DNA replication in crude extracts. The 4X174-RFI DNA molecule is recognized by the 4x174 A protein giving rise to the RFII-A protein complex; this intermediate can follow several pathways on the presence of other proteins as described in the text, Section VI. Panel (c) shows that in order for lagging-strand synthesis to start, the replication factor Y site must be exposed in a single-stranded form. Panel (d) shows that once the protein complex is formed at the replication factor Y site, the complex can migrate in an anti-elongation direction, forming primers that can be extended by the elongation proteins [panel (e)]. Panel (g) shows that when the phage proteins are present, the displaced viral plus strand is packaged into phage particles.

quence of this physical separation of the plus and minus strand origins for R F DNA replication, and the dependence of the minus strand DNA recognition sequence upon synthesis initiated at the plus strand site, is a marked asymmetry in plus and minus strand synthesis.

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Whereas there is asymmetric synthesis of each template strand during 4x174 RF to RF DNA replication, leading and lagging strand synthesis can occur nearly simultaneously in other replicons, such as the E. coli plasmid colicin E l (68). The initiation of leading strand (the L strand) ( 6 9 ) DNA synthesis during colE 1 DNA replication requires RNA polymerase, RNase H, E: coli DNA polymerase I and DNA gyrase. A unique RNA transcript located at the origin of replication is synthesized by RNA polymerase and elongated by DNA polymerase I for about 100 nucleotides (70). This unique transcript serves as primer for the DNA polymerase 111 elongation system. Lagging strand (the H strand) synthesis, however, requires the dnaG, dna B (7f), and probably dnaC proteins (unpublished observations). These requirements closely resemble those for 4x174 minus strand synthesis. Furthermore, colE1 and pBR322 DNAs have been found to contain two discrete chromosomal segments that, when denatured, support replication factor Y ATPase activity (72). These segments are located downstream from the origins of DNA replication. One is located on the L strand and the other further downstream on the H strand. The evidence, therefore, suggests that lagging strand synthesis of colEl plasmid DNA occurs by a mechanism similar to that already described for 4x174 minus strand synthesis. The mechanism by which leading-strand synthesis (after it is initiated) proceeds is less clear. The fact that a replication factor Y site exists on the H strand suggests that after elongation by DNA polymerase I for about 100 nucleotides, a signal is encountered that converts synthesis from a continuous to a discontinuous mode. Staudenbauer ef al. (71) reported that leading-strand synthesis requires the dnaB gene product but not the dnuG gene product. This data is difficult to reconcile with data that show that plasmid pBR322, into which the 4 X A gene protein cleavage site had been inserted (in both orientations), can be replicated under conditions identical to those required for 4Xl74-RF DNA replication (73).Furthermore, when these recombinant plasmids are replicated in extracts prepared from E. coli strains thermosensitive for genes dnaB, C, or G under nonpermissive conditions, the product is exclusively single-stranded circular DNA. This is true whether the orientation is such that the A cleav68. Tomizawa, J., Sakakibara, Y., and Kakefuda, T. (1974). P N A S 71, 2260-2264. 69. Tomizawa, J. (1975). Nrititre (London) 257, 253-254. 70. Itoh, T., and Tomizawa, J . 4 . (1980). P N A S 77, 2450-2454. 71. Staudenbauer, W. L., Scherzinger, E., and Lanka, E. (1979). Molec. Gen. Genet. 177, 113- 120. 72. Zipursky, S. L., and Marians, K. J. (1980). P N A S 77, 6521-6525. 73. Zipursky, S. L., Reinberg, D . , and Hurwitz, J . (1980). P N A S 77, 5182-5186.

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age site is on the H strand or on the L strand. When these same reactions are carried out under permissive conditions, that is when d n a B , C, and G are active, only R F molecules are formed. These data indicate that replication of either strand requires the actions of d n a B , C, and G proteins. These results agree with other in v i m experiments that use extracts from DNA ligase mutants, and indicate that synthesis of both strands proceeds discontinuously (68). Possibly the replication factor Y site present on the H strand acts as an assembly site for d n a B , C, and G proteins, which proceed to convert replication to a discontinuous mode (72). These observations could well reflect the operation of two mechanisms during synthesis of leading-strand DNA. Perhaps with colEl and related plasmids, initiation is catalyzed by RNA polymerase and is continuous in the sense that it does not involve multiple priming sites. Upon exposure of the replication factor Y site, DNA synthesis is converted to a d n a B , C, G-dependent discontinuous mode. Further studies with isolated H and L strands from colEl and related plasmids will help resolve the requirements for leading- and lagging-strand synthesis. VII.

Phage-Encoded Priming Enzymes

A. BACTERIOPHAGE T7 In vivo studies of phage T7 first suggested the participation of the T7 gene 4 protein during the initiation of T7 DNA synthesis. After shifting a T7 gene 4 temperature-sensitive mutant to nonpermissive conditions, electron microscopic studies showed that large single-stranded gaps were formed on one side of each growing replication fork (74, 75). This observation suggested that gene 4 functioned in the initiation of lagging-strand DNA synthesis. Analysis of DNA synthesized in vivo and in v i m in the absence of a functioning gene 4 protein supported this conclusion [for a review see Ref. (7)]. Gene 4 protein has been shown to participate in the synthesis of oligoribonucleotide primers on displaced single-stranded DNA that arises during synthesis on duplex T7 DNA (76-80). DNA synthesized on circular single-stranded templates in the presence of DBP, 74. Dressler, D . , Wolfson, J . , and Magazin, M. (1972). P N A S 69, 998-1002. 75. Wolfson, J., and Dressler, D. (1972). PNAS 69, 2682-2686. 76. Kolodner, R., Masamune, Y., LeClerc, J. E., and Richardson, C. C. (1978). JBC 253, 566-573. 77. Kolodner, R., and Richardson, C. C. (1978). JBC 253, 574-584. 78. Kolodner, R., and Richardson, C. C. (1977). P N A S 74, 1525-1529. 79. Romano, L., and Richardson, C. C. (1979). JBC 254, 10476-10482. 80. Romano, L., and Richardson, C. C. (1979). JBC 254, 10483-10489.

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ribonucleoside triphosphates, T7 DNA polymerase and gene 4 protein was also shown to contain 5’-terminal oligoribonucleotides (80). These reactions have been reconstituted in v i m with purified proteins (76-82). The reactions, representing lagging-strand synthesis, require the E. coli DBP, T7 DNA polymerase, the T7 gene 4 protein, and ribonucleoside triphosphates, as well as deoxynucleoside triphosphates. Oligoribonucleotide primers can be formed by gene 4 protein acting alone in the presence of ATP and CTP on single-stranded templates. These RNA primers are composed of 5’-pppApCpCpC/A-3’ (79, 80). A simple recognition sequence has been found and shown to consist of the sequence complementary to the primer oligonucleotide with the requirement that it must also include a 3‘ C , i.e., 3’-CTGGG/T-5’.This sequence occurs 13 times on the 4x174 chromosome; studies on the relative efficiency of site utilization on this chromosome suggest the possibility that gene 4 protein first binds randomly to the single-stranded DNA and then travels in a 5‘ to 3‘ (antielongation) direction until it finds the first recognition sequence, and then synthesizes the predominantly tetranucleotide primer molecule (83). Unlike the dnaG protein, the T7 gene 4 protein has a single-stranded DNA-dependent nucleoside triphosphatase activity that is coupled to DNA synthesis (78). The gene 4 hydrolysis of NTPs to NDPs and Pi in the presence of single4rande.d DNAs accompanies, and is necessary for, DNA synthesis on duplex T7 DNA catalyzed by the T7 DNA polymerase. It has been suggested that the energy derived from gene 4 hydrolysis of ATP can be used to unwind the duplex in advance of the replication fork. Under appropriate conditions, gene 4 protein and T7 DNA polymerase can form a complex that can be isolated by gel filtration chromatography. Both dncrG and T7 gene 4 proteins are monomeric polypeptides of approximately the same molecular weight (64,000 and 58,000, respectively). In contrast to priming by the dnaG protein on phage templates, gene 4 protein priming activity is not dependent on the presence of a singlestranded DNA binding protein. Moreover, whereas dna G protein synthesizes several families with different length primers of varying sequence, the known primer molecules synthesized by the gene 4 protein are homogeneous in length and sequence. Only ribonucleoside triphosphates serve as substrate for gene 4 protein, while dncrG can use deoxynucleoside triphosphates and adenosine diphosphate in addition to the ribonucleoside triphosphates. The site recognized by gene 4 protein is far simpler than 81. Scherzinger, E . , Lanka, E., and Hillenbrand, G . (1977). Nitclric Acid> Res. 4, 415 1-4163. 82. Hillenbrand, G . Morelli, G . , Lanka, E., and Scherzinger, E. (1978). CSHSQB 43, 449-453. 83. Tabor, S . , and Richardson, C. C . (1981). PNAS 78, 205-209.

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that of the known sites for dnaG action. Studies (84) suggest that the site utilized in vivu in the T7 system is very similar to that determined from in vitro studies. By mapping five initiation sites on newly replicated T7 DNA, the recognition site for RNA primer formation was inferred to be 3’-CTGGN-5’ or 3’-CTGTN-5’. B. BACTERIOPHAGE T4 Synthesis of linear duplex bacteriophage T4 DNA in vivo depends on the products of several T4 genes; these are the genes 32,41,43,44,45,61, and 62, (85). The elongation of bacteriophage T4 duplex DNA has been reconstituted in vitro with purified or partially purified proteins (86-93). The complete in vitru system consists of seven phage T4-encoded proteins, the products of genes 32, 41,43,44,45, 61, and 62. Gene 32 protein is a single-stranded DNA binding protein (86, 94). Gene 43 protein is the T4 DNA polymerase (95-97). The products of genes 44 and 62 copurify as a tight complex that has DNA-dependent ribo- and deoxyribo-ATPase activities (88. 89, 98). Gene 45 protein activates the ATPase activity of the 44/62 complex (88, 89, 98). Both the 44/62 complex proteins and the 41 protein catalyze a single-stranded DNA-dependent hydrolysis of nucleoside triphosphates to nucleoside diphosphates and inorganic phosphate; these hydrolyses can be differentially blocked by (y-S)ATP and (y-S)GTP, respectively. The five proteins, the products of genes 32, 43, 44, 45, and 62, catalyze an efficient strand displacement synthesis 84. Ogawa, T., and Okazaki, T. (1979). Nrrclcic Acids Res. 7, 1621-1633. 85. Epstein, R. H., Bolle, A,, Steinberg, C. M., Kellenberger, E., Boy d e la Tour, E., Chevalley, R., Edgar, R. S., Susman, M., Denhardt, G. H., and Leilausis, A. (1963). CSHSQB 28, 375. 86. Alberts, B., and Sternglanz, R. (1977). Natirre (London) 269, 655-661. 87. Liu, C.-C., Burke, R. L., Hibner, U., Barry, J., and Alberts, B. (1978). CSHSQB 43, 469-487. 88. Alberts, B. M., Morris, C. F., Mace, D. N., Bittner, M., Sinha, N. K., and Moran, L. (1975).1fi “DNASynthesisandIts Regulation”(M. Goulian, P.Hanawalt,andC. F. Fox,eds.), pp. 241-269. Academic Press, New York. 89. Morris, C. F., Sinha, N. K., and Alberts, B. M. (1975). PNAS 72, 4800-4804. 90. Alberts, B. M., Barry, J., Bittner, M., Davies, M., Hama-Inaba, H., Liu, C. C., Mace, D., Moran, L., Morris, C. F., Piperno, J., and Sinha, N. K. (1977). In “Nucleic Acid-Protein Recognition” (H.J. Vogel, ed.), pp. 15-29. Academic Press, New York. 91. Silver, L. L., and Nossal, N. G. (1978). CSHSQB 43, 489-494. 92. Nossal, N. G. (1979). JBC 254, 6026-6031. 93. Nossal, N. G., and Peterlin, B. M. (1979). JBC 254, 6032-6037. 94. Champoux, J. J. (1978). Annu. Rev. Biochem. 47, 449-479. 95. Huang, W. M., and Lehman, I. R. (1972). JBC 247, 3139-3146. 96. DeWaard, A,, Paul, A. V., and Lehman, I. R. (1965). PNAS 54, 1241-1248. 97. Goulian, M., Lucas, Z. J., and Kornberg, A. (1968). JBC 243, 627-638. 98. Barry, J . , and Alberts, B. M. (1972). PNAS 69, 2717-2721.

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(leading-strand synthesis) at a nick in duplex DNA (87, 91). This synthesis is stimulated by the addition of genes 41 and 61 proteins, especially when T4 DNA is used as template (87). These protein components have been grouped into functional classes as follows: (1) The T4 DNA polymerase (gene 43 protein); (2) the polymerase accessory proteins (genes 44/62 complex and 45); (3) the helixdestabilizing protein, a single-stranded DNA binding protein (gene 32); and (4) the RNA priming proteins (genes 41 and 61). All seven proteins described above are required for lagging-strand synthesis (87, 91). Studies have shown that genes 41 and 61 proteins are required for priming synthesis on single-stranded DNAs. The priming activities of the T4 replication proteins have been determined by using single-stranded 4x174, fd and fl DNAs as model templates. The T4 gene 41 protein and a protein controlled by the T4 gene 61, when present together in reactions containing 4x174 DNA (or fd DNA), form pentaribonucleotides with the sequence pppApCpNpNpN, where N can be G, U, C, or A (99, 100). The same group of sequences was found in the RNA at the 5’ terminus of the 4x174 DNA product made by the combined actions of the T4 gene 32, 41, 43 (DNA polymerase), 44/62 complex, 45, and 61 proteins (99). The T4 gene 61 protein was identified as an activity (absent in extracts of E. coli infected with T4 gene 61-deficient mutants) that is required for the formation of oligoribonucleotides and for DNA synthesis with unprimed single-stranded circular templates (99). The activity associated with this protein has been extensively purified by successive chromatography on single-stranded DNA cellulose, phosphocellulose, hydroxylapatite, and valine-Sepharose columns. The apparent molecular weight of the gene 61 protein is 40,000 to 45,000; it is active as a monomer polypeptide that is highly basic (101). In v i m both genes 41 and 61 proteins must be present simultaneously in order for primer synthesis to occur. It is not known whether the genes 41 and 61 proteins will catalyze a pyrophosphate exchange or pyrophosphorylysis reaction. Even though the primers formed are short, primer utilization is efficient; nearly 90% of the primers formed in Iitro are used to prime DNA synthesis on single-stranded circular templates (fd) (100). As with the T7 gene 4 product, the genes 41 and 61 proteins appear to act at multiple, but specific, sites on single-stranded 99. Nossal, N . G . (1980). JBC 255. 2176-2182. 100. Liu, C.-C., and Alberts, B. M. (1980). P N A S 77, 5698-5702. 101. Alberts, B . M., Barry, J . , Bedinger, P., Burke, R . L . , Hibner, U . , Liu, C.-C., Sheridan, R. (1980). 111 “ICN-UCLA Winter Symposium on Mechanistic Studies on DNA Replication and Recombination’’ (B. Alberts.ed.), in press. Academic Press, New York, pp. 449-473.

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DNAs (99-101). The stoichiometry of gene 41 protein to gene 61 protein to template required to form a single primer is not yet known. The T4 gene 41 product appears similar to the E. coli dnaB gene product (that was resolved in the (bX174 system) by virtue of their common single-stranded DNA-dependent nucleoside triphosphatase activities. The T7 gene 4 protein itself has single-stranded DNA-dependent nucleoside triphosphatase activity and is larger than the T4 gene 61 protein (58,000 compared to 40,000-45,000). Both are smaller than the E. coli dnoG gene product, which is 64,000 and has no DNA-dependent triphosphatase activity. Unlike primer site recognition by dnaG protein in a3, st-1, 4K,and G4, and for 4x174, all of which require the action of a single-stranded DNA binding protein (DBP), neither the TCencoded nor the T7-encoded priming proteins require the action of a single-stranded DBP for in vituo priming activity on single-stranded templates. The primer sites recognized by the phage-encoded priming proteins may therefore differ from those recognized by the E. coli proteins, at least in two of the three pathways described above. VIII.

Priming in Eukaryotic Systems

The molecular mechanisms involved in priming eukaryote DNA replication are not well understood. The major obstacle has been the lack of a cell-free system capable of faithfully duplicating the in vivo replication process. Studies with DNAs and proteins from eukaryotic organisms have also been limited by the difficulty of obtaining genetic mutants that affect DNA replication and by the high molecular weight of the DNA, making it difficult to isolate and manipulate. Studies of SV40, polyoma, herpes virus, adenovirus, and yeast DNA replication with either cell extracts or partially purified proteins have offered hope that these limitations can be overcome (102-104). As with the bacteriophages, the smaller size of these viral DNAs means easier isolation and manipulation, as well as dependence on host enzymes for at least some of the functions required for DNA replication. SV40 DNA replication differs from the asymmetrical rolling circle model described for some bacteriophage DNAs. SV40 molecules replicate semiconservatively (105, 106) with the majority beginning replication at a 102. 103. 104. 105.

Kelly, T. J . , and Nathans, D. (1977). Adbwn. Virus. R e s . 21, 86. Winnacker, E. L. (1978). r e / / 14, 761. Kornberg, A. (1980). “DNA Synthesis.” Freeman, San Francisco, California. Hirt, B. (1968). P N A S 55, 997. 106. Magnusson, G . , Winnacker, E. L., Eliasson, E., and Reichard, P. (1972). J M B 72, 539.

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unique origin and proceeding in both directions at approximately equal rates (107-109). Although replication begins at a unique origin in every eukaryote and prokaryote system thus far investigated, replication on both strands is not always synchronized in both directions (110-112). SV40 and polyoma DNA elongation process is predominantly a continuous process in the same direction as fork movement on the leading strand, and a discontinuous process on the lagging strand, which grows in the opposite direction of fork movement (1 13-116). Discontinuous DNA replication occurs through repeated initiations of short pieces of nascent DNA (130-150 bases long), called Okazaki fragments (117-119). Initiation of Okazaki fragments appears to involve an RNA primer (about 10 bases long) that is covalently attached to the 5' terminus of the nascent DNA fragment (120-125). A similar observation has been reported for mammalian cells (126-130). SV40 has two genetic loci (A and D) that affect initiation of DNA replication. Mutants in the D gene that codes for a coat protein, do not uncoat properly and therefore prevent transcription of early genes that are 107. Danna, K.J., and Nathans, D. (1972).P N A S 69, 3097. 108. Fareed, G. C., and Salzman, N. P. (1972).Nrrttrra N e w Biol. 238, 274. 109. Thoren, M. M . , Sebring, E. D., and Salzman, N. P. (1972).J . Virol. 10, 462. 110. Chattoraj, D., and Inman, R. (1973).P N A S 70, 1768. 1 1 1. Bogenhagen, D. A , , Gillum, A., Martens, P., and Clayton, D. (1978).C S H S Q B 43, 253-262, 112. Lechner, R., and Kelly, T. (1977).Cell 12. 1007. 113. Flory, P. (1977).Ntrcleic Acids R1.s. 4, 1449. 114. Hunter, T., Francke, B., and Bacheler, L. (1977).Cell 12, 1021. 115. Perlman, D.,and Huberman, J. (1977).Cell 12, 1029. 116. Kaufman, G..Bar-Shavit, R., and DePamphilis, M. L. (1978).Nltrleic Acids R P S . 5, 2535. 117.*Fareed, G . , and Salzman, N. L. (1972).Nitrttro N i w Eiol. 238, 274. 118. Pigiet, J., Winnacker, E. L., Eliasson, R., and Reichard, P. (1973).Nutrrre N e w Biol. 245, 243. 119. Francke, B.,and Hunter, T. (1974).J M E 83, 99. 120. Hunter, T., and Francke, B. (1974).JMB 83, 123. 121. Pigiet, J., Eliasson, R., and Reichard, P. (1974).J M B 84, 197. 122. Reichard, P., Eliasson, R., and Soderman, G. (1974).P N A S 71, 4901. 123. Anderson, S . , Kaufmann, G., and DePamphilis, M. L. (1977).Biorl7emistry 16, 4990. 124. Eliasson, R., and Reichard, P. (1978).JBC 253, 7469. 125. Eliasson, R.,and Reichard, P. (1979).J M B 129, 393. 126. Edenberg, H., and Huberman, J. (1975).Auutt. Re\.. Getzer. 9, 245. 127. Tseng, B.,and Goulian, M. (1975).JMB 99, 329. 128. Waqar, M., and Huberman, J . (1975).Cell 6, 551. 129. Tseng, B.,and Goulian, M. (1977).Cell 12, 483. 130. Tseng, B., Erickson, J., and Goulian, M. (1979).J M B 129, 531.

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required for DNA replication (131, 132). Gene A codes for two proteinslarge T antigen and small T antigen (133). Studies of thermolabile gene A mutants (134-139) and microinjection of purified T antigen into cell nuclei (140, 141) have shown that only large T antigen satisfies the requirement for initiation of SV40 viral DNA replication and stimulation of host cellular DNA synthesis. T antigen is required for initiation of DNA replication but not for completing elongation of viral DNA (136). T antigen binds preferentially to the 120 nucleotides defining the origin of SV40 DNA replication (142) and is essential for proper initiation of DNA synthesis (143). However, no priming activity associated with T antigen has been demonstrated. Since a DNA-dependent ATPase and a gyrase activity copurifies with T antigen (144, 1 4 9 , additional mechanisms other than direct interaction with the DNA may be involved in initiation. At present, the only eukaryotic system that can initiate nascent DNA chains in v i m upon addition of exogenous DNA and partially purified proteins is the adenovirus DNA system (146, 147). In the adenovirus system, both DNA strands of the duplex isolated from virions are linked to a 55,000 dalton protein (148). The covalent linkage is between dCMP at the 5' terminus of the DNA and the protein (149). DNA-free crude fractions isolated from adenovirus infected HeLa cells catalyze continuous asymmetric semiconservative replication of exogenous adenovirus DNA. This reaction requires the addition of adenovirus DNA that contains the protein attached to the 5' termini. Reagents or reactions that alter the 55,000 dalton protein, such as phenol, heat, or proteinase K, inactivate the viral DNA as template; all adenovirus DNA synthesis carried out in 131. Shenk, T., Rhodes, C., Rigby, P., and Berg, P. (1975). PNAS 72, 989. 132. Chou, J . , and Martin, R. (1975). J . Virol. 15, 145. 133. Crawford, L., Cole, C., Smith, A . , Pancha, E . , Tegtmeyer, P., Rundell, K., and Berg, P. (1978). PNAS 75, 117. 134. Yamuguchi, N . , and Kuchino, T. (1975). J . Virol. 15, 1279. 135. Osborn, M., and Weber, K. (1975). J . Virol. 15, 636. 136. Tegtmeyer, P. (1975). J . Virol. 15, 613. 137. Brugge, J., and Butel, J. (1975). J . Virol. 15, 619. 138. Martin, R., and Chou, J. (1975). J . Virol. IS, 599. 139. Kimura, G . , and Itagaki, A. (1975). PNAS 72, 673-677. 140. Graessmann, M., and Graessmann, A. (1976). PNAS 73, 366. 141. Tjian, R., Fey, G . , and Graessmann, A. (1978). PNAS 75, 1279. 142. Tjian, R. (1978). Cell 13, 165. 143. Shortle, D., and Nathans, D. (197'9). J M E 131, 801-817. 144. Giacherio, D., and Hager, L.P. (1979).JEC 254, 8113-8116. 145. Giacherio, D., and Hager, L. P. (1980). JBC 255, 8963-8966. 146. Challberg, M. D., and Kelly, T. J. (1979). PNAS 76, 655-659. 147. Kaplan, L. M . , Ariga, H., Hunvitz, J . , and Horwitz, M. S . (1979). PNAS 76, 5534-5538. 148. Rekosh, R. M. K . , Russell, W. C . Bellet, A. J . D., and Robinson, A. J. (1977). Cel/ 11, 283-295. 149. Carusi, E. A. (1977). Virology 76, 380-394.

6. PRIMING E N Z Y M E S

181

vitro is covalently linked to protein (150-153). Interestingly, the protein

linked to newly synthesized DNA is predominantly an 80,000 dalton protein; both the 55,000 and the 80,000 dalton proteins have tryptic oligopeptides in common, suggesting a precursor-product relationship (1.50). The requirements for adenoviral DNA replication have been examined with crude extracts from adenovirus infected cells. Cytosol extracts isolated from adenovirus-infected cells require the four deoxyribonucleoside triphosphates, Mg2+,ATP, dithiothreitol, adenovirus DNA-protein complex, and a nuclear extract fraction from uninfected cells. The nuclear extract can be replaced by purified DNA polymerase P preparations (154). These additions lead to the semiconservative synthesis of full genomic length adenovirus DNA. All synthesis is blocked by the potent DNA polymerase Q inhibitor, aphidicolin. Challberg et 01. (150) have shown that all DNA synthesis occurs with newly synthesized DNA covalently linked to an 80,000 dalton protein. In addition, Lichy et al. (155) have shown that [a-32P]dCTPcan be covalently linked to the 80,000 dalton protein independent of DNA synthesis. This reaction requires the adenovirus DNA-protein complex, ATP, nuclear extract from uninfected cells, and cytosol from adenovirus-infected cells. The linkage reaction is unaffected by aphidicolin. In the presence of dCTP, dATP, dTTP, and dideoxy-GTP, a 26-base-long oligonucleotideprotein complex (of 88,000 daltons) is produced and can be isolated. Micrococcal nuclease digestion of this complex converts it to the 80,000 dalton complex. The oligonucleotide component of the 88,000 dalton complex, like the 5’ ends of adenovirus DNA, contains no G residues in the first 25 bases. These results show that initiation occurs specifically on a protein, which leads to the covalent linkage of a 5‘-dCMP moiety to a P-hydroxyl site on the 80,000 dalton protein. Stillman ef al. demonstrated that this adenovirus terminal protein is coded for by the viral genome (156).

The crucial role played by the terminal protein moiety of adenovirus DNA in replication was demonstrated by Horwitz and Ariga, who showed that only the restriction digest adenovirus fragments that contained the terminal protein supported DNA replication (153). 150. Challberg, M . D., Desiderio, S . V., and Kelly, T. J . , Jr. (1980). PNAS 77, 51055109. 151. Ikeda, J . - I . , Enomoto, T., and Hurwitz, J. (1981). PNAS 78, 884-888. 152. Reiter, T. R . , Futterer, R. Weingartner, B., and Winnacker, E. L. (1980). J . Virol. 35, 662-671. 153. Horwitz, M . S . , and Ariga, H. (1981). PNAS 78, 1476-1480. 154. Ikeda, J.-E., Longiaru, M., Horwitz, M. S . , and Hurwitz, J. (1980). PNAS 77, 5827-5831. 155. Lichy, J., Horwitz, M. S. , and Hurwitz, J. (1981). /“AS 78, 2678-2682. 156. Stillmann, B. W., Lewis, J. B . , Chow, L. T., Mathews, M . B . , and Smart, J. E. (1981). Cell 23, 497-508.

182

E . W. BENZ, JR., D. REINBERG, A N D J. HURWITZ

The system that replicates adenovirus DNA has been resolved into multiple protein fractions, which have been partially characterized (151); DNA synthesis in the presence of the adenovirus DNA-protein complex requires the adenovirus DNA binding protein, DNA polymerase a and p and a partially purified protein fraction isolated from the cytosol of adenovirus-infected cells. This fraction contains detectable DNA polymerase a and DNA-dependent (and independent) ATPase, and catalyzes formation of the 80,000 dalton protein-dCMP complex independently of DNA replication. The fraction contains at least three readily discernible polypeptides. All of the DNA molecules synthesized in this reaction are covalently linked to an 80,000 dalton protein, as observed for the crude extract system (150). The requirements for adenoviral DNA replication with partially purified proteins include: The four deoxynucleoside triphosphates, Mg2+, ATP, dithiothreitol, adenovirus DNA-protein complex, adenovirus DNA binding protein, DNA polymerase p, and partially purified protein preparations isolated from the cytosol of adenovirus-infected cells. The latter fraction contains DNA polymerase a. The importance of this polymerase in adenovirus DNA synthesis is supported by the finding that aphidicolin is a potent inhibitor of the reaction with partially purified proteins and with crude extracts. The size of the product formed with the partially purified protein system varies between 25-50% the size of the product formed in the crude extract system (151). Synthesis of full genomic length adenovirus DNA requires the addition of cytosol from uninfected cells (1.50). Unlike the bacterial systems, the elongation of primed templates in the eukaryotic systems does not appear to follow a common pathway. With RNA as primer, only DNA polymerase a can extend the primer end (157, 158). Chain growth due to this enzyme rarely exceeds 200-300 bases, and requires the participation of DNA polymerase p to complete full genome length products (154). Since multiple forms of both polymerases have been reported (159-161), additional factors may be involved. The role played by DNA polymerase y is completely unknown. ACKNOWLEDGMENTS We wish to thank Drs. J. Ikeda and K. J . Marians for helpful advice and Dr. A. Kornberg and co-workers for communicating results prior to publication. 157. Weissbach, A . (1977). Annu. R e v . Biochtvn. 46, 25-47. 158. Bollum, F. J. (1960). JBC 235, 2399-2403. 159. Lamothe, P., Baril, B., Chi, A . , Lee, L., and B a d , E. (1981). PNAS 78,4723-4727. 160. Grummt, F., Waltle, G . , Jantzen, H. M., Hamprecht, R., Huebscher, D., and Kuentzle, C . C. (1979). PNAS 76, 6081-6085. 161. Rapaport, E . , Zamecnik, P. C., and Baril, E. F. (1981). PNAS 78, 838-842.

tRNA Nucleotidyltransferase MURRAY P. DEUTSCHER

I. Introduction . . . . . . . . . . . . . . . . 11. Purification and Structural Studies . . . . . . A.Assay . . . . . . . . . . . . . . . . . B. Purification . . . . . . . . . . . . , . . C . Purity and Specific Activity . . . . . . . D. Physical and Chemical Properties . . . . . 111. Catalytic Properties . . . . . . . . . . . . . A. pH and Cation Effects . . . . . . . . . . B. Nucleoside Triphosphate Donors . . . . . C. Kinetic Mechanism . . . . . . . . . . . D. RNA Acceptors . . . . . . . . . . . . . E. The Active Site and Synthesis of the -C-C-A F. Misincorporation of Nucleotides . . . . . G. Pyrophosphorolysis and Hydrolysis of tRNA IV. Biological Role . . . , , . . . . . . . . . . V. Research Applications . . . . . . . . . . .

.

.

. . . . . .

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

.

,

. .

. . . . . . , ,

. . . . , .

. . . . . . . . . . .

. . . . . Sequence. . . . , . . . . . . . . . . . . . . . . . . . , . . . . , . . , , , . . . . . . . . . .

183 186 186 186 188 190 192 193 196 198 199 207 210 211 213 215

I. Introduction

ATP(CTP)-tRNA nucleotidyltransferase ( I ) catalyzes the incorporation of AMP and CMP residues into tRNA molecules from which all, or part, of the 3' terminal trinucleotide sequence, -C-C-A, has been removed (2, 1. This enzyme (EC 2.7.7.25) has also been called -C-C-A pyrophosphorylase, tRNA adenylyltransferase and tRNA cytidylyltransferase. Under the latter name it has also been given a separate listing by the Commission on Biochemical Nomenclature (EC 2.7.7.21). For this article the name tRNA nucleotidyltransferase is used. 2. Deutscher, M. P. (1973). Progr. Nircleic Acid R e g . Mol. B i d . 13, 51.

183 THE ENZYMES, VOL XV

a

Copyright 1982 by Academic Prebs. Inc. All rlghts of reproduction in any form reserved ISBN 0-1 2- I227 15-4

184

MURRAY P. DEUTSCHER

3). In the presence of both ATP and CTP the enzyme will regenerate the -C-C-A sequence according to the overall reaction (4) tRNA-N

+ ATP + 2 CTP + tRNA-C-C-A +

3 PP,

(1)

In the presence of a single nucleoside triphosphate the enzyme will also catalyze the following partial reactions (2-4):

+ CTP tRNA-C + C T P +

tRNA-N

tRNA-C-C

4

+ ATP

4

tRNA-C or tRNA-C-C tRNA-C-C

+ PPi

tRNA-C-C-A

+ PP,

+ PP,

(2) (3) (4)

Reactions (2), (3), and (4) are generally used to assay tRNA nucleotidyltran sferase during purification. tRNA nucleotidyltransferase activity has been identified in a wide variety of organisms including mammals (5-7), birds ( 8 ) , amphibians ( 9 ) , insects (10),fungi ( I f , 12), higher plants (13, 14) and bacteria (15, 16). In addition, the enzyme has been shown to be present in mammalian mitochondria (17, 181, and to be associated with a variety of RNA tumor viruses (19, 20) and Sendai virus (21). The widespread occurrence of 3. Sprinzl, M., and Cramer, F. (1979). Progr. Niicleic Acid R e s . Mol. Biol. 22, 1. 4. Abbreviations: tRNA-N, tRNA lacking the entire pCpCpA sequence; tRNA-C, tRNA lacking the terminal pCpA sequence; tRNA-C-C, tRNA lacking the terminal AMP residue; PP,, inorganic pyrophosphate. 5. Canellakis, E . S . (1957). BBA 25, 217. 6. Herbert, E. (1958). JBC 231, 975. 7. Hecht, L. I., Zamecnik, P. C., Stephenson, M. L., and Scott, J. F. (1958). JBC 233, 954. 8. Chung, C. W., Mahler, H. R., and Enrione, M. (1960). JBC 235, 1448. 9. Paradiso, P., and Schofield, P. (1976). Exp. Cell R e s . 100, 9. 10. Poblete, P., Jedlicky, E., and Litvak, S. (1977). BBA 476, 333. 1 1 . Morris, R. W., and Herbert, E. (1970). Biochemistry 9, 4819. 12. Hill, R., and Nazario, M. (1973). Biochemistry 12, 482. 13. Cudny, H., Pietrzak, M., and Bartkowiak, S . (1975). Phytorhemistry 14, 85. 14. Dullin, P., Fabisz-Kijowska, A., and Walerych, W. (1975). Acta Biochirn. Pol. 22, 279. 15. Moldave, K. (1960). BBA 43, 188. 16. Coutsogeorgopoulus, C. (1960). BBA 44, 189. 17. Mukerji, S. K . , and Deutscher, M. P. (1972). JBC 247, 481. 18. Bouhnik, J . , Michel, O., and Michel, R. (1973). Biocliimie 55, 1179. 19. Faras, A. J., Levinson, W. E., Bishop, J. M., and Goodman, H. M. (1974). V i r d o g y 58, 126. 20. Thomassen, M. J., Kinsley-Lechner, E., Ohe, K., and Wu, A. M. (1976). BBRC 72, 258. 21. Kolakafsky, D. (1972). J . Virol. 10, 555.

7. tRNA NUCLEOTIDYLTRANSFERASE

185

tRNA nucleotidyltransferase in all types of cells, subcellular organelles, and viruses suggests a significant function for this enzyme that has been conserved during evolution. Activities that catalyze the terminal addition of nucleotides to tRNA have been known since the earliest studies of tRNA structure and function in the late 1950s (5-7, 22-24); in fact, this reaction complicated many of the early studies of RNA synthesis in vivo because it masked de novo incorporation of nucleotides. In detailed studies of the terminal addition reaction in mammalian cell extracts, Hecht et uf. ( 7 , 2 5 ) showed that CTP and ATP were the precursors for nucleotide incorporation, and that CMP and AMP were added in a 2 : 1 ratio. The reaction required Mg2+,and was inhibited by PPi, which reversed the reaction. The first partial purification of the enzyme involved in terminal addition of nucleotides to tRNA was reported by Canellakis and Herbert in 1960 (26, 27). During the next several years the enzyme was purified several hundredfold fromEscherichiu coli ( 2 8 , 2 9 ) ,rat liver (30),and rabbit muscle (31). Specific activities of these preparations were in the range of 1 to 6 pmol of AMP incorporated per hr per mg of protein. Each of these preparations also incorporated CMP in a constant ratio throughout purification, suggesting the involvement of a single enzyme for both activities. However, none of these enzyme preparations was characterized with regard to purity or structural properties of the protein. Further studies of tRNA nucleotidyltransferase lagged until the 1970s when a number of purified preparations of the enzyme became available from a variety of sources (10-12, 32-41). Since then there has been increased interest in 22. Heidelberger, C., Harbers, E . , Liebman, K. C., Takagi, Y., and Potter, V. R. (1956). BBA 20,445. 23. Paterson, A. R. P., and LePage, G . A. (1957). Cancer Res. 17, 409. 24. Edmonds, M., and Abrams, R. (1957). BBA 26, 226. 25. Hecht, L. I., Stephenson, M. L . , and Zamecnik, P. C. (1959). PNAS 45, 505. 26. Canellakis, E. S., and Herbert, E. (1960). PNAS 46, 170. 27. Canellakis, E. S., and Herbert, E. (1960). BBA 45, 133. 28. Furth, J. J., Hurwitz, J. Krug, R . , and Alexander, M. (19611. JBC 236, 3317. 29. Preiss, J., Dieckmann, M., and Berg, P. (1961). JBC 236, 1748. 30. Daniel, V., and Littauer, U . 2. (1963). JBC 238, 2102. 31. Starr, J . L., and Goldthwait, D. A. (1963). JBC 238, 682. 32. Gross, H. J . , Duerinck, F. R . , and Fiers, W. C. (1970). EJB 17, 116. 33. Carre, D. S., Litvak, S . , and Chapeville, F. (1970). BBA 224, 371. 34. Sternbach, H . , von der Haar, F., Schlimme, E . , Gaertner, E., and Cramer, F. (1971). EJB 22, 166. 35. Miller, J. P., and Phillips, G . R. (1971). JBC 246, 1274. 36. Best, A. N . , and Novelli, G. D. (1971). A B B 142, 527. 37. Deutscher, M . P. (1972). JBC 247, 450. 38. Rether, B., Bonnet, J., and Ebel, J-P. (1974). EJB 50, 281.

186

MURRAY P. DEUTSCHER

tRNA nucleotidyltransferase because of its role in processing and repair of tRNA, its study as a model enzyme for phosphodiester bond synthesis, and its usefulness as a reagent for modifying or labeling the 3‘ terminus of tRNA. 11.

Purification and Structural Studies

A.

ASSAY

tRNA nucleotidyltransferase activity is measured by incorporation of radioactive ATP or CTP into an acid-insoluble form (42). tRNA molecules lacking one, two, or three terminal residues may be used as acceptors depending on whether ATP or CTP is the donor. The quality of the tRNA acceptor is important because tRNA-N and tRNA-C can inhibit AMP incorporation into tRNA-C-C, and likewise, tRNA-C-C can inhibit CMP incorporation. Masiakowski and Deutscher (43, 44) determined tRNA nucleotidyltransferase activity by measurement of the release of [ 32P]PPi from [Y-~~PIATP or [ P,y3’P]CTP upon nucleotide incorporation into model acceptor substrates that were acid soluble. Comparison of this assay with the incorporation assay indicated that the two methods gave equivalent results, confirming the stoichiometry of reactions shown in Eqs. (3) and (4).

B. PURIFICATION A number of highly purified tRNA nucleotidyltransferase preparations from a variety of sources (Table I) have become available ( I 0 , 3 3 - 4 / , 4 5 , 46). Since tRNA nucleotidyltransferase is present in cells in extremely small amounts, purifications of 5000- to 25,000-fold are often required (Table I). In fact, it has been estimated that theE. coli enzyme is present in an abundance of only 140 molecules per cell (39). Generally, standard purification procedures have been used to prepare these enzymes, although some newer methods are worthy of note. Affinity chromatography 39. 40. 473. 41. 42. 43. 44. 45.

Schofield, P., and Williams, K . R . (1977). JBC 252, 5584. Leineweber, M., and Philipps, G. R. (1978). Hoppe-Seylev’s 2. Physiol. Chem. 359,

Cudny, H . , Pietrzak, M., and Kaczkowski, J. (1978). Plrntcr 142, 23. Deutscher, M. P. (1974). I n ‘*Methodsin Enzymology,” Vol. 29, p. 706, 1974. Masiakowski, P., and Deutscher, M. P. (1977). FEBS Lett. 77, 261. Masiakowski, P., and Deutscher, M. P. (1980). JBC 255, 11233. McGann, R. G., and Deutscher, M. P. (1980). EJB 106, 321. 46. Deutscher, M. P., and Masiakowski, P. (1978). Nucleic Acids Res. 5, 1947.

TABLE I PROPERTIES O F

Approximate purification" Source 1. E. coli MRE600

2. E . coli B 3. E . coli B 4. E . coli B

(-fold)

HIGHLY P U R I F I E D tRNA

Specific activityb (FmoUhr/rng)

0.2 60 29 36 4,000

5 . E . coli MRE600 6. E. coli A19 7. L. ucidophilus

11. Houseflies

12. Rabbit liver

Molecular weight

37,Oooc -

45,000' 54,000d

Sedimentation rate

(S)

Isoelectric point

2.9 3.4

6.4

-

-

1 ,000 200 244 500

-

(36) (39)

7.5

4.2

40,000'

-

30,000' 31 ,Wd 44,000' 47,000d 48,000'

2.6

' The symbol (H) after the purification indicates that the preparation was reported

Reference

-

5.9

53,000' 51,5Wd 45,000d 50,O0Od 43,000d 70,0OOd 71.000'

8. Bakers' yeast 9. Bakers' yeast 10. L. luteus seeds

NUCLEOTIDYLTRANSFERASES

3.6-4.0

7.2,7.5

to be homogeneous. Specific activities are based on the standard assays used in each preparation, and have been converted to common units. Sephadex. Sodium dodecyl sulfate-acrylamide gel electrophoresis. Equilibrium ultracentrifugation.

(37. 4 6 )

188

MURRAY P. DEUTSCHER

on tRNA-Sepharose has been used for preparation of the E. coli B protein (39) (Table 1,4), and the usefulness of this method has been confirmed for preparation of the enzyme from another strain of E. coli (45) (Table 1,6). Affinity chromatography on commercially available Affi-Gel Blue with selective elution by tRNA was successful for rapid purification of the rabbit liver enzyme (46) (Table 1,lZ). However, this adsorbent was not useful for the E. coli tRNA nucleotidyltransferase because the protein bound so tightly that it could not be eluted in satisfactory yield. The E. coli and L . acidopliilirs enzymes (Table I,5,7) have also been purified in good yield to apparent homogeneity in just two steps by using preparative polyacrylamide gel electrophoresis (40). This method may prove extremely useful for rapid purification of the enzyme. In all cases examined, the highly purified tRNA nucleotidyltransferases incorporate both AMP and CMP into tRNA. In addition, the ratio of the two activities remains constant over several thousandfold purifications (3.5, 38, 40, 45, 47). Both activities also comigrate during acrylamide gel electrophoresis (35,47)and isoelectric focusing (47). These results lead to the conclusion that a single protein catalyzes both AMP and CMP incorporation. This conclusion is supported by studies of E. coli mutants deficient in tRNA nucleotidyltransferase (48), since a single point mutation can equally affect both activities. Several workers have reported on the presence of multiple forms of tRNA nucleotidyltransferase (26, 29, 32, 37, 38, 40), in some cases with different enzymatic properties (32, 37, 40). The explanation for these observations is not yet apparent, although in some cases the difference may represent partial proteolytic digestion during purification, or partial binding to nucleic acid. With the two forms of rabbit liver enzyme, which can be separated by isoelectric focusing (47) or chromatography on phosphocellulose ( 3 7 , 4 6 ) ,it has been shown that the two proteins have similar structural and catalytic properties, but significant differences can be detected (37). It has been suggested (37) that the minor form of rabbit liver enzyme (amounting to one-third of the total activity) may represent the species found in mitochondria (17).

c.

PURITY

A N D SPEClFrC

ACTIVITY

Several of the purified tRNA nucleotidyltransferase preparations are thought to be homogeneous, based on electrophoresis in polyacrylamide 47. Deutscher, M. P. (1970). JEC 245, 4225. 48. Deutscher, M. P., and Hilderman, R. H. (1974). J . Bacteriol. 118, 621.

7. tRNA NUCLEOTIDYLTRANSFERASE

189

gels under native or denaturing conditions (Table I). These preparations include the enzymes from E. coli B (3.5), E. coli MRE 600 (40), L. ctcicl(~philus(401, baker’s yeast (34, 3 8 ) , L . lrrterrs seeds ( 4 / ) , houseflies ( l o ) ,and rabbit liver ( 3 7 ) .However, only in the case of the yeast (34) and rabbit liver enzymes (37) has this conclusion been confirmed by sedimentation equilibrium ultracentrifugation. Both the yeast (34, 38) and rabbit liver enzymes (37) are devoid of nucleic acid, as determined by their UV absorption spectra. The specific activities of purified tRNA nucleotidyltransferases have caused some confusion because different preparations from the same source show extremely wide variations. This is particularly true of the E. coli enzymes for which earlier preparations had specific activities below 100 pmol of AMP incorporated per hr per mg of protein (Table 1,l-3), whereas later preparations (Table I,4-6) had values as high as 8000, despite the fact that all the preparations were thought to be highly purified. Part of the problem is due to the level of purity of the various preparations; this is the reason for the differences between enzymes 4, 5, and 6 in Table I, because 4 and 6 are known to be only about 25-40% pure. Also many of the enzymes were assayed under suboptimal conditions, with ATP concentrations at, or below, K m and with impure tRNA acceptors. However, neither of these points would explain the very large differences between the first and second groups of E. coli preparations. It is known that some of these enzyme preparations are unstable (3.5),and that denatured enzyme tends to copurify with active material (37), so one possible explanation for the difference between preparations may be the amount of inactive protein present (39). The specific activities shown in Table I are for assays under standard conditions used during purification. Schofield and Williams (39) have estimated that the pure E. coli B enzyme (Table I,4) assayed under optimal conditions would actually have a specific activity of about 24,000 pmol of AMP incorporated per hr per mg of protein (converted from their units), corresponding to a turnover number of 350 per second. Where possible, similar estimates have been made for some of the other preparations, correcting for the purity, assay conditions and K m values reported. These estimates are summarized in Table 11. Without considering that the quality of the tRNA acceptor, or some other unknown variable, could influence the specific activity measurements, it appears that the prokaryotic enzyme has a higher turnover number than the enzyme from eukaryotes. Nevertheless, the turnover numbers, under optimal assay conditions, for both the prokaryotic and eukaryotic tRNA nucleotidyltransferases are extremely high for enzymes

1 90

MURRAY P. DEUTSCHER TABLE I1 ESTIMATED SPECIFIC ACTIVITIES AND TURNOVE OF R PURE tRNA DY LTRANSFERASES NUCLEOTI

Source 4." E. coli B

6. 8. 9. 10. 12.

E . coli A19 Bakers' yeast Bakers' yeast L. h r m s seeds Rabbit liver

Maximum specific activityb (pmoVhr/mg)

Turnover number (sec-')

24,000 19,000 6,000 1,500 750 5,000

350 260 120 10 60

Reference (39) (45 ) (54 ) (38)

(41) (37)

Number of the enzymes correspond to those in Table I. Maximum specific activities were estimated by correcting for the reported purity of the enzyme, temperature of the assay and concentration of substrates relative to K,. 'I

synthesizing phosphodiester bonds, especially since these enzymes do not catalyze a processive reaction (as do other nucleic acid polymerases) but must dissociate from tRNA after each AMP addition.

D. PHYSICAL A N D CHEMICAL PROPERTIES 1.

Molecikir Weight

The molecular weight of tRNA nucleotidyltransferases from various sources generally falls in the range of 40,000 to 52,000 (Table I), based on sodium dodecyl sulfate-acrylamide gel electrophoresis or gel filtration. In the case of the rabbit liver enzyme (37), the molecular weight has also been confirmed by sedimentation equilibrium ultracentrifugation. Two exceptions to the narrow molecular weight range for tRNA nucleotidyltransferases have been observed. The first is the enzyme purified from yeast by Sternbach el al. (M), which has a molecular weight of about 70,000, both by gel electrophoresis and equilibrium centrifugation. In addition, its szO,,,, of 4.2 S is somewhat higher than that of the rabbit liver enzyme (3.6-4.0 S). Another partially purified yeast enzyme also has a sedimentation coefficient of about 4 S determined by sucrose gradient centrifugation (11). The second exception is the enzyme purified from houseflies ( l o ) , which was reported to have a molecular weight of about 30,000, both by gel electrophoresis and chromatography on Sephadex G-200, and a sedimentation coefficient of 2.6 S. Since presumably the

7. tRNA NUCLEOTIDY LTRANSFERASE

191

identical reaction is carried out by all tRNA nucleotidyltransferases, and considering that the bacterial and liver enzymes have almost identical molecular weights, it is surprising that the yeast and insect enzymes diverge so far. Further work is necessary to determine whether the substantial differences in size (greater than twofold between the yeast and insect enzymes) has any functional significance. In all cases examined (Table I), the molecular weight of tRNA nucleotidyltransferase determined by gel electrophoresis under denaturing conditions is essentially identical to that measured by nondisruptive methods, indicating that all of these preparations are single polypeptide chains. Thus, despite the fact that tRNA nucleotidyltransferases must interact with and distinguish between tRNAs with different 3' termini, as well as distinguish between ATP and CTP in order to synthesize an accurate -C-C-A sequence, they are relatively small and uncomplicated struct urall y . 2. Chemical Properties

An amino acid composition has been obtained only for the two forms of the rabbit liver enzyme (37). These homogeneous enzymes contain no unusual amino acids and display no unusual patterns in their amino acid composition. Although ammonia was not determined, a considerable fraction of the glutamic acid and aspartic acid residues must be amidated because, despite an excess of potential acidic over basic residues (about 92 versus 69), the isoelectric points of the two proteins are slightly basic, at 7.2 and 7.5. Interestingly, the yeast enzyme, which has a very different molecular weight, also has an isoelectric point of 7.5 (34). On the other hand, the E. coli enzyme, which has a similar molecular weight, has a considerably lower isoelectric point of 5.9 (39) or 6.4 (49). The two forms of rabbit liver enzyme differ with respect to half-cystine residues; one contains seven, the other four. Many of these residues may exist as the disulfide (37). Both forms of the rabbit liver enzyme are relatively insensitive to reagents reacting with sulfhydryl groups, the one with only four half-cystines being almost totally resistant (37). Other partially purified mammalian tRNA nucleotidyltransferases from rabbit muscle (50) and Ehrlich ascites cells (51) are also quite insensitive to these reagents. In contrast, the enzymes from,?. coli ( 5 2 , 5 3 ) ,yeast (54,55), and 49. 50. 51. 52. 53. 54. 55.

Carre, D. S., and Chapeville, F. (1974). Biochirnie 56, 1451. Anthony, D. D., Starr, J. L., Kerr, D. S., and Goldthwait (1963). JBC 238, 690 Girgenti, A. J., and Cory, J. G. (1976). Intern. J . Biochem. 7, 275. Miller, J. P., and Philipps, G. R. (1971). JBC 10, 1280. Carre, D. S., and Chapeville, F. (1974). BBA 361, 176. Sternbach, H., Sprinzl, M.,Hobbs, J. B., and Cramer, F. (1976). EJB 67, 215. Kroger, M., Sternbach, H., and Cramer, F. (1979). EJB 95, 341.

192

MURRAY P. DEUTSCHER

plants (56) are affected by sulfhydryl inhibitors. In the case of the E. coli enzyme, one report (52) indicates that both AMP and CMP incorporation are affected equally, whereas another (53) demonstrates that only AMP incorporation is sensitive to the reagents. This discrepancy remains to be explained. Cramer and co-workers have shown that modified tRNA molecules derivatized near the 3' terminus with N-hydroxysuccinimide esters of bromoacetic acid (54),mercuriacetic acid (54), or iodoacetamide (55) can form covalent 1 : 1 adducts with the yeast enzyme and inactivate it. Since these reagents are known to react with -SH groups, and the modified enzyme can be reactivated with 2-mercaptoethanol, these workers have suggested that an essential cysteine residue is located at the active site of yeast tRNA nucleotidyltransferase. Also, since the enzyme can be protected against inactivation by ATP or CTP ( 5 4 ) ,these results suggest that the cysteine residue interacts with both the 3' terminus of the tRNA acceptor and the nucleoside triphosphate donor. 3. Stnrctural Metml

It has been suggested that many nucleotidyltransferases may be metalloenzymes (57). Some support for this idea in the case of tRNA nucleotidyltransferase has been obtained for the E. coli enzyme. Treatment with EDTA (35)or o-phenanthroline (39, 58) can irreversibly inactivate this enzyme. It appears that prior to the irreversible inactivation, the enzyme goes through an intermediate stage in which inhibition can be reversed by dilution or by addition of divalent cations (39). Addition of o-phenanthroline to assays affects AMP incorporation by the E. coli enzyme (58, 59), but does not affect CMP incorporation (58). On the other hand, tRNA nucleotidyltransferases from rabbit liver (59), yeast (57), or Rous sarcoma virus (57)are not inhibited by preincubation or inclusion of a-phenanthroline in the assay. If these proteins are also metalloenzymes, the structural metal must be buried and inaccessible to the chelating agent. Further experimentation is required to resolve this issue. 111.

Catalytic Properties

Since purified tRNA nucleotidyltransferases contain no nucleotide material that could act as a template, the ability of these enzymes to regener56. 57. BBRC 58. 59.

Cudny, H., Pietrzak, M . , and Kaczkowski, J. (1978). Planrci 142, 29. Valenzuela, P., Morris, R. W., Faras, A., Levinson, W., and Rutter, W. J. (1973). 53, 1036. Williams, K. R., and Schofield, P. (1975). BBRC 64, 262. Evans, J. A . , and Deutscher, M. P. (1976). JBC 251, 6646.

7. tRNA NUCLEOTIDYLTRANSFERASE

193

ate an intact -C-C-A sequence at the 3' end of tRNA must be determined by a specific organization of subsites within the active site of the protein. Inasmuch as the. correct 3' terminal structure is required for the proper functioning of tRNA, the mechanism whereby tRNA nucleotidyltransferase achieves its high degree of fidelity is of considerable interest. Unfortunately, although a number of tRNA nucleotidyltransferases have been characterized with respect to simple catalytic parameters (Table 111), detailed studies of the mechanism, specificity, and active site of this enzyme have been quite limited. A. pH

AND

CATION EFFECTS

All tRNA nucleotidyltransferases studied to date have alkaline pH optima, generally in the range of pH 9- 10 for nucleotide incorporation (Table 111). The rate at pH 7.0-7.5 is about 25-40% of that at the pH optimum. The explanation for the high pH optimum in v i m is not yet clear, although kinetic analysis of the rabbit liver enzyme has suggested that at pH 7.0 dissociation of the tRNA product begins to affect the rate of the reaction, whereas this is not the case at pH 9.4 (60). In addition, examination of nucleotide incorporation into the model acceptor substrates (43),CpC and cytidine, has shown that this reaction occurs at pH 7.0 at less than 5% of the rate at pH 9.4 (44), but that the model acceptors are potent inhibitors at the lower pH. It was suggested ( 4 4 ) that the low rate of reaction with the model acceptors at pH 7.0 is due to the inability of the products to dissociate from the enzyme in the absence of the rest of the tRNA molecule, although dissociation can take place at pH 9.4. If we assume that the enzyme works under optimal conditions in vivo (presumably at about pH 7 . 3 , these results may indicate that in v i m the reaction follows a different pathway than in viva, or perhaps that in viva a more rapid dissociation of the product can be promoted by other factors. tRNA nucleotidyltransferases require a divalent cation for activity. Because the cation must interact with both the nucleoside triphosphate and the tRNA acceptor, the relative ability of different cations to function in tRNA nucleotidyltransferase catalysis represents a balance between their effects on each substrate. Generally, the highest rates of incorporation are observed with Mg'+, although Mn'+ and Co2+also support activity (36,53, 56, 61). However, the actual rates of nucleotide incorporation found with different divalent cations are also influenced by the nucleoside triphosphate substrate, the divalent cation concentration, the ionic strength and pH of the reaction mixture, and the acceptor RNA. 60. Evans, J . A., and Deutscher, M . P. (1978). JBC 253, 7276. 61. Deutscher, M . P. (1972). JBC 247, 459.

TABLE 111 CATALYTIC PROPERTIES OF HIGHLY PURIFIED tRNA NUCLEOTIDYLTRANSFERASES Apparent

Source 1." 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12.

E. coli MRE600 E . coli B E. coli B E. coli B E. coli M E 6 0 0 E. coli A19 L. acidophilus Bakers' yeast Bakers' yeast L. litreus seeds Houseflies Rabbit liver

Mg2' pH Optimum Optimum (mM) 9.5 9.0-9.4 9.5 8.5-9.3

9.0-9.4

-

10 >5 5-10

5

-

-

9.5

1

9.5

-

-

9.3-10

5-10

K, (mM)

Apparent K, ( F M )

ATP

CTP

tRNA-C-C

0.33 0.095 0.16 0.14 3.6 0.31 0.71

0.017 0.015 0.06

1.5 0.21 9 6.3 1.7 11 0.4

0.6 0.45 0.25 0.2 2

0.017 0.03 0.029 0.2 0.26 0.07 0.033 0.03'

5.5

-

-

12

tRNA-C

Inhibitor of incorporationb

tRNA-N

AMP

CMP

-

-

0.18 6.5

0.20

CTP (NC) CTP (C) CTP ( C ) CTP -

ATP (NC) A- (C) ATP ATP

13

-

11

-

6

7.8

-

4

-

CTP(C) CTP (NC)

-

(C) ATP ( S ) A-

Reference (33,49) (52 ) (36 )

(71 (40 1 (45 ) (40 ) (5-4 1

(381 (56 1 (10)

(61, 70)

Numbers of the enzymes correspond to those in Table I. Symbols in parenthesis are as follows: NC, noncompetitive inhibitor; C, competitive inhibitor; S, activator; no symbol, type of inhibition not determined. ' Determined under conditions in which saturation curve was hyperbolic, not biphasic.

7. tRNA NUCLEOTIDYLTRANSFERASE

195

The divalent cation can also influence the specificity of nucleotide incorporation. For example, in the presence of Mn2+a variety of anomalous reactions catalyzed by tRNA nucleotidyltransferase are stimulated, whereas normal reactions are inhibited (62-64). In fact, under these conditions 5 S RNA is a better substrate for CMP incorporation than tRNA-C (63), and tRNA-C-C-C is a better substrate for AMP incorporation than tRNA-C-C (64). From these observations one might conclude that tRNA nucleotidyltransferase actually catalyzes nucleotide incorporation into a variety of RNAs in vivo. It is only from studies of tRNA nucleotidyltransferase mutants (48), and the ability of the enzyme to accurately synthesize a complete -C-C-A terminus on tRNA, that one assumes that its in vivo activity is on this substrate. These results point out the pitfalls associated with extrapolating from in v i m reactions to in vivo functions. A further question about tRNA nucleotidyltransferase function in vivo and cation effects is raised by the finding that the rabbit liver enzyme is greatly stimulated by polyamines ( 5 9 ) ; the rates of both AMP and CMP incorporation are increased to levels higher than found at any Mg2+concentration. In addition, spermine increases the specificity of the enzyme for AMP and CMP incorporation into tRNA. Although polyamines decrease the requirement for Mg2+(from about 10 to 1 mM), the presence of a divalent cation is still essential for tRNA nucleotidyltransferase activity (53,56, 5 9 ) . These results demonstrate two separate requirements for cations in tRNA nucleotidyltransferase catalysis, only one of which can be satisfied by polyamines. Since polyamines are known to interact with tRNA (65), these results have been interpreted to indicate that the divalent cations are required for binding nucleoside triphosphates to the enzyme, but polyamines are the preferred counterion for tRNA, perhaps because they induce a subtle conformational change that cannot occur with metal cations. Direct evidence for interaction of the polyamine with the tRNA rather than the enzyme was obtained from studies with the model acceptors, CpC and cytidine (44). In this system spermine had no effect on the rate of either AMP or CMP incorporation. These data and others (66) raise the interesting possibility that the physiological counterions for tRNA in vivo are, at least partly, polyamines. 62. Klemperer, H. G . , and Haynes, G . R. (1967). BJ 104, 537. 63. Deutscher, M. P. (1973). JBC 248, 3108. 64. Deutscher, M. P. (1973). JBC 248, 3116. 65. Quigley, G . J., Teeter, M. M . , and Rich, A. (1978). PNAS 75, 64. 66. Sakai, T. T., and Cohen, S. S. (1976). Progr. Nucleic Acid Res. M a / . B i d . 17, IS.

196

B.

MURRAY P. DEUTSCHER

NUCLEOSIDE TRIPHOSPHATE DONORS

The substrate specificity of tRNA nucleotidyltransferase with respect to the nucleoside triphosphate donors has been reviewed by Sprinzl and Cramer ( 3 ) ,and will not be discussed in detail here. The enzyme displays a high degree of specificity for ATP and CTP among the natural ribonucleoside triphosphates, presumably to ensure that only a -C- C-A sequence is synthesized in vivo. The purified liver (61) and yeast (67) enzymes are devoid of activity with GTP or ITP. However, purified tRNA nucleotidyltransferases from a number of sources (10, 45, 49, 61, 67, 68) can utilize UTP in place of CTP, although a U-U sequence cannot be readily made (49, 68, 69). The rate of UMP incorporation varies between 2 and 10% of the rate of CMP incorporation, and the apparent K m is from 10-to 40-fold higher than that for CTP (45, 49, 61, 68). In addition, CTP is a potent competitive inhibitor of UMP incorporation (10, 49, 70), ensuring that only CMP is incorporated under normal conditions. Although tRNA nucleotidyltransferases from several sources have many similar properties, they differ considerably with respect to their nucleoside triphosphate sites. Thus, the apparent K m values for ATP generally vary from about 0.1 to 0.3 mM for the E. coli enzymes (Table 111), although one report gives the K m as 3.6 mM (40). The enzymes from yeast, higher plants, and houseflies give slightly higher apparent K m values for ATP, a range of 0.2 to 0.6 mM (Table 111). In contrast, the mammalian enzymes have K m values apparently one order of magnitude higher. Partially purified enzymes from rat liver (30) and rabbit muscle (50) have apparent K m values for ATP of 1.0 and 3.8 mM, respectively, and the purified rabbit liver enzyme has a K m of about 2 mM (61). However, in the case of purified rabbit liver enzyme, it was shown that the apparent K m for ATP increases with increasing concentrations of the tRNA-C-C acceptor. The binding constant for ATP in the presence of tRNA-C-C is 4.2 mM, but for binding to free enzyme it is 0.18 m M (Table IV). Thus, it is possible that ATP binds to all the tRNA nucleotidyltransferases with similar affinities, but the effect of the tRNA substrate, in the case of the mammalian enzymes, distorts the apparent K, values. In fact, no such effect of tRNA was observed in a kinetic analysis of the E. coli enzyme (71). In all cases the apparent K m values (or binding constants) for CTP are considerably lower than those for ATP, and generally are in the range of 67. 68. 69. 70.

Best, A. N . , and Novelli, G . D. (1971). A B B 142, 539. Sprinzl, M., Sternbach, H., von der Haar, F., and Cramer, F. (1977). EJB 81, 579. Deutscher, M. P. (1972). JBC 247, 469. Masiakowski, P., and Deutscher, M. P. (1980). JBC 255, 11240. 71. Williams, K. R . , and Schofield, P. (1977). JBC 252, 5589.

7 . tRNA NUCLEOTIDYLTRANSFERASE

I97

0.01 to 0.03 mM (Table 111). The values reported for the yeast enzymes (38, 54), about 0.2 mM, appear to be an exception. The apparently high affinity of tRNA nucleotidyltransferase for CTP has important implications for the specificity of synthesis of -C-C-A and will be discussed in more detail later (Section 111,E).Earlier reports (50, 6 / ) have shown that the rabbit muscle and rabbit liver enzymes give nonlinear double reciprocal plots for CTP (with estimated K , values for the liver enzyme of 0.004 and 0.4 mM), suggesting the presence of multiple CTP binding sites. Further studies of the rabbit liver enzyme (70) provide evidence that this nonlinearity results because CTP also binds at the ATP site and stimulates CMP incorporation from the CTP site. Since CTP binds to the ATP site with about the same affinity as ATP (70) and increases the V,,,, of CMP incorporation, it is clear that the curved double reciprocal plots arise from a combination of effects at two distinct sites. When CTP is prevented from binding to the ATP site, apparent K , values for CTP of about 0.03 m M are obtained (70). From inhibition studies, a binding constant for CTP of about 0.01 mM has been estimated (70). A further difference between the various tRNA nucleotidyltransferases is observed in their response to the presence of a second nucleoside triphosphate. I n all cases examined (Table 111) CTP inhibits AMP incorporation into tRNA-C-C; however, in some cases this inhibition was reported to be competitive (36,38,52),and in others not (33, 70). In the case of the rabbit liver enzyme (70), the noncompetitive inhibition was attributed to the binding of CTP at both the ATP site where it competes with ATP, and at the CTP site where it competes with the end of tRNA-C. Assuming that other tRNA nucleotidyltransferases have separate binding sites for ATP and CTP (see Section III,E), it is difficult to understand how CTP inhibition of AMP incorporation could be competitive with ATP, since high concentrations of ATP would not be expected to overcome binding of CTP at its own site. Perhaps for some enzymes the terminal C of tRNA binds so tightly that it cannot be displaced by CTP, and all the inhibition is due to competition at the ATP site. With the bacterial and yeast enzymes, ATP also inhibits CMP incorporation (Table HI), and in some cases this inhibition is competitive (38, 5 2 ) . However, for the mammalian enzymes, ATP stimulates CMP incorporation (50, 5 / , 70). In the case of the rabbit liver enzyme this stimulation does not require AMP incorporation, since the nonincorporated analogs, ADP and AMP-(CH2)-PP, also stimulated (70). This stimulation is evidence that ATP and CTP bind to the enzyme at separate sites, and also shows that the two sites interact, leading to stimulation of CMP incorporation. The mechanism for this stimulation remains to be explained.

198

MURRAY P. DEUTSCHER

C. KINETIC MECHANISM Detailed kinetic analyses of AMP incorporation into tRNA-C-C have been carried out for the E. coli (71) and rabbit liver (60) enzymes. Based on an initial velocity bisubstrate analysis, both enzymes were shown to proceed by a sequential mechanism (72). These results indicate that both substrates are bound to tRNA nucleotidyltransferase prior to release of any product, and suggest that enzyme-AMP is not an intermediate in the reaction. Earlier studies (43) with the rabbit liver enzyme designed to detect reaction of [‘TIATP with the protein, or release of [32PlPPifrom [Y-~’P]ATPin the absence of acceptor, also gave no evidence for formation of a covalent intermediate. Stereochemical analysis (73) of phosphodiester bond formation with the yeast enzyme and ATPaS as substrate have shown that no racemization occurs upon nucleotide incorporation, and that the reaction proceeds by inversion of configuration at the a-phosphorus (74). The simplest interpretation of these data is that the reaction proceeds by an SN2 mechanism in which the 3’ hydroxyl on tRNA-C-C displaces PPi by a nucleophilic attack at the (Y phosphorus of ATP. If the reaction proceeds in this manner it would also eliminate the possibility of an enzyme-AMP intermediate. Dead-end inhibition studies using the competitive inhibitors ADP and AMP-(CH&PP (60,71) and tRNA-C-Cp (60) supported a random mode of substrate addition for both the E . coli and rabbit liver enzymes. However, product inhibition studies using tRNA-C-C-A and PPi differed for the two enzymes. In the case of the E. coli enzyme (71) all the product inhibitions are noncompetitive, leading to the conclusion that the reaction catalyzed by the E . coli enzyme proceeds by a random mechanism. In contrast, product inhibition studies of the rabbit liver enzyme revealed that tRNA-C-C-A is a competitive inhibitor with respect to tRNA-C-C, and PPi is competitive with respect to ATP. The other two product inhibitions were noncompetitive, suggesting the formation of enzyme * tRNA-CC .PPi and enzyme. tRNA-C-C-A ATP ternary complexes. These data were interpreted to indicate that the rabbit liver enzyme acts by a rapid equilibrium random mechanism; this conclusion was confirmed by isotope exchange studies during the net reaction, and at equilibrium. It is not yet clear whether the difference in reaction mechanism between the two enzymes is significant, or whether it is due to the fact that the two studies were carried out at different pH values. The rapid equilibrium mechanism

-

72. Cleland, W. W. (1970). “The Enzymes,” 3rd ed., Vol. 11, p. 1 . Cleland’s nomenclature is used throughout. 73. Eckstein, F., Sternbach, H., and von der Haar, F. (1977). Biochernistrv 16, 3429. 74. Burgero, P. M. J . , and Eckstein, F. (1978). P N A S 75, 4798.

199

7. tRNA NUCLEOTIDYLTRANSFERASE

for the rabbit liver enzyme indicates that the rate-determining step is the interconversion of the central complexes, whereas for the E. coli enzyme it is probably the release of one of the products. However, studies of the rabbit liver enzyme at pH 7.0 (60) have shown that at the lower pH dissociation of the product also becomes more important. Thus, the apparent difference in mechanism between the two enzymes may reflect a relatively small difference in rate constants for the interconversion or dissociation steps. The finding that rabbit liver tRNA nucleotidyltransferase acts by a rapid equilibrium mechanism means that the kinetic constants determined for substrates and products are actually dissociation constants, which are summarized in Table IV. As noted in Section III,B, tRNA-C-C decreases the affinity for ATP, and ATP decreases the affinity of tRNA-C-C. It is also clear that ADP and AMP-(CH2)PPbind more weakly than ATP, and that the terminal residue of tRNA-C-C-A affects binding to the enzyme. This latter point is considered in more detail in the next section. D. RNA ACCEPTORS 1. Specijicity

tRNA nucleotidyltransferases display little or no specificity with respect to tRNA substrates. Thus, purified enzymes from E. coli (35,45) can add AMP or CMP to tRNA preparations from E. coli, yeast, or liver at similar rates. Likewise, purified enzymes from eukaryotes (13, 61) are equally active with tRNAs from various species. In addition, the enzymes TABLE IV S U M M A ROF Y DISSOCIATION CONSTANTS F O R RABBIT L I V E RtRNA NUCLEOTIDY LTRANSFERASE" Ligand

Source Free enzyme Enzyme + ATP Enzyme + tRNA-C-C Enzyme + tRNA-C ____

~~

ATP (mM)

PPi (mM)

tRNA-C-C

tRNA-C-C-A

(PW

(CLW

0.18 4.2

3.0(1.0)'

0.60 12 -

7.8( 16) 97 -

-

-

1.8

-

-

CTP (mM)

(0.01)

~~

Data from Evans and Deutscher (60) and Masiakowski and Deutscher (70). Values for dissociation constants were obtained from bisubstrate kinetic analysis or from inhibition studies (values in parentheses). "

200

MURRAY P. DEUTSCHER

can act on all the different amino acid-specific tRNA species since close to stoichiometric amounts of AMP and CMP can be added to bulk tRNA, and amino acid acceptor activity can be restored for all amino acids (35, 75). In fact, neither the E. coli nor rabbit liver enzymes discriminate between the different tRNA species in a preparation of total tRNA because the percentage repair for individual acceptor species (measured by aminoacylation) follows exactly the incorporation of AMP into the total population (76). Thus tRNA nucleotidyltransferases must recognize some common features of tRNA structure; this is an interesting contrast to the aminoacyl-tRNA synthetases, which recognize subtle differences among tRNA molecules. Although tRNA nucleotidyltransferases can utilize any type of tRNA as substrate, they are almost totally inactive with other types of nucleic acid. Synthetic polynucleotides and DNA are ineffective with either the E. coli (33)or rabbit liver ( 6 1 ) enzymes. Likewise, the E. coli enzyme is inactive with 5 S RNA, high molecular weight rRNA, and R17 phage RNA (33); however, it can incorporate AMP and CMP into a number of phage and plant viral RNAs after they have been treated with snake venom phosphodiesterase (77, 78). Venom diesterase removes the terminal -C-C-A sequence on these molecules, and possibly also fragments them to release a tRNA-like structure (77, 78). The purified rabbit liver enzyme can add AMP and CMP to 5 S RNA and rRNA (63, 64), but the rates of these reactions are slow compared to incorporation into tRNAs (1 to 15% with various 5 S RNAs, 1 to 4% with rRNAs). However, the apparent K , for 5 S RNA is almost identical to that for tRNA-C-C. The studies with 5 S RNAs revealed that the 3' terminal nucleotide of the RNA acceptor plays an important role in the specificity of tRNA nucleotidyltransferase (64).The rate of AMP incorporation into E. coli 5 S RNA, which contains a 3' terminal UMP residue, is only 6% as rapid as incorporation into wheat germ 5 S RNA, which has a 3' terminal CMP. Similarly, AMP incorporation into high molecular weight rRNA is solely adjacent to a CMP residue (64). A direct comparison of the rates of AMP incorporation into tRNAs with different 3' termini showed that addition to tRNA-C-A or tRNA-C-U proceeds only 2-4% as rapidly as into tRNAC-C (64). Nevertheless, it is possible to synthesize tRNA-C-A-A and tRNA-C-U-A with rabbit liver tRNA nucleotidyltransferase. Substitution of Mn2+for Mg2+increases the rate of AMP incorporation into tRNA-C-A 75. Sprinzl, M . , and Cramer, F. (1975). PNAS 72, 3049. 76. Deutscher, M. P., and Evans, J. E. (1977). J M B 109, 593. 77. Prochiantz, A . , B6nicourt, C . , Carre, D., and Haenni, A-L. (1975). W B 52, 17. 78. Busto, P., Carriquiry, E . , Tarrago-Litvak, L., Castroviejo, M., and Litvak, S. (1976). Atin. Microbiol. 127A, 39.

7. tRNA NUCLEOTIDYLTRANSFERASE

20 1

and tRNA-C-U, but decreases the rate into tRNA-C-C such that the anomalous acceptors become 30-40% as active. It appears that Mn2+may serve to anchor or orient the 3' terminal residue adjacent to the AMPdonating site. 2. Binding Kinetic studies of purified tRNA nucleotidyltransferases have generally given apparent Km values in the range of 5-15 pM for the various tRNA substrates (Table 111).One report (52) has suggested that the K, values for individual tRNA species may vary as much as 10-fold, but it is difficult to explain these results in light of the observation (76) that all tRNAs are acted on randomly. Measurements of dissociation constants for tRNA substrates are extremely limited. For the rabbit liver enzyme, the dissociation constant for tRNA-C-C binding to free enzyme is 0.6 pM, and this is increased to 12 pM in the presence of ATP (Table IV), which is identical to the apparent K m for this substrate. Protection constants for tRNA-N, tRNA-C and tRNA-C-C for stabilizing the E. coli enzyme against thermal inactivation are in the range of 0.2 to 0.4 p M (79). The high affinity of tRNAs for tRNA nucleotidyltransferase indicated by these experiments suggests that enzyme-tRNA complexes should be detectable, and they have been observed by gradient centrifugation ( I0, I I , 80, 8 I ) , binding to nitrocellulose filters (10, I I , 82), and gel filtration chromatography (37). For all the purified enzymes studied these data have revealed the presence of 1 : 1 complexes between tRNA and the enzyme (IO, I f , 37,81). In some kinetic studies tRNA-C-C-A has been shown to be a noncompetitive inhibitor of nucleotide incorporation into acceptor tRNAs (38, 71), whereas in others the inhibition was reported to be competitive (52,60,81). Since the direct binding experiments suggest that only a single tRNA can be bound to the enzyme at one time, it would be expected that high concentrations of substrate tRNA would completely prevent binding of the tRNA-C-C-A inhibitor, leading to competitive inhibition. The reason for this discrepancy is not yet apparent, although it has been suggested that tRNA-C-C-A binds to the enzyme solely through its terminal residues at the ATP and CTP sites (38) and perhaps this binding is too weak to be detected by physical methods. The kinetic and direct binding studies that have been carried out suggest that tRNA-N, tRNA-C and tRNA-C-C bind to tRNA nucleotidyltransferase with approximately equal affinities. In contrast, intact 79. Miller, J. P., and Philipps, G. R. (1971). Biochrmisfry 10, 1001. 80. Hondo, H. (1969). BBA 195, 587. 81. Carre, D. S . , Litvak, S., and Chapeville, F. (1974). BBA 361, 185. 82. Igarashi, S. J., and McCalla, J. I . (1971). Can. J . Biocliem. 49, 1308.

202

MURRAY P. DEUTSCHER

tRNA-C-C-A binds about an order of magnitude more weakly, as determined by inhibition studies (52, 60), direct binding ( / / ) , protection of the enzyme against thermal inactivation (38, 79), substrate properties (63), and ability to elute tRNA nucleotidyltransferase from affinity columns (46). The mechanism whereby addition of a single nucleotide residue to a nucleic acid can decrease affinity for the protein is not understood, but it undoubtedly serves to promote dissociation of the tRNA-C-C-A product to allow the enzyme to recycle. This phenomenon may be related to the ATP stimulation of CMP incorporation, such that binding of any ligand to the ATP site, including the terminal A of tRNA-C-C-A, would lead to accelerated dissociation of the tRNA molecule from the enzyme.

3 . tRNA Recognition and Model Acceptors Since tRNA nucleotidyltransferase acts on all tRNAs the mechanism by which the enzyme recognizes its tRNA substrate must utilize a structural feature, or features, common to all tRNAs. Early chemical modification studies demonstrated that modification of tRNA by hydroxylamine, which destroys cytosine residues (83), cyanoethylation, which affects pseudouridines (84), or sodium borohydride reduction, which destroys dihydrouracil residues (85, 86) have relatively little effect on the ability of tRNA to accept AMP. Likewise, substitution of uracil by fluorouracil(87) or low levels of bromination or methylation (88), do not impair AMP incorporation. On the other hand, treatment with nitrous acid (83,89) and UV irradiation (90) decrease AMP acceptance, but cross-linking of E. coli tRNA with near-UV light does not alter the rate or extent of -C-C-A addition (91). Removal of nucleotides from tRNA also may have little effect on recognition by tRNA nucleotidyltransferase. Elimination of as many as five nucleotides from the anticodon region (92) does not decrease AMP incorporation, and some fragments of tRNA can function as acceptors (32, 93). However, removal of 15% of the nucleotides from the 5' end of tRNA 83. Takanami, M., and Miura, K. I. (1%3). BBA 72, 237. 84. Rake, A. V., and Tener, G . M. (1966). Biochemistry 5, 3992. 85. Cerutti, P. (1968). BBRC 30, 434. 86. Igo-Kemenes, T., and Zachau, H. G. (1969). EJB 10, 549. 87. Giege, R., Heinrich, J., Wed, J. H., and Ebel, J-P. (1969). BBA 174, 53. 88. Rether, B., Weil, J. H . , and Ebel, J-P. (1965). Bull. Soc. Chim. B i d . 47, 1591. 89. Carbon, J. A. (1965). BBA 95, 550. 90. Harriman, P. D . , and Zachau, H. G. (1966). J M B 16, 387. 91. Carre, D. S . , Thomas, G., and Favre, A. (1974). Biochimie 56, 1089. 92. Chuguev, I. I., Axelrod, V. D., and Bayer, A. A. (1970). BBRC 41, 108. 93. Overath, H . , Fittler, F., Harbers, K . , Thiebe, R., and Zachau, H. G. (1970). FEES Lerf. 11, 289.

203

7. tRNA NUCLEOTIDYLTRANSFERASE TABLE V COMPARISON OF KINETIC CONSTANTS FOR tRNAs SUBSTRATES"

Substrate AMP incorporation tRNA-C-C CPC Cytidine CMP incorporation tRNA-C APC Cytidine

AND

MODELACCEPTOR

Apparent K, (mM)

(pmoVhr/ml enzyme)

0.004 12 80

59 4.5 3.1

0.002 8 90

89 2.4 1.3

Vmax

" Data from Masiakowski and Deutscher (43. 44, 96) using rabbit liver tRNA nucleotidyltransferase.

with spleen phosphodiesterase decreases incorporation by 50% (94), and half-molecules of tRNA do not protect against thermal inactivation (79). These results indicate that many aspects of tRNA structure can be dispensed with, leaving tRNA nucleotidyltransferase function unaffected, but that gross changes or removal of certain residues cannot be tolerated. Furthermore, the overall structure of tRNA must be important since most other RNAs are not acceptors, and denaturation of tRNALeU leads to loss of acceptor function (95). Considerable clarification of the recognition problem has been obtained from studies of model acceptor substrates. Compounds as small as dinucleoside monophosphates and nucleosides can function as AMP and CMP acceptors with rabbit liver tRNA nucleotidyltransferase (43, 96), although with reduced V,,, values and greatly increased apparent K, values (Table V). Nevertheless, at sufficiently high substrate concentrations, measurements of rate of nucleotide incorporation are quite easily made. The products of the reaction are the same as with tRNA (i.e., a single AMP residue is added to CpC, and only ATP and CTP function as donors) (43). Also, these compounds bind to the enzyme in the same position as the 3' end of tRNA (44). The fact that these compounds can substitute for tRNA as nucleotide acceptors indicates that a complete tRNA molecule is not required for catalysis. However, it is also clear that even though the small analog of the tRNA terminus is sufficient for catalysis to occur, the rest of the tRNA molecule plays an important role in stabilizing the binding of 94. Bernardi, A., and Cantoni, G . L. (1969). JBC 244, 1468. 95. Lindahl, T., Adams, A., Geroch, M., and Fresco, J . R. (1967). PNAS 57, 178.

204

MURRAY P. DEUTSCHER

Time (min) FIG. 1. Stimulation of CMP incorporation into the model substrate, cytidine, by the nonreacting fragment, tRNA-Xp. Reproduced from Masiakowski and Deutscher (96).

the acceptor moiety and, more importantly, in the efficiency of catalysis. This latter point can be demonstrated directly (96) since readdition of the rest of the tRNA molecule to the model system stimulates nucleotide incorporation as much as 60-fold, to levels obtained with the natural tRNA acceptor. For example, addition of tRNA-X, to the system in which CMP is incorporated into cytidine stimulates incorporation to the level attained with tRNA-C (Fig. 1). This is accomplished without any significant change in the apparent K , for either cytidine or CTP. Thus, two recognition regions of the tRNA substrate can be defined: (1) The reacting end, which is sufficient to trigger catalysis; and (2) the nonreacting portion, which is essential for obtaining the optimum catalytic efficiency. Each part of the substrate functions independently, and the covalent bond between the two parts is not necessary. This model system permits study of each of the two recognition regions of the tRNA separately, one by its ability to act as an acceptor, and the other by its ability to stimulate the reaction. In addition, comparison of nucleotide incorporation into tRNAs and into the model acceptors makes it possible to sepa96. Masiakowski, P., and Deutscher, M. P. (1979). JBC 254, 2585.

7. tRNA NUCLEOTIDYLTRANSFERASE

205

rate effects on tRNA structure from effects on the reaction. Several of these effects have been considered in Section II1,A. A substantial amount of information dealing with the first point has been obtained; i.e., the structural features necessary for the reacting end to function as an acceptor of AMP or CMP ( 4 4 ) . Of all the dinucleoside monophosphates tested (11 of a possible 16), only CpC is an active AMP acceptor (Table VI). A low level of activity is observed with the other three compounds containing a 3' terminal C (ApC, GpC and UpC) but all other dinucleoside monophosphates are devoid of activity (Table VI). Thus, the specificity of tRNA nucleotidyltransferase in this model system conforms exactly to the structure present at the 3' terminus of the natural acceptor, tRNA-C-C, and further emphasizes the importance of a 3' terminal cytidine residue for a compound to function as an active AMP acceptor. Deoxycytidine and (pdC)z are also inactive, indicating that the vicinal hydroxyls of ribose are necessary, However, ribose itself, or ribose plus cytosine, are not acceptors. Furthermore, increasing the charge on cytidine, as in CMP, CDP or CTP, eliminates acceptor activity (Table VI). The structural requirements for a compound to function as a CMP acceptor differ from those for AMP (44). In contrast to the very high degree of specificity observed for AMP incorporation into CpC, CMP can be incorporated into a variety of acceptors, although at different rates (Table VI). Since CMP is normally incorporated into tRNA-C, which terminates with -C, or into tRNA-N, which terminates with any one of the four nucleotides, it might be expected that the specificity for CMP incorporation is less demanding than that for AMP incorporation. In fact, it is interesting that the best dinucleoside acceptor for CMP is ApC (Table VI); this sequence is the one found most frequently at the 3' terminus of tRNA-C because the residue preceding the -C-C-A sequence is generally AMP (97). Thus, as observed with AMP incorporation, the model acceptor system for CMP incorporation appears to closely parallel the specificity seen with the natural tRNA acceptors. Since tRNA nucleotidyltransferase works equally well with all tRNA species, but recognizes tRNA-C-C-A, tRNA-C-C, tRNA-C and tRNA-N as different, the region of the protein that interacts with the 3' end of tRNA might be expected to determine the enzyme's specificity. The use of model substrates has proved that this assumption is correct, and has also shown that specificity and catalytic efficiency are separable features of enzymatic reactions. However, these results naturally raise the ques97. Sprinzl, M . , Grueter, F., Spelzhaus, A . , and Gauss, D. H. (1980). Nucleic Acids Rrs. 8, r l .

206

MURRAY P. DEUTSCHER TABLE VI NUCLEOTIDE INCORPORATION INTO MODELACCEPTOR SUBSTRATES" Pyrophosphate release (nmoV15 min) Substrateb CPC APC GPC UPC CPA CPG CPU APA GPU UPA UPU Cytidine CMP CDP CTP (pdC)z Deoxycytidine Ribose Ribose + cytosine

ATP

CTP

7 .I 0.2 0.3

1.2 4.4 1.4 0.5 0.4

0.5 <0.05 10.05 <0.05 <0.05 <0.05 io.05 <0.05 0.7 0.1 <0.05 C0.05 0.1 10.05 C0.05

0.3 0.3 0.7 0.2 0.5 0.2 0.3 0.3 0.2 10.05 0.4 10.05

-

co.05 __

Data from Masiakowski and Deutscher (44). All acceptor substrates present at 5 mM except ribose at 250 mM and cytosine at 50 mM.

tion of how the enzyme distinguishes between the various model acceptors such that CpC can be several orders of magnitude more active than a compound such as CpA, or 40-fold more active than ApC, for AMP incorporation (Table VI). Theoretically, this specificity could come from a higher affinity to bind CpC at the acceptor site, or alternatively, only CpC would be able to trigger the catalytic process. Measurements of apparent K,,,values for ApC and UpC as AMP acceptors, and comparison of various dinucleoside monophosphates as inhibitors of AMP incorporation into cytidine, have suggested that the different dinucleoside monophosphates have similar affinities (44).Thus, the inability of many of these compounds to act as AMP acceptors is probably due to an effect on the V,,, of the reaction. Although many of these compounds can bind to the active site, perhaps they are not anchored or oriented properly for reaction with

207

7. tRNA NUCLEOTIDYLTRANSFERASE

ATP unless the 3' residue is C, and an even better fit is obtained if both residues are C. In this connection it is interesting that Mn2+can stimulate AMP incorporation into the model acceptors and also change somewhat the specificity of the reaction (44), just as it does with tRNA-C-A and tRNA-C-U (64). E. THEACTtVE SITE A N D SEQUENCE

S Y N T H E S I S OF T H E

-C-C-A

Inasmuch as tRNA nucleotidyltransferase contains no nucleotide template, accurate synthesis of the -C-C-A sequence must be determined solely by the specificity of donor and acceptor subsites within the active site of the enzyme. The high specificity of AMP incorporation for the CpC model acceptor (44) clearly defines 3 subsites within the active site of the rabbit liver enzyme, one that binds the donor ATP moiety, and two others that specifically recognize the acceptor cytidine residues. Furthermore, the stimulation of the model reaction by nonreacting fragments of tRNA (96) defines an additional subsite that recognizes tRNA. Several lines of evidence indicate that the subsites binding ATP and CTP are different: First of all, as shown in Table 111, the apparent K , values for the two substrates are very different, and their ability to protect the enzyme from thermal inactivation also differs (38, 79); second, for the mammalian enzymes, ATP stimulates CMP incorporation (50,51, 63), and ADP and AMP-(CH2)PP, which are competitive inhibitors of ATP (60), stimulate CMP incorporation (70); third, the acceptor specificities for AMP and CMP incorporation in the model system are different; fourth, using the affinity reagent, periodate-oxidized ATP, it is possible to totally inactivate AMP incorporation under conditions in which substantial CMP incorporation remains (70); fifth, an E. coli mutant has been isolated (48) that has lost most of its AMP-incorporating activity, but retains completely its ability to incorporate CMP (45). Additional evidence suggests that the site that binds the CTP donor overlaps, or is identical, with the subsite that recognizes the terminal C residue on the tRNA-C-C acceptor. Clearly, this would be expected since once a CMP residue is added to tRNA-C to generate tRNA-C-C, the residue becomes the acceptor for incorporation of the subsequent AMP residue. CTP strongly inhibits AMP incorporation into CpC or cytidine, much more than it inhibits AMP incorporation into tRNA-C-C (70), despite the fact that competition between CTP and ATP should be the same in all cases. The simplest interpretation is that CTP competes with CpC and cytidine when these compounds are used as AMP acceptors. Furthermore, CTP is a potent inhibitor of the anomalous incorporation of

208

MURRAY P. DEUTSCHER

AMP into tRNA-C (70), which generates tRNA-C-A (see Section 111,F). In the absence of CTP the terminal residue of tRNA-C must occupy a position adjacent to the AMP-donating site, which is normally occupied by the terminal residue of tRNA-C-C. Addition of very small amounts of CTP, as low as 5 p M , leads to CMP incorporation followed by AMP incorporation. Thus, binding of CTP at its donating site must move tRNA-C to the position where it accepts CMP rather than AMP, indicating that either CTP or the terminal C residue of an AMP acceptor can occupy the CMP-donating site. We conclude from all these data that the active site, at least of rabbit liver tRNA nucleotidyltransferase, consists of multiple donor and acceptor subsites arranged in tandem that recognize the tRNA, the terminal residues of tRNA, ATP, and CTP, as shown in Fig. 2 (61). The subsites binding the terminal residues of tRNA also bind the donor triphosphate residues. The tRNAs that lack one, two, or three terminal residues would A. Normal Incorporation:

+

+

+

+ tRNA tRNA

+

+

-X - X -C

tRNA--

CTP CTP

-C

X-C

ATP

6. Anomalous Incorporation:

+

+

+

+

+

+

tRNA-

tHNA

X

-

C

ATP

tRNA

-

X

ATP

c

CTP

-c -

FIG, 2. Model for tRNA nucleotidyltransferase catalysis. (A) For normal nucleotide incorporation each of the tRNA acceptors is positioned adjacent to the appropriate donor subsite to ensure the accurate synthesis of the -C-C-A sequence. Subsites exist for recognizing the tRNA, each of the terminal C residues, and ATP. CTP binds in the same subsites as the terminal C residues of tRNA. (B) For anomalous incorporation in the absence of CTP, tRNA-C and tRNA-X may also be positioned a fraction of the time adjacent to the AMPdonating site and accept AMP to generate tRNA-C-A and tRNA-A. In the absence of ATP, CTP may be incorporated into tRNA-C-C from the ATP site. Reproduced from Masiakowski and Deutscher (44).

7. tRNA NUCLEOTIDYLTRANSFERASE

209

position themselves in the active site adjacent to the donor subsites in a way that depends on the terminal structure of the tRNA and the availability of nucleoside triphosphates. The accurate synthesis of the -C-C-A sequence obviously depends on the specificity of the subsites; at each step the enzyme must make a choice between incorporation of a correct or an incorrect nucleotide, and this will be determined by the specificity of the donating site and the concentration of nucleoside triphosphates, which also will affect the positioning of the acceptor. CMP will be preferentially incorporated into tRNA-N because the high affinity of CTP for its binding site(s) (see Table 111) would prevent this tRNA from binding adjacent to the AMP-donating site at normal CTP concentrations. Similarly, even low concentrations of CTP are known to be sufficient to prevent tRNA-C from binding adjacent to the AMPdonating site. On the other hand, CTP is a relatively poor inhibitor of AMP incorporation into tRNA-C-C, most likely because the two C residues anchor the tRNA adjacent to the ATP site. Furthermore, even though CMP can be incorporated into tRNA-C-C from the ATP-donating sites (see Section III,F), the rate of this reaction is only a few percent of AMP incorporation into tRNA-C-C, and the higher concentrations of ATP in cells, as compared to CTP, would also preclude CTP binding at the ATP site. Thus, even though the subsites of tRNA nucleotidyltransferase may not be absolutely specific for binding or nucleotide incorporation, synthesis of the -C-C-A sequence would be assured as long as both ATP and CTP were present. However, if either of these triphosphates is missing, the potential for misincorporation would increase dramatically. This is exactly what occurs in vitro (see Section 111,F). One feature of this model has not been verified experimentally; i.e., whether one or both sites that recognize C residues are also CMPdonating sites. In view of this ambiguity an alternative model that utilizes only a single CTP site must also be considered (Fig. 3). In this case, CMP would be incorporated into tRNA-N from the single CMP-donating site (step 1); the product, tRNA-C, which would be blocking the site (step 2), would translocate back one position allowing another CTP molecule to bind (step 3); the second CMP would be incorporated from the same donating site (step 4);and the product tRNA-C-C would be adjacent to the AMP-incorporating site for synthesis of tRNA-C-C-A (step 5 ) . This model predicts that tRNA-C can bind to the enzyme in two positions, adjacent to either the CTP or ATP subsites, and this obviously occurs since tRNA-C is both an AMP and a CMP acceptor (69). Since after step 1 tRNA-C would be adjacent to the ATP site, the potential for synthesis of tRNAC-A exists. Thus, one consequence of this model would be that tRNA-C must translocate more rapidly than AMP incorporation occurs.

210

MURRAY P. DEUTSCHER t

+

t

t

t

+

tRNA

1.

2.

tRNA

3.

tRNA - X

4.

tRNA - X

5.

tRNA - X

__ ~

~

X X

CTP

-C

C

CTP

__ c - c ~

C-C

ATP - A

3. Alternative model for tRNA nucleotidyltransferase catalysis. CMP incorporation into tRNA-X and tRNA-C comes from the single CTP subsite. In the presence of CTP the tRNA-C product translocates back one position and is fixed in position by the binding of CTP. In this position it accepts the second CMP residue. In the absence of CTP the tRNA-C product would be adjacent to the ATP site, and in position for synthesis of the anomalous product, tRNA-C-A. Reproduced from Masiakowski and Deutscher (70). FIG.

F. MISINCORPORATION OF NIJCLEOTIDES Under certain conditions purified tRNA nucleotidyltransferases from E. coli (45,49,67), yeast (98), and rabbit liver (69) can catalyze the misincorporation of nucleotides into tRNA (i.e., the synthesis of sequences other than -C-C-A). These anomalous reactions occur when either ATP or CTP are omitted from reaction mixtures. Thus, in the absence of ATP, additional CMP residues can be added to tRNA-C-C. In the case of the E. coli (49, 67) and yeast (98) enzymes misincorporation is limited to a single additional CMP residue, whereas with the liver enzyme as many as 7 or 8 additional residues can be added (69). This reaction can be inhibited by ATP, and for the liver enzyme, also by KC1 (63). On the other hand, the reaction is promoted by Mn2+(63). The rate of CMP addition to tRNAC-C proceeds at about 4% of the rate of normal CMP incorporation into tRNA-C (63, 98). It is also possible to add a single AMP residue to tRNA-C-C-C to generate tRNA-C-C-C-A (64,691. In the case of the rabbit liver enzyme, CMP residues also can be added to intact tRNA-C-C-A (63, 69, 99). This reaction proceeds at 1% the rate of normal CMP incorporation, and also can be stimulated by Mn2+(63). In the absence of CTP anomalous AMP incorporation occurs. AMP can be incorporated directly into tRNA-N (49, 98, 69) or tRNA-C (45, 49, 67, 69,981. For theE. coli (45) and yeast (98) enzymes these reactions proceed at less than 10% the rate of AMP into tRNA-C-C, whereas for the liver enzyme, rates as high as 40% are obtained for AMP incorporation into tRNA-C (61), but less than 10% for AMP into tRNA-N (61). At even 98. Rether, B . , Gangloff, J., and Ebel, J-P. (1974). EYE 50, 289. 99. Deutscher, M. P., and Ghosh, R. K. (1978). Nucleic Acids Res. 5, 3821.

7. tRNA NUCLEOTIDYLTRANSFERASE

21 1

TABLE VII S U M M A ROF Y ANOMALOUS PRODUCTS SYNTHESIZED W I T H tRNA NUCLEOTIDVLTRANSFERASE~ Acceptor substrate

Donor triphosphate

Product

tRNA-N tRNA-C tRNA-C-C tRNA-C-C tRNA-C-C-A tRNA-C-C-A

ATP ATP CTP CTP, then ATP CTP CTP, then ATP

tRNA-A, tRNA-A-A, tRNA-A-A-A tRNA-C-A, tRNA-C-A-A tRNA-C-C-C, tRNA-C-C-(C). tRNA-C-C-C-A tRNA-C-C-A-C-C tRNA-C-C-A-C-C-A

~~

“ References are in text. slower rates it is also possible to add as many as two or three AMP residues to tRNA-N (69, 98). A summary of the various anomalous products synthesized by tRNA nucleotidyltransferase is given in Table VII. The mechanism of misincorporation can be partly understood by the model presented in Fig. 2. In the absence of ATP, CTP can bind to the AMP-donating site and at a slow rate can be added to tRNA-C-C, which occupies its normal position on the enzyme (70). The presence of ATP prevents this reaction, both by competing with CTP for binding and by its own incorporation to remove the tRNA-C-C acceptor. On the other hand, the mechanism by which the rabbit liver enzyme adds CMP to tRNA-CC-A is still not understood. In the absence of CTP, tRNA-C can sit adjacent to the AMP-donating site and act as an AMP acceptor. Most likely, the presence of the 3’ terminal cytidine residue enhances this reaction. Presumably, the identical situation occurs with tRNA-N, although in this case the acceptor may occupy the correct position for AMP incorporation very infrequently, leading to the extremely slow rate of this reaction. No evidence has been obtained for AMP incorporation from a CMP-donating site (70). A N D HYDROLYSIS OF tRNA G . PYROPHOSPHOROLYSIS

The reaction catalyzed by tRNA nucleotidyltransferase is reversible. The apparent equilibrium constant for the forward reaction at pH 9.4 is approximately 1500, and at pH 7.0 it is 50 (60). The difference between the two pH values is mainly due to release of a proton during the incorporation reaction. The reverse reaction, the pyrophosphorolysis of tRNA can be detected by conversion of [3’PlPPi to [32P]ATPor CTP (29, 30) or by

212

MURRAY P. DEUTSCHER TABLE VIII COMPARISON O F FORWARD A N D REVERSE REACTIONS OF RABBIT L I V E RtRNA NUCLEOTIDYLTRANSFERASE ~~

~

~~

~

pH (pmoVhr/ ml enzyme) Reaction and additions" Incorporation (ATP + tRNA-C-C) None +-PP, (2 m M ) + tRNA-C-C-A Pyrophosphorolysis (tRNA-C-C-A None + ATP (0.5 mM) + tRNA-C-C

+ PP,)*

7.0

9.4

3.24 0.96 3.10

11.0 6.00 10.9

0.24 0.48 0.12

0.02 0.13 0.005

a All tRNA substrates and inhibitors are present at equal concentrations (0.75 mg/ml). * Since this reaction is extremely sensitive to the presence of tRNA-C-C, especially at pH 9.4, the actual rates of the reaction and the level of stimulation by ATP or inhibition by tRNA-C-C, may vary somewhat depending on the amount of tRNA-C-C contaminant initially present in the tRNA-C-C-A substrate.

release of radioactivity from terminally labeled tRNAs in the presence of PP, (100). As expected from the pH dependence of the equilibrium constants, the pH optimum for pyrophosphorolysis is only about 10% as rapid at pH 9.4 as at pH 7.0 (Table VIII). At its optimum (pH 7.0) the reverse reaction proceeds at only about 2% of the maximum rate of the forward reaction (Table VIII). These results suggest that the pyrophosphorolysis reaction is not significant in vivo. However, in vitro the reverse reaction has proven useful for preparation of terminally labeled tRNAs (101) and [32P]CTP(44) by isotope exchange. The pyrophosphorolysis and PPi exchange reactions have not been well studied with purified preparations of tRNA nucleotidyltransferase. Earlier work (29,30) suggests that the specificity of the reverse reaction is similar to that of nucleotide incorporation with respect to both nucleoside triphosphates and RNAs. Blockage of the vicinal hydroxyl groups of tRNA by aminoacyl or peptidyl groups prevents pyrophosphorolysis (102), and periodate oxidation and borohydride reduction of the 3' ter100. Deutscher, M. P. (1973). BBRC 52, 216. 101. Ghosh, R. K . , and Deutscher, M. P. (1978). Nucleic Acids Res. 5, 3831. 102. Pulkrabek, P., and Rychlik, I. (1969). BBA 179, 245.

7. tRNA NUCLEOTIDYLTRANSFERASE

213

minus of tRNA decreases the rate of reaction with PPi (103). Further studies of the reverse reactions should prove useful for understanding the action of tRNA nucleotidyltransferase. At high pH values purified rabbit liver tRNA nucleotidyltransferase also catalyzes an unusual hydrolytic reaction, the removal of CMP from tRNA-C-C (100). This reaction is optimal at pH 10, and is undetectable at pH 7.0. Hydrolysis is not observed with tRNA-C-C-A, tRNA-C-A, tRNA-C-U or tRNA-C. The reaction requires Mg2+, is stimulated by Mn2+, and is inhibited by nucleoside triphosphates and increased ionic strength. The significance of this reaction is unknown, but its specificity suggests that it is associated with only one of the subsites in the active site.

IV.

Biological Role

For many years the biological role of tRNA nucleotidyltransferase was something of a mystery. The presence of a -C-C-A sequence on all tRNA molecules, and the existence of an enzyme that could synthesize this sequence, naturally led to the conclusion that the enzyme is involved in the biosynthesis of these residues in tRNA ( 2 ) . In addition, it was known that the terminal residues of tRNA, especially the terminal AMP, turned over independently of the rest of the'tRNA chain ( 2 ) , and it was assumed that tRNA nucleotidyltransferase also played a role in this process. These ideas were thrown into doubt, however, with the isolation and sequencing of a tRNA precursor (104), since the -C-C-A sequence was already present in the primary transcript and was undoubtedly encoded within the gene for the tRNA. Studies with other precursors confirmed that encoding of the -C-C-A sequence within the gene is a general phenomenon in E. coli (105). This raised the question of the function of tRNA nucleotidyltransferase, since in most cases turnover of the terminal residues is limited to the terminal AMP residue, whereas the enzyme can accurately synthesize the complete -C-C-A sequence. Some clarification of this paradox has been obtained from continued sequencing of tRNA precursors and tRNA genes, and from isolation of mutants deficient in tRNA nucleotidyltransferase. It is now clear from further sequence studies that the situation in E. c d i differs from that in 103. von der Haar, F., Schlimme, E., Gornez-Guillen, M., and Cramer, F. (1971). EJB 24, 296. 104. Altrnan, S . , and Smith, J. D. (1971). Nature New Biol. 223, 3 5 . 105. Chang, S . , and Carbon, J. (1975). JBC 250, 5542.

214

MURRAY P. DEUTSCHER

other organisms. In all other cases, including mitochondria and chloroplasts, tRNA genes do not encode the -C-C-A sequence (106-110) and must be added posttranscriptionally, presumably by tRNA nucleotidyltransferase. Also in E. coli, upon infection with T-even bacteriophages, some of the phage-specific tRNA precursors are devoid of an intact-CC-A sequence, although it is present in others ( I l l ) . Despite the fact that E. coli tRNA genes encode the -C-C-A sequence, studies with E. roli mutants deficient in tRNA nucleotidyltransferase (48) revealed that the enzyme is required for the normal growth of these cells (112).These studies also provided direct evidence that tRNA nucleotidyltransferase actually plays a role in tRNA metabolism since mutant cells contain tRNA molecules with defective 3' termini (48, 113). Thus, the enzyme must be involved in repair of tRNA molecules that are missing part of the 3' terminus. However, in agreement with the sequencing data, no evidence for involvement of tRNA nucleotidyltransferase in the biosynthesis ofE. coli tRNAs was obtained (113, 114). On the other hand, the biosynthesis of some bacteriophage TCspecified tRNAs does require tRNA nucleotidyltransferase (112, 115). The enzyme may also be involved in other aspects of bacteriophage development, but this function is still not understood (116). Thus, from these studies it appears that in most organisms tRNA nucleotidyltransferase plays a role both in tRNA biosynthesis and in tRNA repair. The involvement of tRNA nucleotidyltransferase in tRNA biosynthesis in eukaryotes is reenforced by the presence of this enzyme in mitochondria (17, 18), which are known to have a distinct tRNA-synthesizing system. However, some question has arisen with respect to the subcellular localization of the remainder of tRNA nucleotidyltransferase activity. One study of rat liver cells indicated that all the remaining activity is in the cytoplasmic fraction, and that nuclei are devoid of the enzyme ( 17 ) . However, recent studies of tRNA maturation in Xenopus laevis oocytes have 106. Goodman, H . M., Olson, M. V., and Hall, B. D. (1977). PNAS 74, 5453. 107. Valenzuela, P., Venegas, A., Weinberg, F., Bishop, R . , and Rutter, W. J . (1978). PNAS 75, 190. 108. Silverman, S., Schmidt, O . , SOU, D., and Hovemann, B. (1979). JBC 254, 10290. 109. Garber, R. L., and Gage, L. P.(1979). Cell 18, 817. 110. Eperon, I. C . , Anderson, S . F., and Nierlich, D. P. (1980). Nafiire (London) 286, 460. 111. McClain, W. H . , Seidman, J. G . , and Schmidt, F. J. (1978). J M B 119, 519. 112. Deutscher, M. P., Foulds, J . , and McClain, W. H. (1974). JBC 249, 6696. 113. Deutscher, M. P., Lin, J. J. C . , and Evans, J. A. (1977). J M B 117, 1081. 114. Morse, J. W., and Deutscher, M. P. (1975). JMB 95, 141. 115. Seidman, J. G . , and McClain, W. H. (1975). P N A S 72, 1491. 116. Morse, J . W., and Deutscher, M. P. (1976).BBRC 73, 953.

7. tRNA NUCLEOTIDY LTRANSFERASE

215

suggested that addition of the -C-C-A sequence takes place in the nucleus (117). It is not yet clear whether these differences represent different tRNA maturation pathways in the two tissues, or whether liver nuclear tRNA nucleotidyltransferase is very susceptible to leakage during the isolation procedure.

V.

Research Applications

tRNA nucleotidyltransferase has found widespread use for substituting unnatural nucleotides within the -C-C-A sequence of tRNA, and for incorporating radioactive residues into this sequence [see Ref. (3) for a review of this subject]. The enzyme has aiso proved useful for changing the length of the 3' terminal sequence, such as the synthesis of tRNA-C-A (98, 118) and tRNA-C-C-C-A (98, 119), in order to elucidate the role of these residues in tRNA function. Finally, tRNA nucleotidyltransferase has been used for synthesis of the model tRNA precursors tRNA-C-C-A[14C](C),(99) and tRNA-C-[**C]U (99, 120). These synthetic tRNA precursors have been used as substrates for isolation of tRNA processing nucleases (101, 120, 121).

117. 143. 118. 119. 120. 121.

Melton, D. A . , De Robertis, E. M . , and Cortese, R. (1980). Nrrtirre (London) 284, Tal, J . , Deutscher, M. P., and Littauer, U. 2. (1972). EJB 28, 478. Kirschenbaurn, A . H., and Deutscher, M. P. (1976). BBRC 70, 258. Schmidt, F. J . , and McClain, W. H. (1978). N u d e i c Acids Res. 5, 4129. Cudny, H., and Deutscher, M . P. (1980). P N A S 77, 837.

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Poly(A) Adding Enzymes MARY EDMONDS

I . Introduction and Perspective . . . . . . . . . . . . . . . . . . I1 . Purification and Properties . . . . . . . . . . . . . . . . . . . A . Ion Exchange Techniques . . . . . . . . . . . . . . . . . B . Af€inity Chromatography . . . . . . . . . . . . . . . . . . C . Criteria of Purity . . . . . . . . . . . . . . . . . . . . . D . Contaminating Enzymes . . . . . . . . . . . . . . . . . . I11. Multiple Poly(A) Polymerases . . . . . . . . . . . . . . . . . I v. Properties of Poly(A) Polymerase Proteins . . . . . . . . . . . . A . Size and Subunits . . . . . . . . . . . . . . . . . . . . . B . Other Physical Properties . . . . . . . . . . . . . . . . . . V.Assay . . . . . . . . . . . . . . . . . . . . . . . . . . . . VI . Stoichiometry . . . . . . . . . . . . . . . . . . . . . . . . VII . Substrates . . . . . . . . . . . . . . . . . . . . . . . . . . VIII . Primer Requirement . . . . . . . . . . . . . . . . . . . . . . IX . Ion Requirements . . . . . . . . . . . . . . . . . . . . . . . A . Divalent Cations . . . . . . . . . . . . . . . . . . . . . . B . Monovalent Ions . . . . . . . . . . . . . . . . . . . . . . C . Hydrogen Ions . . . . . . . . . . . . . . . . . . . . . . X . Inhibitors . . . . . . . . . . . . . . . . . . . . . . . . . . A . Product and Substrate Analog Inhibitors . . . . . . . . . . B . Antibiotics . . . . . . . . . . . . . . . . . . . . . . . . C . Intercalating Drugs . . . . . . . . . . . . . . . . . . . . D . Other Inhibitors . . . . . . . . . . . . . . . . . . . . . . XI . Kinetics and Reaction Mechanism . . . . . . . . . . . . . . . XI1 . Biological Role . . . . . . . . . . . . . . . . . . . . . . . . A . Polyadenylation in the Nucleus and Cytoplasm

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B . Subcellular Localization of Poly(A) Polymerases C . Nuclear Poly(A) Polymerases . . . . . . . . . XI11 . Regulation of Poly(A) Polymerases . . . . . . . . A . The Cell Cycle . . . . . . . . . . . . . . . .

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THE ENZYMES VOL . XV

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Copyright @ 1982 by Academic Press Inc . All rights of reproduction in any form reserved

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B. Mitogen Stimulation . C. Hormonal Stimulation D. Differentiation . . . . E. Metabolic Activation . XIV. Research Applications . .

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Introduction and Perspective

Poly(A) adding enzymes, commonly called poly(A) polymerases but more accurately RNA terminal riboadenylate transferases, were first described in 1960 (I). Within a few years many such enzymes had been partially purified and characterized from a variety of prokaryotes and eukaryotes [for a review of this early work see Ref. (2)]. Interest in these enzymes eventually diminished since poly(A) was not known to exist in animal cells. It was quickly revived, however, by the discovery of poly(A) sequences covalently linked to the 3’ end of both messenger RNA and heterogeneous nuclear RNA of animal cells. It is commonly assumed, although not yet convincingly demonstrated, that poly(A) polymerases are responsible for this addition. [The important characteristics of these enzymes from prokaryotes, eukaryotes, and animal viruses are reviewed in Ref. (2); for a review comparing poly(A) polymerases from normal and neoplastic tissues with an emphasis on extraction and purification procedures and specific inhibitors, see Ref. (.?)I. Recent work on these polymerases has centered less on enzymology and more on clarification of their role in polyadenylation reactions in the cell. Information accumulated since 1975, particularly on the multiplicity, cellular localization, and regulation is emphasized in this chapter. The poly(A) adding reactions considered here are restricted to enzymes that attach AMP residues from ATP to the 3’ end of polyribonucleotides through 3‘- to 5’-phosphodiester bonds according to the following reaction: Polyribonucleotide + n ATP

Mg” or Mn’+

polyribonucleotide (A).

+ n PP,

(1)

This definition excludes terminal deoxyribonucleotide transferases, other homopolynucleotide polymerases, primer-dependent ribonucleotide polymerases, and polynucleotide phosphorylases, although the last two activities may also add AMP residues to the 3’ end of polyribonucleotide primers. No attempt is made here to differentiate poly(A) polymerases on 1. Edmonds, M . , and Abrams, R. (1960). JBC 235, 1142. 2. Edmonds, M . , and Winters, M . A. (1976).Progr. Nitcleic Acid Res. Mol. B i d . 17, 149. 3. Jacob, S . , and Rose, K . (1978). Merhods Cancer R e s . 14, 191.

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the basis of lengths of the poly(A) synthesized since this depends on a number of obvious variables in the reaction conditions that have rarely been considered in the assay for any given poly(A) polymerase. Lengths falling within the 100-200 range characteristic of the poly(A) sequences in animal cell RNA are synthesized in v i m by several of the polymerases purified from animal cells, but sequences as short as 2 or longer than 1000 have been reported for other preparations. Poly(A) adding enzymes as defined here show a high specificity for ATP but a rather low specificity for primer. Most of the purified poly(A) polymerases are able to use a variety of polyribonucleotides and oligoribonucleotides differing in sequence and length as primers, although variations in their effectiveness have usually been noted. The divalent cation requirement has been the source of some confusion since poly(A) polymerases purified from different sources, and in some cases even from the same source ( 2 ) , respond differently to the presence of Mg2+ and MnZ+.As expected, the existence of different poly(A) polymerases was predicted from these data, and in the case of HeLa cells it has been quite clearly shown by the actual separation of two poly(A) polymerase activities that differ in their Mg2+and Mn*+requirements ( 4 ) . The significance of multiple poly-(A) polymerases remains unclear since it is not yet possible to correlate precisely the activity of any one polymerase with a polyadenylation reaction in the cell. Some proposals for specific roles for different poly(A) polymerases are considered in later sections. 11.

Purification and Properties

Poly(A) polymerases should be present in all living organisms since poly(A) sequences are apparently ubiquitous [for a compilation of their distribution, see Ref. (S)]. Purifications have been reported from a variety of viruses, bacteria, yeasts, plant and animal tissues, and cultured cells (2, 3). The most highly purified preparations from calf thymus, rat liver nuclei, HeLa cells, and E. coli have provided the basic information that defines the reaction in v i m . A.

ION

EXCHANGE TECHNIQUES

Purification schemes have usually relied on remarkably similar protocols. Except for organisms with cell walls, poly(A) polymerases, in contrast to RNA polymerases, are readily extracted from tissues, cells, or 4. Nevins, J . , and Joklik. W. (1977). JBC 252, 6939. 5. Karpetsky, T., Boguski, M., and Levy, C. (1979). Subcell. Biuchern. 6, 1.

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organelles by homogenization in neutral buffers of low or moderate ionic strength. A typical purification scheme begins with an ammonium sulfate fractionation of a crude extract to obtain a somewhat purified concentrate of enzyme that can be applied to a DEAE-cellulose column. The weak affinity of poly(A) polymerases for anionic binding sites is exploited by selecting a low ionic strength that prevents it, but not most other proteins including RNA polymerases, from binding. Primer dependence develops at this stage since endogenous polynucleotides are tightly bound. A subsequent binding to a negatively charged exchanger, usually phosphocellulose (or CM-cellulose or CM-Sephadex), results in a substantial purification. A final step on hydroxylapatite usually raises the purity significantly. A tabulated summary of the extraction and purification techniques used in poly(A) polymerase purifications from several animal and tumor tissues has been published (3).

B. AFFINITY CHROMATOGRAPHY Recent purifications, particularly those from limited sources such as cultured cells, embryos, and viruses, have incorporated affinity chromatographic steps into the protocol. Usually these steps replace more laborious ion exchange techniques, but in some cases they have been added to an existing,protocol as a final step. A variety of polynucleotides attached to celluloses or Sepharoses have been used. An early success involved a major purification of the poly(A) polymerase from core particles of vaccinia virions on DNA cellulose (6). Affinity chromatography has also been used to purify the poly(A) poiymerases from extracts of HeLa cells infected with vaccinia virus. In this case host HeLa cell poly(A) polymerases were also purified substantially by using DNA cellulose at a terminal step (4). A single-step purification on DNA cellulose of the rat liver nuclear enzyme from a crude nucleoplasmic extract has also been reported (7). Poly(A) polymerase from mouse embryo extracts was purified on Sepharose-bound transfer RNA (8). Chromosomal RNA prepared from rat liver chromatin and attached to Sepharose was effective as a terminal step in the purification of the polymerase from rat liver nuclei (9).

The molecular basis for binding poly(A) polymerases to polynucleotides attached to solids is unknown, but ionic interactions of the type displayed by phosphocelluloses may be involved since relatively high 6. 7. 8. 9.

Moss, B . , Rosenblum, E., and Gershowitz, A. (1975). JBC 250, 4722. Rose, K . , Roe, F., and Jacob, S. (1977). BBA 478, 180. Hadidi, A., and Sethi, S. (1976). BBA 425, 95. Antoniades, D., and Antonoglou, 0. (1978). BBA 519, 447.

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22 1

anion concentrations ( 100-200 mM) are apparently needed to displace the bound enzyme (10-13). A successful purification of a poly(A) polymerase from a crude extract of rat liver nuclei on ATP-Sepharose may represent a true affinity purification since the enzyme can be released by low levels of ATP (14). Oddly enough use of poly(A) attached to a solid support has been reported only once. This was for the purification on poly(A) Sepharose of the vaccinia virus poly(A) polymerase from infected HeLa cells ( 4 ) . There is little data with which to compare the effects of affinity chromatographic methods on the purity and properties of any one poly(A) polymerase preparation. In the few cases reported, the specific activities of such preparations have not greatly exceeded those of the same enzymes purified by conventional protocols. A possible exception is a chromatinbound rat liver nuclear preparation that has a specific activity after purification on chromosomal RNA Sepharose ( 9 ) about five times that of a rat liver nuclear enzyme reported to be homogeneous after a conventional purification (13).

C. CRITERIA OF PURITY Poly(A) polymerases fromE. coli ( l o ) ,calf thymus (11, 12), HeLa cells ( 4 ) , and rat liver nuclei ( 9 , 13) have been extensively purified. One preparation from calf thymus (12) and another from rat liver nuclei (13) are described as nearly homogeneous on the basis of correlation of enzyme activity with the electrophoretic mobility of a major protein band on a nondenaturing gel. Both preparations, however, showed significant amounts of nonenzymatically active protein that was ascribed to aggregation. A second piece of evidence to support claims for homogeneity was obtained when the calf thymus enzyme focused at the same pH as a major protein component during isoelectric focusing in gel (12), although again other nonenzymatically active proteins were present in the pH gradient. This preparation, that is nuclease-free, appears to be the most highly purified preparation reported thus far. Meaningful comparisons of the specific activities of different preparations cannot be made unless similar assays are used and divalent ion effects are considered. This has rarely been the case, but it is interesting to note that values for a number of preparations from similar and different 10. Sippel, A. (1973). EJB 37, 31. 11. Winters, M . , and Edmonds, M . (1973). JBC 248, 4756.

12. Tsiapalis, C . , Dorson, J . , and Bollum, F. (1975). JBC 250, 4486. 13. Rose, K . , and Jacob, S. (1976). EJB 67, 11. 14. Grez, M . , and Niessing, J . (1977). FEBS Lett. 77, 57

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sources range from 2000 to 10,000 nmoles/mg/hour. Two different highly purified Mn2+-activatedpreparations from rat liver nuclei that have been assayed with poly(A) as primer fall within this range. The specific activity of one preparation (13) was reported in the 1500-2500 range for different preparations, and the other as 6000 (9). A strikingly different value approximately two orders of magnitude higher has been reported for the highly purified Mn2+-activated calf thymus polymerase discussed previously (12). This discrepancy may result from the use of a small riboadenylate primer, ( A p ) A in the assay rather than poly(A), which is used for the assay of other poly(A) polymerases, although for this preparation poly(A) was said to be nearly as effective as (Ap)A on a molar basis (15). If not related to differences in the assay, then either this polymerase has a particularly high turnover number or most other preparations are far from pure. The latter seems unlikely in view of the relative homogeneity reported for the proteins of several highly purified enzymes.

D.

CONTAMINATING

ENZYMES

Activities that might interfere with the poly(A) polymerase assay, such as RNA polymerase, tRNA nucleotidyltransferase, polynucleotide phosphorylase, and ATPases, are apparently removed at early stages of purification since they are not detected in highly purified preparations from most animal tissues (11-13). Polynucleotide phosphorylase in bacterial extracts in the presence of an ATPase mimics a poly(A) polymerase activity and may have been mistaken for poly(A) polymerase in E. coli extracts (10). In one case, the activity of a primer-dependent ribonucleoside triphosphate polymerase in E. coli that was able to polymerize all four nucleotides was originally attributed to poly(A) polymerase (16, 17). Contamination with nucleases is a far more severe problem, and has not been adequately assessed with sufficiently sensitive assays in most cases. The Mn2+-activatedpoly(A) polymerase from calf thymus is an exception in that a systematic survey of both exo- and endonucleases throughout the purification showed nuclease activity remained until the final purification step (12). This preparation was judged nuclease-free when it was able to polyadenylate the 3' end of QP RNA without loss of its biological activity, although the activity is presumably lost by a single phosphodiester bond break (18). The purified Mg2+-activatedcalf thymus poly(A) polymerase 15. 16. 17. 18.

Tsiapalis, C., Dorson, J., deSante, D., and Bollurn, F. (1973). BERC 50, 737. Ohasa, S., and Tsugita, A. (1972). Nature New B i d . 240, 39. Ohasa, S., and Tsugita, A. (1976). J M E 105, 545. Gilvarg, C., Bollurn, F., and Weissrnann, C. (1975). P N A S 72, 428.

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does not alter the size of poly(A) or 28 S ribosomal RNA during a long incubation (11). The purified preparation from rat liver nuclei was originally reported to be free of both poly(A)- and RNA-degrading activity (f3),but was later reported to contain a 3’ exonuclease that degrades poly(A), but not poly A : U duplexes (19). Since this preparation had previously been described as homogeneous, the authors concluded that the exonuclease and polymerase functions were associated with the same protein.

111.

Multiple Poly(A) Polymerases

Perhaps the most important finding to come from recent purification studies is evidence for two different polyfA) polymerases in a single cell type. The possible existence of multiple enzymes was recognized many years ago when poly(A) polymerases from different sources showed striking differences in their response to Mg2+ and Mn2+ (2). Evidence for separate Mg2* and Mn2+-stimulatedenzymes in extracts of calf thymus nuclei was also reported, but the two activities were not separated (20). At that time a cautious approach to the interpretation of responses of polymerases to Mg2+and Mn2+seemed appropriate in view of the low purity of the preparations and the well-known anomalous effects substitution of Mn’+ may have on the substrate and template specificity of polynucleotide polymerases (21, 22). When two highly purified calf thymus poly(A) polymerases prepared by different laboratories were found to have different responses to Mg2+and Mn2+as well as to identical primers (If, 12), it appeared that different proteins had been purified. This conclusion was supported by obvious differences in apparent molecular weight, although some reservations exist about the significance of the value obtained for the Mg2+activated polymerase (If). Neither group reported the isolation of a second enzyme, although a Mn’+-activated poly(A) synthesis was lost during purification of the Mg’+-activated enzyme, and was not found among the other protein fractions (If). Other investigators have separated from a single extract as many as three fractions on cation-exchange celluloses that show polyf A) 19. Abraham, A . , and Jacob, S . (1978). PNAS 75, 2085. 20. Edmonds, M., and Abrams, R. (1965). “Symposium on Nucleic Acids,” p. 1 . Council of Scientific and Industrial Research, New Delhi. 21. Berg, P., Chamberlin, M., and Fancher, H. (1963). I n “Informational Macromolecules” (H.Vogel, V. Bryson, and J. Lampen, eds.), p. 467. Academic Press, New York. 22. Karkas, J . (1973). P N A S 70, 3834.

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polymerase activity. Two of the fractions from a rat liver nuclear extract were stimulated by Mn2+,but a third was activated by Mg2+(23). Three Mn2+-activatedfractions were separated from Chinese hamster embryo fibroblasts during phosphocellulose chromatography (24). Two of these fractions that had a requirement for Mn2+showed only minor differences in enzymatic properties, and no differences in isoelectric point, sedimentation coefficients, or size by gel filtration analysis. It is clear that separation of active fractions does not constitute proof for different enzymes since separations may result from incomplete removal of polynucleotides or from aggregation of enzyme proteins. Observed differences in primer or divalent cation requirements of separated fractions may also merely reflect differences in nuclease contamination of each fraction. Compelling evidence for the presence of two different poly(A) polymerases in the cell has come from a comparison of the physical properties and reactivity of two enzymes that were resolved during phosphocellulose chromatography of HeLa cell extracts (4). After each had been further purified on DNA-cellulose, their sedimentation coefficients and electrophoretic mobilities were compared. Although the apparent molecular weights calculated from velocity sedimentation analysis in density gradients were quite similar, the electrophoretic mobilities of the single protein bands observed for each enzyme in denaturing gels were clearly dBerent. The molecular weight calculated for each enzyme is shown in Table I. Properties of the two enzymes differ significantly. One uses only Mn2+ as the divalent cation, whereas the other uses either Mn2+or Mg2+.The Mn2+-activatedenzyme uses small oligomers such as (Ap)JA almost as well as total HeLa RNA as a primer, whereas the Mg2+-activatedenzyme showed a strong preference for RNA. The oligomer (Ap)JA in this case was about tenfold less effective than it was for the Mn2+-activated polymerase. These differences resemble those reported previously for the two poly(A) polymerases purified from calf thymus. Again in this case the Mn2+-activatedenzyme used small oligomers or poly(A) equally well as primers ( I 2 ) , but the Mgz+-activatedenzyme definitely preferred larger polynucleotides ( I I ) . The presence of two distinct poly(A) polymerases in animal cells (multiple enzymes in prokaryotes have not been reported) indicates that caution is needed in evaluating data describing the properties of a poly(A) polymerase. Ambiguities introduced by multiple polymerases could best be avoided by a separation of the activities, although extensive purifica23. Niessing, J. (1975). W B 59, 127. 24. Pellicer, A . , Salas, J . , and Salas, M. (1978). BBA 519, 149.

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tion of a single activity (as has apparently been done with the two calf thymus poly(A) polymerases) may also suffice. Unfortunately an inhibitor specific for only one of the two activities that could be used to assess their separation has not been found. IV.

Properties of Poly(A) Polymerase Proteins

A.

SIZE A N D SLJ8UNtTS

Aside from size estimates little is known of the molecular structure of any poly(A) polymerase. Since the subject was last reviewed ( 2 ) , molecular weights of enzymes purified from other sources have been obtained, and are shown in Table I. In denaturing gels a number of different preparations show a major band moving with a mobility similar to a 60,000 molecular weight polypeptide marker. The most reliable estimates of molecular weight are those for the Mn2+-activatedcalf thymus (12) enzyme where the molecular weight calculated from gel filtration and from velocity sedimentation analysis was 60,000. This value was also observed for the major protein band seen after electrophoresis in denaturing gels, suggesting that this poly(A) polymerase is apparently a single polypeptide. The TABLE I MOLECULAR WEIGHT O F PoLY(A)POLYMERASES Nondenatured Source Calf thymus Mn2+-activated Mg2+-activated Rat liver Nucleoplasm Hepatoma Mouse L cells HeLa cells Mn"-activated Mg2+-activated CHO fibroblasts Mouse embryos E . coli Vaccinia Cores Infected cytoplasm

Sedimentation

Gel filtration

60,000 -

60,000 120,000-140,000

80.000 63,000 58 ,OOO 45,000-60,000 65,000

80,000 70,000

Denatured gel electrophoresis

60,000 48,000

-

60,000

> 150.000 145,000- 155,000 65,000

75,000 50,000 50,000 51,000; 38,000 57,000; 37,000

Ref.

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two enzymes from HeLa cells also appear to be single polypeptides although they differ somewhat in size ( 4 ) . The molecular weights calculated from gradient sedimentation velocity analyses of each enzyme differed only slightly from the values estimated for single polypeptides in the denaturing gels. Most of the values reported in Table I have not been confirmed by a second analytical technique and must be considered as tentative. The molecular weights estimated for polypeptides in denaturing gels depend on the assumption that the enzyme is the predominant (if not the only) species in the preparation, since activities have not been measured in the major band. Other polypeptides with the same mobility may of course be present since analysis has been limited to a single set of conditions in all cases.

B. OTHERPHYSICAL PROPERTIES The physical properties of poly(A) polymerases have yet to be studied in detail. Isoelectric points of about pH 6.0 have been reported for the Mg2+-activated polymerases from calf thymus ( 1 1 ) and for the Mn2+activated enzymes of mouse L cells (25) and Chinese hamster ovary fibroblasts (24), suggesting these are weakly acidic proteins. The pH for the Mn2+-activatedpolymerase of calf thymus was estimated as 7.4 (12). The amino acid composition of the Mn2+-activated polymerases from rat liver and rat hepatoma showed no unusual features, although the two poly(A) polymerases differed significantly from each other in the content of 3 amino acids (13).

V.

Assay

Measurement of the moles of AMP from radio-labeled ATP incorporated into an acid-insoluble product provides the basis for assaying poly(A) polymerases. Although rapid and inherently accurate the assay is not specific, and may be quite unsuitable for assaying poly(A) polymerases in crude or partially purified extracts. As noted before, contaminating polymerases may produce acid-insoluble products labeled by ATP. Nucleases may also degrade the reaction product or produce new priming sites during the course of the reaction. Additional characterization of the reaction product can confirm the presence of a poly(A) polymerase. The simple kinetic parameters that characterize enzymes in general must, of course, be known before the assay can give meaningful data. 25. Giron, M. L. and Huppert, J. (1972).BBA 287, 438.

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VI.

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Stoichiometry

Equation (1) expresses the overall stoichiometry of the reaction. A direct correlation of AMP disappearance with pyrophosphate appearance has been reported only for E. coli polymerase (26)and the Mn2+-activated calf thymus poly(A) polymerase (12). The stoichiometry of the reaction with respect to primer is necessarily complicated by the fact that the product of the reaction also serves as a primer. This problem has been neatly solved in a kinetic study of poly(A) polymerase from E. coli (27) that is discussed in detail in the Section XI. VII.

Substrates

The utilization of ribonucleoside triphosphates other than ATP either singly or in the presence of ATP is usually less than one percent of that for ATP. The same is true for rADP and dATP. Levels higher than one percent most likely result from contaminating polymerases that polymerize single ribonucleoside triphosphates into homopolymers, or from ribonucleotide polymerases found in several tissues and organisms that synthesize RNA on RNA primers (16, 28, 29). Two types of ATP analogs have been tested as substrates. One, adenosine 5’-(/3,y-methylene)triphosphatewas efficiently polymerized into poly(A) with a polymerase from quail oviduct (30). The other, cordycepin triphosphate, 3’-dATP, which has been reported to be incorporated at the 3’ terminus of the primer (31) prevents further chain elongation. VIII.

Primer Requirement

The specificity of the primer requirement became a subject of great interest after poly(A) sequences were shown to be covalently attached to the 3’ end of both hnRNA and mRNA (32-34). Early reports of homology 26. August, T., Ortiz, P., and Hurwitz, J. (1962). JBC 237, 3786. 27. Sano, H . , and Feix, G.(1976). EJB 71, 577. 28. Longacre, S . , and Rutter, W. (1977). JBC 252, 273. 29. Boguslawski, G., Zehring, W., Meyer, R . , and Parr, J. (1977). JBC 252, 4337. 30. Miiller, W. E . , Totsuka, A , , Kroll, M., Nusser, I . , and Zahn, R. (1975). BBA 383, 147. 31. Miiller, W. E., Seibert, G., Beyer, R., Breter, H . , Maidhof, A., and Zahn, R. (1977). Cancer Res. 37, 3824. 32. Edmonds, M., Vaughan, M., and Nakazato, H . (1971). PNAS 68, 1336. 33. Lee, S., Mendecki, J., and Brawerman, G . (1971). PNAS 68, 3165. 34. Darnell, J., Wall, R . , and Tushinski, R. (1971). PNAS 68, 1321.

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among nucleotide sequences in the 3’ untranslated regions adjoining poly(A) in a- and P-globin mRNAs of rabbit and mouse and in a mouse immunoglobulin K light-chain mRNA (35) suggested polyadenylation recognition sites might reside either in the three-dimensional folding of this region or in a short sequence at the 3’ end. Newer sequence data on these untranslated regions of several other mRNA species has made this quite unlikely. Among all mRNAs examined thus far there is no common sequence at the RNA-poly(A) junction, nor is there extensive homology among these mRNAs in the 3‘ untranslated region (36). One striking exception that might serve as a poly(A) recognition site is the hexanucleotide, A-A-U-A-A-A, found about 13 to 20 nucleotides from the 5’ end of the poly(A) sequence in most eukaryotic mRNA sequences reported (36). There is evidence from other experiments, however, that specificity for polyadenylation may not reside in a specific polymerase site, but in an endonuclease site that when cleaved produces a 3’-hydroxyl end to which poly(A) can be added. For example, prematurely terminated transcription products recovered from UV-irradiated vaccinia virus particles are polyadenylated with a normal length poly(A) tail (371, yet the entire 3‘untranslated region of these shortened transcripts is missing, suggesting a specific addition site is unnecessary. This lack of a specific structural requirement for polymerase is compatible with the lack of specificity reported for primers of all poly(A) polymerases (2). A free 3’-hydroxyl end of an oligo- or polyribonucleotide appears to be the sole structural requirement for successful priming. Most poly(A) polymerases do not use polydeoxynucleotides as primers, although the enzymes purified from vaccinia virions ( 3 8 ) ,maize ( 3 9 ) ,and hamster fibroblasts (24) apparently do. This low specificity does not mean that polymerases are completely indifferent to primer composition, size, or shape but only that none have shown a striking dependence on one or more of these properties. The Mg2+-activatedcalf thymus enzyme, for example, uses poly(A), transfer RNA, and small RNA fragments from total calf thymus RNA equally well; but it uses HeLa 18 and 28 S ribosomal RNA, and MS-2 RNA poorly if at all. The small amount of radioactivity from ATP associated with the reaction products when the latter serve as primers does not cosediment with the recovered intact primers but with smaller fragments, apparently 35. Proudfoot, N . , and Brownlee, G . (1975). Nrrtiire (London) 252, 428. 36. Jou, M., and Fiers, W. (1978). “Biochemistry of the Nucleic Acids,” Vol. 11, p. 100. University Park Press, Baltimore, Maryland. 37. Gerschowitz, A . , and Moss, B. (1979). J . Virol. 31, 849. 38. Moss, B . , Rosenblum, E., and Gerschowitz, A. (1975). JBC 250, 4722. 39. Mans, R., and Huff,N . (1975). JBC 250, 3672.

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contaminating ribosomal RNA and MS-2 preparations (40). Primer studies that have not examined reaction products or assessed nuclease contamination of the polymerase are not readily interpreted, nor are data comparing priming efficiency of polynucleotides that are mixtures of unspecified molecular weights. Lack of specificity for the 3' terminal nucleotides was suggested when poly(C) and poly(I), but not poly(U), primed poly(A) synthesis with the Mg2+-activated calf thymus enzyme. This was confirmed when the product synthesized from [CI-~~PPIATP,using a mixture of calf thymus RNA fragments, showed ["PI transfer in about equal amounts to UMP, GMP, and CMP at 3' termini. The total ["PI in each nucleotide relative t o that in the AMP was close to that expected from the average length of the poly(A) made in the experiment (40). Similar low specificities have been reported for the Mn'+-activated calf thymus polymerase (12) and for the E. coli enzyme (10). Although the nucleotide sequence of primers appears to be unimportant, this is not necessarily the case for primer length. The Mg'+-activated enzyme from calf thymus (11) is primed far more effectively by longer poly(A) molecules than by short oligomers of AMP (A3-AI0). Although quantitatively less striking, the same is true for the E. coli poly(A) polymerase (10, 27). As noted earlier, the Mn2+-activated polymerase of calf thymus shows little preference for lengths of poly(A) primers (15). These differences constitute a part of the evidence for two distinct poly(A) polymerases in calf thymus. Observations on the two activities separated from HeLa cell extracts show similar differences in that the Mg'+activated polymerase preferred either longer poly(A) or RNAs rather than short oligomers of AMP, whereas the Mn*+-activated HeLa polymerase was indifferent to primer length (4). IX.

Ion Requirements

A. DIVALENT CATIONS The divalent cation requirement may be filled by Mg2+, MnZ+,or a combination of the two depending on the source of the polymerase. The optimal concentration is related to ATP levels, being either approximately equimolar or about twice that of the ATP concentration. The specificity of the divalent cation requirement previously noted that has provided part of the evidence for distinctive poly(A) polymerases (see Section 111) may in fact be influenced by the primer concentration. The enzyme from calf 40. Winters, M . , and Edmonds, M. (1973). JBC 248, 4763.

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thymus (12), which is more active in Mn2+than Mg2+when primed with low levels of p(A),, shows an increased ability for Mg2+to replace Mn2+as divalent ion as the concentration of this primer is raised. The maximal velocity of the reaction was the same in Mg'+ as Mn2+,although the K,,, for p(A), was four times higher in Mg". Some caution is needed in classifying polymerases on the basis of a specificity for divalent cations since partially purified enzymes are likely to contain nucleases. A partially purified poly(A) polymerase from calf thymus nuclei, for example, contains a Mg*+-activatednuclease capable of degrading poly(A) that is completely inhibited in the presence of Mn2+ (20).

B. MONOVALENT IONS Poly(A) polyrnerases are particularly sensitive to ionic strength. Monovalent cation concentrations needed to activate RNA polymerases (see Ref. 51) actually inhibit poly(A) polymerases. Levels of NaCl or KCl above 50 mM inhibit many poly(A) polymerases (6, I I-13). C. HYDROGEN IONS Most poly(A) polymerases are active in the range of pH 7 to 9 with an optimum somewhat above pH 8, where assays are performed.

X.

Inhibitors

Most of the compounds that inhibit DNA and RNA polymerases have been tested on poly(A) polymerases. With a few interesting exceptions to be discussed, the results have been those expected for a primer rather than a template-dependent polymerase. For investigational purposes the ideal inhibitor would be highly specific, not only for poly(A) polymerases in vitro, but also for cellular polyadenylation reactions. Such an inhibitor could clarify the role of multiple polymerases in v i m and correlate individual poly(A) polymerases with polyadenylation reactions in vivo, in much the way a-amanitin has been used to distinguish transcription carried out by each of the three eukaryotic RNA polymerases. In a broader sense, such an inhibitor could be used to examine the function of poly(A) sequences if polyadenylation could be uncoupled from transcription and other posttranscriptional processing reactions. Unfortunately such an inhibitor has not been found, but a systematic search has never been reported.

8. POLY(A) ADDING ENZYMES

A.

23 1

PRODUCT A N D SUBSTRATE ANALOGINHIBITORS

Pyrophosphate is an inhibitor of poly(A) polymerases (10-13). It has been shown to be noncompetitive for ATP and primer (tRNA in this case) in the E. coli poly(A) polymerase reaction (27). Inorganic phosphate has also been reported to be an inhibitor for the polymerases from E. coli (10) and from calf thymus nuclei ( I ] ) , but not for the rat liver nuclear polymerase (13). The basis for the inorganic phosphate inhibition has not been investigated. The primer analog ApA was a competitive inhibitor for tRNA primer but not itself a primer in the poly(A) polymerase reaction from E. coli (10, 27). Concentrations of the other ribonucleoside triphosphates, GTP, UTP, and CTP equivalent to that of ATP inhibit poly(A) polymerases (1 1-13). This property has often been used to distinguish these enzymes from RNA polymerases in crude extracts. The ATP analogs 2’-dATP and 3’-dATP are inhibitors of about equal potency for poly(A) polymerases (41-43). Arabinose ATP (Ara-ATP) inhibits the nuclear poly(A) polymerase from rat liver at levels that have little effect on the RNA polymerases ( 4 4 ) , a selectivity that differentiates it from 3’-dATP. Interest in 3’-dATP as an inhibitor stems from the fact that it is the biologically active form of the nucleoside antibiotic, cordycepin or 3’deoxyadenosine. Cellular poly(A) synthesis is highly sensitive to this drug, but heterogeneous nuclear RNA synthesis is much less so (45, 46). The transport or processing of nuclear RNA into cytoplasmic mRNA is greatly diminished apparently as a result of 3’-dATP inhibition of poly(A) synthesis (47). Although RNA polymerase I1 transcripts are less affected than RNA polymerase I (preribosomal RNA) transcripts in vivo, it has not been possible to distinguish effects of 3’-dATP on RNA polymerases I and I1 in vitro (48). The ATP analog, 3’-dATP, is not a particularly potent inhibitor of poly(A) polymerases in vitro, no more so in fact than 2’-dATP, the natural substrate for DNA polymerases (41-43). The inhibition with 3’-dATP is competitive with ATP (41-43), but there is conflicting data on the nature of the inhibition by 2’-dATP of the rat liver nuclear poly(A) 41. 42. 43. 44. 45. 46. 47. 48. 325.

Maale, G . , Stein, G . , and Mans, R. (1975). Nature (London) 255, 80. Rose, K . , Bell, L . , and Jacob, S . (1977). Nriture (London) 268, 178. (1978). FEES Left. 96, 354. Koch, G . , and Niessing, .I. Rose, K . , and Jacob, S. (1978). EBRC 81, 1418. Philipson, L., Wall, R., Glickman, R., and Darnell, J. E. (1972). PNAS 68, 3806. Mendecki, J . , Lee, S., and Brawerman, G. (1972). Biochemistry 11, 792. Penman, S., Rosbash, M., and Penman, M. (1970). PNAS 61, 1878. Desrosier, R., Rottman, F., Boezi, J . , and Towle, H. (1976). Nucleic Acids Res. 3,

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polymerase. One group observed competitive (42) and another noncompetitive inhibition with ATP in the reaction (43). Two obvious possibilities for the effects of 3’-dATP on cellular poly(A) synthesis are either that it serves as a competitive inhibitor of ATP in the poly(A) polymerase reaction as shown in vitro, or that it terminates chain elongation by being incorporated. If the latter occurs there would be no 3’-hydroxyl group on which to continue poly(A) synthesis. Evidence for the incorporation of 3’-dATP has not been found in vitw. Poly(A) chains recovered from cells treated with 3’-deoxyadenosine appeared to have adenosine, not cordycepin, at 3’ ends (46). A similar observation was made on the poly(A) chains synthesized in the presence of 3‘-dATP by the purified rat nuclear enzyme (13). A different conclusion was reached for the purified poly(A) polymerase of L cells (31). In this case the oligo(A) primer reisolated from a reaction inhibited by the presence of 3’-dATP was no longer effective as a primer of poly(A) synthesis in vitw. From these observations it is apparent that the mechanism of 3’-dATP inhibition of poly(A) polymerases in vitro is not completely clear, although competitive inhibition with ATP seems likely to play a major role. The situation in vivo is even less clear. The possibility that some mechanism related to another metabolic function of ATP altered by the presence of 3’-dATP in vivo should perhaps be considered.

B. ANTIBIOTICS High levels of actinomycin D, streptolydogen, puromycin, and a-amanitin have little effect on poly(A) polymerases ( 3 ) . However, certain derivatives of rifamycin that inhibit eukaryotic RNA polymerases also inhibit poly(A) polymerases (49, SO). The o-n-octyloxime of 3-formylrifamycin SV, AF/013, that inhibits RNA polymerases by preventing the formation of the polymerase-template initiation complex (51 ) also inhibits poly(A) polymerase apparently by binding to the enzyme (49). In this case chain elongation rather than initiation has been shown to be the site of inhibition (49). Although originally reported as competitive with ATP ( 5 2 ) , more recent studies show AF/013 to be a noncompetitive inhibitor of ATP (50). 49. Jacob, S., and Rose, K . (1974). Nircleic Acids Res. 1, 1549. 50. Nutter, R., and Glazer, R. (1979). Biorliem. P h r m c i r o l . 28, 2503. 51. Roeder, R. G. (1976). “RNA Polymerase,” p. 285. CSH Monograph. Cold Spring Harbor, New York. 52. Rose, K., Ruch, P., and Jacob, S. (1975). Biochemistry 14, 3598.

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C. INTERCALATING DRUGS As might be expected for a template-independent polymerase, ethidium bromide has no effect on poly(A) polymerases (8, 12), whereas proflavine inhibits only at very high levels (8, 12).

D.

OTHERINHIBITORS

Low levels of naturally occurring polyamines, such as spermine, that stimulate RNA polymerase I1 of rat liver inhibit the poly(A) polymerase from the same source (53).The inhibition observed when poly(A), nuclear RNA, or transfer RNA served as primer was not seen with short oligonucleotide primers such as (Ap),A (53). Polyamines appear to exert their inhibitory effects by binding to the primers. The sulfhydryl blocking reagent N-ethylmaleimide inhibits the Mn2+activated polymerases of rat liver (54) and calf thymus (55).

XI.

Kinetics and Reaction Mechanism

Published data on the kinetics of poly(A) polymerases have been limited, with one notable exception (27), to a few measurements of the time course of the reaction and to the effects of ATP or primer concentration on reaction rates. The latter rate measurements have been limited to variations in ATP concentrations at a single concentration of primer, and vice versa. Michaelis constants and maximal velocities have been calculated in some cases from such data. A K m for ATP of 70 phi' for the rat liver enzyme (13) and 50 pM for theE. coli enzyme (10) were reported. The K , for transfer RNA as primer for the E. coli polymerase is 0.2 pM (10).A K m of 50 ph4 was reported for p(A)3 as primer for the Mn2+-activatedcalf thymus polymerase that increases to 200 ph4 when Mg2+is substituted for Mn2+(12). Poly(A) synthesis observed with several purified polymerases (1I , 13, 25) has shown a pronounced lag that precedes the linear uptake of AMP. In some preparations this lag is abolished by raising the enzyme concentration (25),but not in others (11. 13). Preincubation of the Mg2+-activated calf thymus enzyme with o r without primer does not reduce the subsequent lag in AMP uptake (11). A much more rigorous kinetic analysis has 53. Rose, K . , and Jacob, S. (1976). ABB 175, 748. 54. Rose, K . , Roe, F., and Jacob, S. (1977). BBA 478, 180. 55. Coleman, M . S., Hutton, J . , and Bollum, F. (1974). Nuture (London) 248, 407.

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been applied to the purified E. coli polymerase (27). This study contains the requisite measurements to permit some predictions of the mechanism of this two-substrate reaction, and includes some clever techniques that allow the separation of the initiation and elongation stages characteristic of linear polymer formation. The effects of ATP and primer concentration (a mixed transfer RNA of E. coli was used) on the reaction velocities produces data that when transformed to a double-reciprocal plot gives a series of nonparallel lines that indicate a sequential reaction mechanism of either an ordered bi-bi, or a rapid equilibrium bi-bi reaction type (56). Inhibition of the reaction by product (pyrophosphate in this case) and by a substrate analogue allows these two possibilities to be distinguished. Pyrophosphate is a noncompetitive inhibitor for both substrates (ATP and tRNA). The dinucleoside phosphate, ApA, selected as a primer analogue because it is an inhibitor but not a substrate for the reaction, is a competitive inhibitor of tRNA, but not ATP. These results suggest an ordered mechanism in which primer is bound first and then ATP, which releases pyrophosphate followed by the polyadenylated primer product. In this same study an elegant double-label technique was developed to allow simultaneous measurement of initiation and elongation. The oligomer p(A,)A, labeled with 32P at the 5' end by the polynucleotide kinase reaction, was used as primer for the reaction with [3H]ATP. The assay conditions were adjusted so that only the primer molecules elongated by the addition of AMP residues were acid-insoluble. The primer added initially could then be readily separated from the polyadenylated reaction product, which allowed evaluation of the number of chains initiated. With this system kinetic analyses were done at varied ratios of ATP to primer, and of enzyme to primer. The initiation reaction was shown to reach a plateau rather quickly, whereas elongation proceeded continuously. Primer initiation was far more dependent on enzyme concentration than was the elongation reaction, indicating that the two reactions are essentially independent of each other. At low enzyme concentrations the rate of elongation remained quite constant at all levels of primer, whereas the initiation rate increased with increased primer concentration. From this observation it can be predicted that shorter poly(A) chains should be made at low concentrations of enzymes and high levels of primer. High levels of primer had previously been shown to reduce the poly(A) chain length, but the effect of enzyme concentration was not considered in this study ( 1 1 ) . In the case of the E. coli poly(A) polymerase under consideration here, a 40-fold increase in primer at low levels of enzyme reduced the average length of the poly(A) 56. Cleland, W.J. (1963). BBA 67, 104.

8. POLY(A) ADDING ENZYMES

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population from more than 300 nucleotides to about 50 (27). This ability to control the heterogeneity and length of the poly(A) synthesized on any RNA serving as a primer has practical uses that are discussed later. At higher concentrations of the E. coli polymerase both initiation and elongation rates increase as the primer concentration is raised. A V,,, for initiation of 6.7 and for elongation of 30 pmoYminute/unit was calculated. Thus, under the conditions of this experiment, the elongation rate is about five times faster than the initiation rate. If this is a general property of poly(A) polymerases it may account for the lag in AMP uptake noted previously for several of the polymerases from animal tissues and cells. The general similarity among properties of this E. coli poly(A) polymerase and those of eukaryotes makes it likely that kinetic parameters will also be similar, but the necessary experiments with a purified eukaryotic polymerase have yet to be reported. XII.

Biological Role

Fruitful speculation on the biological function of poly(A) polymerases was not possible until poly(A) sequences were found to be covalently bound to cellular RNA. When it was subsequently shown that poly(A) sequences are added posttranscriptionally to the 3' end of nuclear RNA it was obvious that this polyadenylation could be carried out by the primerdependent poly(A) polymerases previously found in animal cell nuclei ( I ). This correlation provided a satisfactory explanation of the function of poly(A) polymerases in cells that in essence remains valid. New information suggests, however, that this is an incomplete view of the functions of these enzymes in animal cells. A.

POLYADENYLATION I N THE NUCLEUS A N D CYTOPLASM

Evidence from two different lines of investigation indicates in one case that poly(A) synthesis in cells is more complex than first believed, and in the other that multiple poly(A) polymerases may exist in a single cell. It is logical to assume that these two findings are related. The existence of multiple poly(A) polymerases in a single cell (see Section 111) was at first puzzling since poly(A) synthesis was believed to occur only in the cell nucleus. Polyadenylated mRNA molecules in cytoplasm were presumed to be processed from polyadenylated RNA in the nucleus and transported to cytoplasm~(S7).Although this pathway appar57. Darnell, J . E . , Jelinek, W., and Molloy, G . (1973). Science 181, 1215.

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MARY EDMONDS

ently accounts for most of the poly(A) sequences in cytoplasm, an independent poly(A) synthesis also occurs there. Use of inhibitors of nuclear RNA and poly(A) synthesis, and labeling experiments with time periods shorter than the time needed for transport of mRNA out of the nucleus, clearly show poly(A) synthesis in cytoplasm. Cytoplasmic poly(A) synthesis differs from nuclear synthesis in being primarily an elongation of preexisting poly(A) rather than the de novo synthesis characteristic of synthesis in the nucleus. This was deduced in these experiments from the fact that lengths of labeled cytoplasmic poly(A) averaging 5-15 AMP residues, were found at the 3' end of much longer poly(A) sequences (58, 5 9 ) . A similar poly(A) elongation reaction is found in the nucleus. Poly(A) sequences of the usual 200-nucleotide length recovered from the nuclear RNA of mouse ascites cells (58) and HeLa cells (see 69) labeled for less than 5 minutes with adenosine also have short stretches of labeled AMP at the 3' end. Distinctive polyadenylation reactions in the cell may account for the presence of distinct poly(A) polymerases (see Section 111). However, from what is known of poly(A) polymerases from different cells, or even from the same cell, the properties (if any) that would assign participation of a single polymerase to one or the other of those reactions in vivo is not clear. The primer requirement might be expected to differentiate the reactions since the poly(A) adding enzyme might need only to recognize one or a short sequence of AMP residues, whereas the synthesis of poly(A) de novo could require a more complex sequence for initiation of poly(A) synthesis. In fact differences do exist in the primer requirements for the two poly(A) polymerases from calf thymus (11, 12) and HeLa cells ( 4 ) , but they are more apparent with respect to size than to sequence of the primer (see Section VIII). Compartmentalization of the polymerases may be a device used to segregate these two types of polyadenylation reactions it7 sifu. Evidence for compartmentalization of distinct poly(A) polymerases is considered in more general discussion of the intracellular localization of poly(A) polymerases.

B . SUBCELLULAR LOCALIZATION OF Poly(A) POLYMERASES Some important reservations should be noted before assigning subcellular locations to poly(A) polymerases because nearly all data have been obtained from aqueous homogenates of cells and tissues. The problem is less severe for a nuclear location since several poly(A) polymerases remain bound to well-washed (including detergent-washed) nuclei during 58. Diez, J., and Brawerman, G. (1974). PNAS 71, 4091. 59. Brawermm, G., and Diez, J . (1975). Cell 5, 271.

8. POLY(A) ADDING ENZYMES

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cell fractionation (11, 13, 30). The question is more complicated for cytoplasm since major translocations of nuclear enzymes may occur during cell fractionation (60). A case in point is the DNA polymerase a of eukaryotic cells that is found in cytoplasm rather than in the nucleus unless nonaqueous fractionation techniques are used (61). A major transfer of poly(A) polymerase from rat liver nuclei occurs in isotonic sucrose homogenates but is avoided in hypertonic sucrose (62). Although nonaqueous techniques have not been used to assess quantitatively the locations of poly(A) polymerases at subcellular sites, a polymerase from rat liver normally present in the cytosol of aqueous homogenates was reported to be absent in cytosol obtained from cells ruptured and fractionated in nonaqueous glycerol (63). In spite of the limitations of aqueous fractionation methods, it is usually assumed that poly(A) polymerases are present in the cytoplasm. Furthermore, from the results with calf thymus, it is assumed that the Mn2+activated polymerase found almost exclusively in cytosol(12) is responsible for the poly(A) elongation reaction that occurs in cytoplasm, whereas the Mg’+-activated enzyme from the nucleus (11) carries out de n o w poly(A) synthesis. Direct support for this view has come from a cell fractionation experiment in HeLa cells where two poly(A) polymerases with distinctive divalent cation requirements were chromatographically separated (see Section 111) ( 4 ) . Although activities overlapped to some extent, the Mn’+-activated enzyme was predominantly cytoplasmic, whereas the Mg‘+-activated enzyme was primarily nuclear. The fact that the hypotonic swelling used to rupture these HeLa cells results in a nuclear swelling that would favor leakage of nuclear proteins, again suggests that some caution be used in interpreting these results as evidence for cytoplasmic poly(A) polymerases. More convincing evidence for cytoplasmic poly(A) polymerases comes from experiments with the eggs of sea urchins. These eggs can be quite clearly separated into enucleate and nucleate halves (64). Two groups of investigators have found that most of the poly(A) polymerase activity of the egg is in the enucleate half (65, 66), as might have been expected from its concentration in the cytosol fraction of egg homogenates (65, 66). A 60. deDuve, C. (1971). J. C i 4 . B i d . 50, 20. 61. Kornberg, A . (1979). “DNA Replication,” p. 206. Freeman, San Francisco, California. 62. Rose, K., Lin, Y.C., and Jacob, S. (1976). FEBS Lett. 67, 193. 63. Avramova, Z., Milchev, G., and Hadjiolov, A. (1980). EJB 103, 99. 64. Harvey, E . N. (1956). “The American Arbacia and Other Sea Urchins.” Princeton University Press, Princeton, New Jersey. 65. Slater, D., Slater, I., and Bollum, F. (1978). De\v/op. B i d . 63, 94. 66. Egrie, J., and Wilt, F. (1979). Bioclremistry 18, 269.

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correlation has also been found in sea urchin eggs between the location of poly(A) polymerase activity and the rapid poly(A) synthesis that occurs on underadenylated cytoplasmic mRNA immediately after fertilization (67, 68).

C. NUCLEAR PoLY(A)POLYMERASES The evidence for nuclear poly(A) polymerases rests on claims that the activities purified from nuclei of calf thymus ( I f ), quail oviduct (30),and HeLa cells (4) account for nearly all of the Mg2+-activatedpolymerase extractable from those cells or tissues. Whether more than one poly(A) polymerase is present in the nucleus is less clear, although different activities would be compatible with the two mechanisms apparently needed to account for the poly(A) addition reaction and de nova poly(A) synthesis observed in the nucleus (59, 69) and in isolated nuclei as well (70). It is not easy to distinguish two such activities on the basis of divalent ion requirements since the Mg2+-activatedenzyme is also equally active in Mn2+. An actual separation of activities has been achieved with nuclei from rat liver (23) and from HeLa cells ( 4 ) by ion exchange chromatography of nuclear extracts. The separated fractions from each source differed in their divalent cation preference, and as already noted the two polymerases from HeLa cells are apparently different polypeptides, a criterion needed to establish the presence of different polymerases in the nucleus (see Section 111). The detection of different poly(A) polymerases in the nucleus would not necessarily dispose of the problem of contamination of the nucleus with cytoplasmic poly(A) polymerases, however. Fractionation into subnuclear components has provided conflicting evidence for different polymerases in the nucleus. Two groups of investigators compared poly(A) polymerases found in a soluble nucleoplasmic extract and a chromatin fraction from which polymerase was released in one case with DNase (71) and in the other by high salt detergent extraction (9). Poly(A) polymerase released by the latter technique was Mg2+activated, but the nucleoplasmic enzyme used only Mn2+. The enzyme released from chromatin could use nonpolyadenylated RNA as a primer but not poly(A), whereas the nucleoplasmic polymerase used only poly(A) or polyadenylated RNA. Other investigators found that the nucleoplasmic- and chromatin-released activities did not differ in divalent ion prefer67. 68. 69. 70. 71.

Slater, D . , Slater, I., and Gillespie, D. (1972). Notrrue (London) 240, 333. Wilt, F. (1973). PNAS 70, 2345. Sawicki, S . , Jelinek, W., and Darnell, J . E. (1977). JMB 113, 219. Kieras, R., Almendinger, R . , and Edmonds, M. (1978). Biochemistry 17, 3221. Rose, K., Roe, F., and Jacob, S. (1977). BBA 478, 180.

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ence or primer requirements (71). They did report, however, that in the chromatin-bound state, the poly(A) polymerase was activated by Mg2+, but on release to a soluble form the enzyme preferred Mn", a characteristic of the nucleoplasmic enzyme as well. The chromatin-released activity also showed a strong preference for exogenous poly(A) as primer, similar again to the nucleoplasmic enzyme. Since the activity released from chromatin appeared to account for most of the chromatin-bound activity, it was concluded that poly(A) polymerase bound to chromatin is the same enzyme found in the nucleoplasmic portion of the nucleus, but has altered properties when bound to chromatin. The explanation for these differences, which lead to opposite conclusions, is not immediately obvious but may arise from differences in techniques used to extract and purify each enzyme. Aqueous cell fractionation techniques obviously cannot give a completely satisfactory picture of the location of poly(A) polymerases in sitrr. While it seems certain that the nucleus contains a Mg'+-activated polymerase, and likely that cytoplasm contains a Mn2+-activatedpoly(A) polymerase, the number and type of each at any designated cell site remains uncertain. The situation is even less clear for other cell organelles from which poly(A) polymerases have been purified. A mitochondrial poly(A) polymerase from rat liver (72, 73) was shown to be distinct from polymerases in the cytosol(72). The significance of ribosomal-bound polymerases cannot be evaluated since there is a high probability of adventitious binding of cytosolic proteins, including poly(A) polymerase, to ribosomes (63, 74).

XIII.

Regulation of Poly(A) Polymerases

Interest in the regulation of poly(A) polymerase activity stems from its role in the processing of nuclear transcripts, where polyadenylation is apparently an early step in mRNA production (75). Regulation of gene expression at the level of translation may also involve poly(A) polymerases since underadenylated or nonadenylated mRNA molecules may require readenylation before they are translated. Data that might correlate changes in poly(A) polymerase activity with changes in the rates of synthesis of poly(A) or poly(A)-containing mRNA would be useful, although few experiments of this sort have been reported. This is not surprising in view of the difficulties in quantifying poly(A) polymerase 72. 73. 74. 15.

Rose, K . , Morris, H . , and Jacob, S. (1975). Bkdietnisfry 14, 1025. Cantatore, P., DeGorgi, C., and Saccone, C. (1976). BBRC 70, 431. Hardy, S., and Kurland, C. (1966). Biochemistry 5, 3668. Darnell, J. E. (1979). Prop-. Nucieir Acid Res. Mnl. B i d . 22, 327.

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activities in crude extracts and in measuring net changes in total mRNA content of the cell. Changes in either of these quantities must not only be reproducible but should be rather large to be convincing. A. THECELLCYCLE Cells dividing in suspension culture have rather constant levels of poly(A) polymerase activity throughout the cell cycle. Erythroleukemic (Friend) cells synchronized by isoleucine starvation showed twice as much poly(A) polymerase activity per cell during the period of DNA synthesis as cells in resting phase (76). Another study with mouse L cells synchronized by a double thymidine block showed only a small change in activity per cell on passing from G I to S phase. This contrasted with the activity of certain exo- and endonucleases in these cells whose activity increased severalfold on entering S phase (77). These results suggest that levels of poly(A) polymerase remain rather constant throughout the cell cycle. The doubling of activity observed in early S phase with the Friend cells may reflect de novo enzyme synthesis needed to restore enzyme activity after cell division.

B. MITOGENS T l M U L A T i O N Resting lymphocytes stimulated to divide with phytohemagglutinin showed a five- to sixfold increase in poly(A) polymerase activity during the time of DNA synthesis (55). Other enzymes such as DNA polymerases a and p showed similar increases at this time. A study of concanavalin A-stimulated lymphocytes (78) showed a marked increase in the labeling of poly(A) sequences in cytoplasm that preceded any change in the rate of RNA synthesis. The increased labeling was attributed to poly(A) turnover since neither the total poly(A) nor its length changed. STIMULATION C. HORMONAL

Two studies of poly(A) polymerase activity in target tissues of animals treated with sex hormones have been reported. Poly(A) polymerase activity in oviducts of quail treated with estrogens showed no change after primary stimulation, estrogen withdrawal, or restimulation (30). After 76. Adolf, G . , and Swetly, P. (1978). BBA 518, 334. 77. Miiller, W., Schroder, H., Arendes, J . , Steffen, R., Zahn, R . , and Dose, K . (1977). EIB 76, 531. 78. Hauser, H . , Knippers, R., and Schafer, K . (1978). ECR 111, 175.

8. POLY(A) ADDING ENZYMES

24 1

several days of progesterone administration to estrous rabbits, poly(A) polymerase activity of the uterus increased severalfold, whereas RNA polymerases I and I1 remained unchanged (79). The increase is abolished if estrogen is administered with progesterone. The increase is not due to decreased poly(A) degrading activity because the same nuclear extracts show marked increases in poly(A) degrading activities as well. Along with the enhanced poly(A) synthesis there is a 50-fold increase in the uteroglobin mRNA content of uteri after the progesterone treatment. Although the basis for the increase in poly(A) polymerase activity is unknown, the correlation with the enhanced production of a specific messenger RNA suggests that cells may regulate poly(A) synthetic and degradative activities when patterns of RNA processing are altered by progesterone treatment.

D. DIFFERENTIATION Poly(A) polymerase activity during cell differentiation has been examined in fertilized sea urchin eggs. These eggs are particularly suitable because the poly(A) content of cytoplasmic mRNA doubles by the time the fertilized egg reaches the two-cell stage (67, 68). Two studies with different species of sea urchin have shown that total poly(A) polymerase activity does not vary during development from the unfertilized egg to the 32-cell embryo stage (66, 80). However, there is a striking rearrangement in the subcellular location of the enzyme that is primarily in the cytosol of the egg, but is progressively transferred to the nucleus during embryogenesis (66). Eggs apparently have enough poly(A) polymerase to carry out this large increase in the rate of poly(A) synthesis that follows fertilization. Animal cells that can be stimulated in vitm to undergo differentiation, such as Friend cells and neuroblastoma cells, have given different results with respect to changes in poly(A) polymerase activity during differentiation. No increase in activity was detected after butyric acid was used to induce globin synthesis in Friend cells (76), from which it was concluded that no increase in enzyme is needed to ensure polyadenylation of the newly synthesized globin mRNAs. On the other hand neuroblastoma cells stimulated by dibutyryl CAMP to differentiate and form axons showed a significant increase in the poly(A) content of messenger RNA and a fourto sixfold increase in nuclear poly(A) polymerase activity (81). It has been shown that not all neuroblastoma cell lines show either increased 79. Orava, M . , Isornaa, V., and Jhne, 0. (1979). EJE 101, 195. 80. Morris, P., and Rutter, W. (1976). Biochemistry 15, 3106. 81. Sirnontov, R., and Sachs, L. (1975). EJB 55, 9.

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rates of synthesis or net increases in poly(A)-containing mRNA on treatment with cyclic AMP (82). Poly(A) polymerase activities were not reported for these cell lines, however.

E. METABOLIC ACTIVATION Organ perfusion studies have indicated that the catalytic function of poly(A) polymerases is metabolically regulated. Rapid changes in poly(A) polymerase activity were observed in rabbit hearts perfused with noradrenaline (83). Within 1 to 2 min of treatment poly(A) polymerase activity in the cytosolic fraction of homogenates of the left ventricle had increased nearly tenfold but returned to normal after 10 min. When dibutyryl cAMP was substituted for noradrenaline the same rapid increase in activity was seen, but in this case it did not return to normal after 10 min (84). This difference from noradrenaline perfusion, whose functions are largely mediated by CAMP, was interpreted as an overcoming of phosphodiesterase activities as a result of continuous perfusion of CAMP.The rapidity of this response along with the well-known protein phosphorylations mediated by cAMP led the authors to postulate that a protein kinase activates poly(A) polymerase in the heart. Evidence for the effects of phosphorylation on poly(A) polymerases in vitro has been reported (85,86). Incubation of the purified rat liver poly(A) polymerase with protein kinases enhanced the reaction rates for poly(A) synthesis, presumably by abolishing the characteristc lag in the reaction rate rather than by altering net poly(A) synthesis. It is not possible to fit the data from these various experimental systems into a coherent view of poly(A) polymerase regulation. In some cases it is contradictory; in others the evidence clearly argues against a simple relationship between alterations in poly(A) synthesis and poly(A) polymerase activity. The short-term in vivo regulation of poly(A) polymerase activity by a CAMP-dependent protein kinase is an interesting possibility for which some support has been provided by in vitro experiments with purified poly(A) polymerases. This is clearly an area where more detailed investigation could provide new insights into the relation of poly(A) polymerase structure to its function. 82. Morrison, M., Hall, C., Pardue, S . , Brodeur, R., Baskin, F., and Rosenberg, R. (1980). J . Neurochern. 34, 50. 83. Corti, A . , Casti, A., Reali, N . , and Caldarea, C. (1976). BBRC 71, 1125. 84. Cask A., Corti, A . , Reali, N., Mazetti, G . , Orlandini, G . , and Caldarea, C. (1977). BJ 168, 3 3 3 . 85. Rose, K., and Jacob, S. (1979). JBC 254, 10,256. 86. Rose, K., and Jacob, S. (1980). Biochemistry 19, 1472.

8. POLY(A) ADDING ENZYMES

XIV.

243

Research Applications

The properties conferred on RNA molecules by poly(A) allows them to be purified with ease from complex mixtures, and also allows them to act as templates for reverse transcriptase in the presence of a deoxythymidylate oligomer. RNA molecules can also be labeled in vifro at the 3' end by the addition of poly(A) sequences. These properties can be acquired by RNA molecules if they can serve as primers for poly(A) polymerases. The low specificity for primers along with the limited reversibility of the reaction makes them suitable reagents for this modification provided they are free of nucleases. The Mn"-activated poly(A) polymerase from calf thymus (12) is able to polyadenylate the RNA from Q p bacteriophage without destroying infectivity (18). The RNA of QP progeny does not acquire poly(A) sequences. RNA from MS-2 bacteriophage was polyadenylated with the purified E . coli polymerase (10) without apparent reduction in size (87). Studies of the migration of the 40 S ribosomal subunits during the translation of brome mosaic virus RNA (88) were clarified by labeling the 3' end of the RNA with poly(A) introduced with the Mg2+-activated poly(A) polymerase of calf thymus (11). Poly(A) polymerases have been used in several studies of the role of poly(A) sequences in the translation and lifetime of specific mRNAs. The studies use either mRNA not containing poly(A), such as those for histones, or mRNAs from which poly(A) has been enzymatically removed. In this last case, recovery of mRNA function on readenylation serves as a control for the specificity of the earlier enzymatic deadenylation. The functional lifetime of deadenylated globin mRNA in Xenopus oocytes is greatly reduced relative to native poly(A)-containing globin mRNA. Readdition of 30 to 40 AMP residues to the deadenylated globin mRNA using the E. coli polymerase prolongs its translatability to control levels (89). The functional lifetime of the nonpolyadenylated histone mRNAs in Xenopr~soocytes was greatly increased if they were polyadenylated in the same system before injection into oocytes (90). Polyadenylation in vifm has often been used to convert non-poly(A)containing RNA into a template for reverse transcriptase to obtain cDNA copies. Complementary copies were first prepared from histone mRNAs 87. Devos, R . , Gillis, E . , and Fiers, W. (1976). EJB 62, 401. 88. Kozak, M . , (1980). Cell 22, 459. 89. Huez, G., Marbaix, G., Hubert, E., Cleuter, Y., Leclerq, M., Chantrenne, H . , DeVOS, R . , Soreq, H., Nudel, U . , and Littauer, U. (1975). EJB 59, 589. 90. Huez, G., Marbaix, G., Gallwitz, D . , Weinberg, E., Devos, R . , Hubert, E., and Cleuter, Y. (1978). Nrrtrrrr (London)271, 572.

244

MARY EDMONDS

polyadenylated by the polymerase from maize (91). Globin rnRNA, deadenylated by polynucleotide phosphorylase and readenylated with the E. coli poly(A) polymerase, when used as a template for reverse transcriptase gave a cDNA indistinguishable from that prepared with native globin mRNA by hybridization analysis (92). A histone-H5 mRNA polyadenylated with maize polymerase (39) became an efficient template for reverse transcriptase (93). The ease of sequencing RNA from complementary DNA copies prompted other investigators to polyadenylate ribosomal RNA species to make them templates for reverse transcriptase. The 3' end of 18 S ribosomal RNA of several different species polyadenylated with the maize enzyme (39) were copied with reverse transcriptase in the presence of primers with appropriate nucleotides inserted at 3' side of the oligo(dT) primer (94). Similar use has been made of a poly(A) polymerase from calf thymus (95) to polyadenylate the 5 S RNA from rat liver for the preparation of cDNA with reverse transcriptase (96). Ac KNOWL EDGMENT Appreciation is extended to the National Cancer Institute of the National Institutes of Health for support of this project from research grant ROI CA 18065-5.

91. 1443. 92. 93. 94. 95. 96. 3232.

Thrall, C . , Park, W., Rashba, H., Stein, J., Mans, R., and Stein, G. (1974). BBRC 61, Getz, M., Birnie, G., and Paul J. (1974). Biochemisrry 13, 2235. Scott, A., and Wells, J. R. (1976). Nature (London) 259, 635. Haugenbuchle, O., Santer, M., and Steitz, J. A. (1978). Cell 13, 551. Keshgegian, A., Meltzer, S.,and Furth, J. (1975). Cancer Res. 35, 1141. Ackermann, S . , Keshgegian, A., Henner, D., and Furth, J. (1979). Biochemistry 15,

Capping Enzyme STEWART SHUMAN

JERARD HURWITZ

I . Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . A . Structure of the RNA Cap . . . . . . . . . . . . . . . . . . B . Cap Synthesis in Viral Systems . . . . . . . . . . . . . . . . C . Purified Capping Enzyme Systems . . . . . . . . . . . . . . I1 . Vaccinia Virus Capping Enzyme . . . . . . . . . . . . . . . . . A . Purification . . . . . . . . . . . . . . . . . . . . . . . . . B . Molecular Properties . . . . . . . . . . . . . . . . . . . . C . Reactions Catalyzed . . . . . . . . . . . . . . . . . . . . . D . Activities Associated with Vaccinia Capping Enzyme . . . . . . III . HeLa Cell Capping Enzyme . . . . . . . . . . . . . . . . . . . A . Purification, Enzyme Assay, and Molecular Properties . . . . . B . Characteristics of the HeLa Cell Capping Reaction . . . . . . . C . Cap Donor and Acceptor Specificities . . . . . . . . . . . . . D . Pyrophosphate Inhibition . . . . . . . . . . . . . . . . . . . IV. Capping Enzyme from Rat Liver Nuclei . . . . . . . . . . . . . A . Purification and Molecular Properties . . . . . . . . . . . . . B . Characteristics of the Capping Reaction . . . . . . . . . . . . V. Role of the Capping Enzyme System in Vivo . . . . . . . . . . . A . Function of the RNA Cap . . . . . . . . . . . . . . . . . . B . Relationship Between RNA Guanylylation and Guanine-7 Methylation . . . . . . . . . . . . . . . . . . . . . . . . . C . Mechanism of Transguanylylation . . . . . . . . . . . . . . . D . Relationship Between Capping and Transcription . . . . . . . . E . Control of Capping Enzyme Activity in Vivo . . . . . . . . . . VI . Research Applications of Vaccinia Virus Capping Enzyme . . . . .

246 246 246 248 249 249 249 250 254 256 256 256 256 257 257 257 258 258 258 260 261 262 264 264

245 THE ENZYMES. VOL . XV Copyright @ 1982 by Academic Ress lnc . All rights of reproduction in any form reserved . ISBN 0-12-122715-4

.

STEWART SHUMAN AND JERARD HURWITZ

1.

Introduction

A. STRUCTURE OF T H E RNA CAP Eukaryotic mRNAs contain a modified 5’ terminal “cap” structure consisting of 7-methylguanosine linked to the penultimate nucleoside of the RNA via a 5’-5‘ triphosphate bridge. The presence of this cap structure or variants thereof (see Fig. 1) on almost all eukaryotic cellular and viral mRNAs has stimulated a great deal of research on the mechanism of cap synthesis, and on the role played by the cap in mRNA function. An excellent summary of the capping literature, including an account of the discovery of the cap and of cap structure and function may be found in an article by Banerjee ( I )as well as in earlier reviews by Shatkin (2) and Filipowicz (3). In this chapter we focus on the properties of the enzymes that modify the RNA terminus to generate the cap structure m7GpppX, known as clip zero. Particular emphasis is placed on RNA guanylyltransferase (referred to as capping enzyme), the enzyme that catalyzes the guanylylation step (i.e., the formation of GpppX) in the capping pathway. The mRNA methylating enzymes are covered in detail in Chapter 18 of this volume.

B.

CAP SYNTHESIS IN V I R A L SYSTEMS

The earliest and most productive biochemical studies on RNA capping were performed using eukaryotic viruses that synthesize capped RNA in vitro, e.g., reovirus (4-a), vaccinia (9-1 I ) , cytoplasmic polyhedrosis virus (12), and vesicular stomatitis virus (13). In the case of reovirus and vaccinia, capping occurs after the initiation of transcription. In these two 1. Banerjee, A. (1980). Microbiol. Rev. 44, 175-205. 2. Shatkin, A. (1976). Cell 9, 645-653. 3. Filipowicz, W. (1978). FEBS Lett. 96, 1-11. 4. Furuichi, Y., Morgan, M., Muthukrishnan, S., and Shatkin, A. (1975). PNAS 72, 362-366. 5 . Furuichi, Y., Muthukrishnan, S., Tomasz, J., and Shatkin, A. (1976). JBC 251, 50435053. 6. Furuichi, Y.,and Shatkin, A. (1977). Virology 77, 566-578. 7. Faust, M., Hastings, K., and Millward, S. (1975). Nircleic Acids Res. 2, 1329-1343. 8. Furuichi, Y., Muthukrishnan, S., and Shatkin, A. (1975). PNAS 72, 742-745. 9. Wei, C., and Moss, B. (1975). PNAS 72, 318-322. 10. MOSS,B., Gershowitz, A., Wei, C., and Boone, R. (1976). Virology 72, 341-351. 11. Ensinger, M., Martin, S . Paoletti, E., and Moss, B. (1975). PNAS 72, 2525-2529. 12. Furuichi, Y.,and Miura, K. (1975). Nrrtrrre (Londuri) 253, 374-375. 13. Abraham, G., Rhodes, D., and Banerjee, A. (1975). Cell 5, 51-58.

9. CAPPING ENZYME 0

247

CH,

H2N

Base,(CH,A)

I 0

FIG. 1. Structure of the 5' RNA cap. The basic cap structure common to all capped mRNAs is m7GpppX. When no additional 5' modifications are present this structure is called m p zero. Cap 1 structures (m'GpppXm) have a methyl group at the 2' position of the penultimate nucleoside sugar. When the cap I penultimate nucleoside is adenosine, the adenine base can be methylated at the N-6 position (m'Gpppm6Am). Cap 2 structures (m7GpppXmpYm)are methylated at the 2' position of both the penultimate nucleoside and the adjacent nucleotide sugar. The relative abundance of cap 0, cap 1, and cap 2 RNA termini varies from one species to another. In general, the complexity of the 5' modification increases in going from lower to higher eukaryotic organisms. In higher eukaryotes, the penultimate base can be any one of the common four or m6A.

systems, the 5' end of the primary transcript, presumed to be a triphosphate terminus (f4),is modified in the following sequence of reactions (1-3). PPPXPYPZP-

+

-

PPXPYPZP-

+

(1)

PI

GPPP + PPXPYPZP- + GPPPXPYPZP- + PP, GpppXpYpZp- + S-adenosylmethionine

m'GpppXpYpZp-

(2)

+ S-adenosylhomocysteine (3)

In the first reaction, the terminal phosphate of triphosphate-terminated RNA is cleaved by RNA triphosphatase. The diphosphate-terminated RNA is then guanylylated (or capped) by RNA guanylyltransferase, an enzyme that transfers GMP from GTP to the diphosphate terminus of 14. Maitra, U., and Hurwitz, J. (1965). /"AS

54. 815-822.

248

STEWART SHUMAN AND JERARD HURWITZ

RNA, with release of PPi. The capped RNA is then methylated by RNA (guanine-7)-methyltransferase, which transfers a methyl group from S- adenosylmethionine to the unmethylated RNA cap. The virusassociated capping systems contain an additional activity-mRNA (nucleoside-2)-methyltransferase-that adds a methyl group at the 2 ' - 0 position of the ribose sugar on the penultimate nucleoside, as shown in reaction (4),to yield a cap 1 structure (as defined in Fig. 1). m'GpppXpYpZp-

+ S-adenosylmethionine

+

m'GpppXmpYpZp+ 5'-adenosylhomocysteine

(4)

It is likely that a similar mechanism of capping of primary transcripts is operative in the cell nucleus. In viral and cellular systems, all of the 5' modifications in reactions (1-4) appear to occur on nascent RNA chains (5, 11, 15, 16).

C. P U R I F I ECAPPING D ENZYME SYSTEMS Efforts to study the capping and methylating enzymes in detail, in the case of many viral and cellular systems, have been limited by the difficulty in purifying the enzymes from virus cores or from nuclear extracts. In the case of vaccinia virus, however, the capping and methylating enzymes have been solubilized and obtained in pure form. The vaccinia capping and methylating system consists of two separable components. The first is a multifunctional capping enzyme complex that contains RNA triphosphatase, RNA guanylyltransferase and RNA (guanine-7)-methyltransferase activities(17-22). This enzyme catalyzes reactions, ( / - 3 ) individually or in concert. The properties of the vaccinia capping enzymes complex are discussed in detail in Section 11. The second component (not discussed here) is an mRNA (nucleoside-2)-methyltransferase( 2 3 , 2 4 )that catalyzes reaction (4).Cellular enzymes that catalyze the capping and methylation 15. Salditt-Georgieff, M., Harpold, M., Chen-Kiang, S., and Darnell, J. (1980). Cell 19, 69-78. 16. Babich, A., Nevins, J . , and Darnell, J. (1980). Norrtre (London) 287, 246-248. 17. Martin, S . , Paoletti, E., and Moss, B. (1975). JBC 250, 9322-9329. 18. Martin, S . , and Moss, B. (1975). JBC 250, 9330-9335. 19. Martin, S . , and Moss, B . (1976). JBC 251, 7313-7321. 20. Venkatesan, S., Gershowitz, A., and Moss, B. (1980). JBC 255, 903-908. 21. Monroy, G., Spencer, E., and Hurwitz, J. (1978). JBC 253,4481-4489. 22. Shuman, S., Surks, M., Furneaux, H . , and Hurwitz, J. (1980). JBC 255, 1158811598. 23. Barbosa, E . , and Moss, B. (1978). JBC 253, 7692-7697. 24. Barbosa, E., and Moss, B. (1978). JBC 253, 7678-7702.

9. CAPPING ENZYME

249

reactions have been purified to varying extents (25-31) and the capping and methylating activities have been separated from each other. The properties of the purified guanylyltransferases from HeLa cell nuclei and from rat liver nuclei are discussed in Sections I11 and IV, respectively.

II. Vaccinia Virus Capping Enzyme

A. PURIFICATION Vaccinia, a member of the poxvirus group, is an ideal system forin vitro studies of RNA processing by virtue of the fact that the virus particle contains all the enzymes necessary for the synthesis and proper processing of functional mRNA (32). The vaccinia capping enzyme [RNA guanylyltransferase and RNA (guanine-7)-methyltransferase] was purified by Martinet (11. in 1975 (17). In the original purification procedure the enzyme was solubilized from viral cores by detergent treatment and purified further by column chromatography. More recent methods (20,22) have retained the initial steps of Martin et al., but have relied upon different ion exchange and affinity chromatography techniques. Initial studies of the vaccinia enzyme demonstrated that the purified enzyme possesses both guanylyltransferase and 7-methyltransferase activities (17, 18,20). It was subsequently shown that the purified enzyme also contains an RNA triphosphatase component ( 2 0 , 2 2 ) .These three activities are inseparable during purification. A summary of the enzymatic reactions catalyzed by the vaccinia capping enzyme complex is shown in Table I. Of these activities, the GTP-PPi exchange activity (22) affords the simplest and least expensive method of assaying capping enzyme during enzyme purification.

B. MOLECULAR PROPERTIES The capping enzyme complex has a sedimentation coefficient of 6.5 S, as determined by centrifugation through a 15-35% linear glycerol gradient 25. 26. 27. 28. 29.

Ensinger, M . , and Moss, B. (1976). JBC 251, 5283-5291. Keith, J., Ensinger, M . , and Moss, B. (1978). JBC 255, 2835-2842. Laycock, D. (1976). F P 36, 770. Venkatesan, S . , Gershowitz, A . , and Moss, B. (1980). JBC 255, 2829-2834. Venkatesan, S . , and Moss, B. (1980). JBC 255, 2835-2845. 30. Mizumoto, K . , and Lipmann, F. (1979). P N A S 76, 4961-4965. 31. Gemershausen, J . , Goodman, D . , and Somberg, E. (1978). BBRC 82, 871-878. 32. Moss, B. (1978). In “Molecular Biology of Animal Viruses” (D. P. Nayak, ed.), Vol. 2, p. 849. Dekker, New York.

250

STEWART SHUMAN AND JERARD HURWITZ TABLE I ACTIVITIES ASSOCIATED WITH VACCINIA CAPPING ENZYME COMPLEX Activity

Reaction catalyzed

( 1 ) GTP-RNA guanylyltransferase (2) RNA (guanine-7)methyltransferase (3) RNA triphosphatase (4) GTP-PP, exchange (5) Nucleoside triphosphate phosphohydrolase

GTP + ppRNA + GpppRNA + PPi GpppRNA + SAM + m'GpppRNA + SAH pppRNA --* ppRNA + Pi GTP + Is2P1PP,+ [P,y 32PlGTP NTP --* NDP + Pi [NTP= dATP, ATP, dGTP, or GTP]

containing 1 M NaCl (22). A value of 6.0 S was observed using a 5-20% linear sucrose gradient containing 0.25 M NaCl (17). Using the latter sedimentation coefficient, and a Stokes radius of 5.03 nm (determined by Sephadex G-200 gel filtration), Martin et al. (17) calculated a native rnolecular weight bf 127,000. The capping enzyme is composed of two subunits of 95,800 and 26,400 molecular weight, as determined by SDSpolyacrylamide gel electrophoresis (22). The 95,000 dalton subunit contains the active site of the guanylyltransferase and GTP-PPi exchange activities (33). It is not clear, however, whether the 26,000 dalton subunit is required for transguanylylation, nor is it clear where the functional domains of the methyltransferase and the RNA triphosphatase activities are located. Reconstitution of the native enzyme from the isolated subunits has not been reported. C. REACTIONS CATALYZED 1. RNA Guanylyltransferase

a . Assay. Guanylyltransferase catalyzes the transfer of GMP from GTP to the 5' terminus of RNA to form the cap structure GpppX. Polyadenylic acid (with triphosphate or diphosphate termini) is generally used as the RNA cap acceptor. Activity is determined by the incorporation of labeled GMP (from [3H]GTP or [C~-~~P]GTP) into acid-insoluble material, b. Requirements for Activity. Guanylyltransferase activity is dependent on a divalent cation and an appropriate cap acceptor (see Section II,C,d below). The metal ion requirement is satisfied by MgZt and to a lesser extent by Mn*+,but not by Ca2+or Zn2+.Enzyme activity is optimal at pH 33. Shurnan, S . , and Hurwitz, J. (1981). PNAS 78, 187-191.

9. CAPPING ENZYME

25 1

7.8 (Tris-HC1buffer). S -Adenosylmethionine is not required for RNA capping, although guanylyltransferase activity may be stimulated up to twofold by added S-adenosylmethionine (18). The basis for the stimulation is discussed in Section V,B. c . Cup Donor SpeciJicity. Only guanine nucleoside triphosphates serve as donors in the capping reaction. GTP, dGTP, and GTPyS can serve as substrates (19, 22). The K , for GTP is 15 pM (34).7-Methyl GTP cannot act as a cap donor (19) nor can ATP, CTP, UTP, GDP, or GMP. The inactivity with 7-methyl GTP confirms that guanine methylation occurs subsequent to RNA capping, as indicated in Section 1,B. d. RNA Acceptor Specificity. Guanylyltransferase caps RNAs that contain a 5'-triphosphate or diphosphate end (20). RNAs that contain 5'monophosphate or 5'-hydroxyl termini are not utilized in the capping reaction (18). Kinetic studies of y-phosphate cleavage of RNA versus RNA capping indicate that the diphosphate RNA terminus is the true cap acceptor, and that the capping of triphosphate-terminated RNA is facilitated by prior conversion of the RNA to a diphosphate-terminated RNA by the RNA triphosphatase activity (20, 22). There is no apparent base specificity for the penultimate nucleotide, since a variety of synthetic homoribopolymers and naturally occurring mRNAs are effective substrates, provided the phosphorylation state of the 5' end is suitable. The effect of RNA secondary structure on the efficiency of the guanylyltransferase reaction has not been adequately evaluated. Capping is not restricted to ribonucleotide polymers because the dinucleotides pppGpC, pppApG, ppGpC, and ppApG act as cap acceptors in the guanylyltransferase reaction, although the K m for dinucleotides is substantially higher than that for long RNA chains (19, 20). The effect of incremental increases in RNA chain length on the efficiency of the capping reaction has not been adequately documented. Martin and Moss (19) have shown that mononucleoside diphosphates (XDPs) can be capped to form GpppX, but that the K , for XDPs is higher still than that for dinucleotides. Thus, the condensation of GTP with XDPs is not likely to be a significant reaction in vivo. e. Inhibition of Guanylyltransferase Activity. Capping activity is inhibited by NaCl concentrations in excess of 0.1 M . At 0.3 M NaCl the enzyme is inhibited by 90%. Guanylyltransferase is completely inactivated by incubation for 5 min at 50". The enzyme is extremely sensitive to inhibition by PPi, a product of the capping reaction. In the presence of 9 p M GTP donor, capping activity is inhibited 50% by 4 p M PPi and 97% by 100 pM PPi (18). Phosphate, on the other hand, has no effect on the rate of 34. Monroy, G . , Spencer, E., and Hunvitz, J. (1978). JBC 253, 4490-4498.

252

STEWART SHUMAN AND JERARD HURWITZ

reaction at concentrations up to 20 mM Pi; activity is inhibited 55% by 40 mM Pi (22). The basis for the inhibition by PPi is discussed in Section II,C,3. f. Reversal of the Guanylylation Reaction. The capping reaction is readily reversible. Capping enzyme catalyzes pyrophosphorolysis of capped The pyrophosphorolysis RNA (GbppXpYpZ-) to regenerate [ CX-~~P]GTP. reaction requires Mg2+.N-7 methylation of the blocking guanosine moiety prevents pyrophosphorolysis of capped RNA (18). 2 . GTP-PP, Exchange a . Assay. Vaccinia guanylyltransferase catalyzes a [32P]PPiexchange reaction with GTP in the absence of an RNA cap acceptor (22). The GTP-PP, exchange activity is assayed by the incorporation of [32P]PPiinto material that is acid-soluble and adsorbable to Norit charcoal. The product of the exchange reaction is [32PlGTP. b. Requirements for the PPt Exchange Reaction. The PPi exchange reaction requires Mg2+.The divalent cation requirement is not satisfied by Mnz+,Cu2+,Ca2+,Co2+,or Zn2+.No exchange activity is observed in the absence of GTP. dGTP supports the PPi exchange reaction but it is approximately 10% as effective as GTP. ATP, CTP, UTP, ITP, GDP, GMP, and 7-methyl GTP do not support enzyme activity. Optimum exchange activity occurs at 0.2-0.3 mM GTP when assayed in the presence of 1 mM PP,. The pH optimum is from pH 7.9 to 8.5 (Tris-HC1 buffer). Activity at pH 7.3 is 37% of the activity at pH 8.3. c. Inhibition of GTP-PP, Exchange Activity. Enzyme activity is reversibly inhibited by p- hydroxymercuribenzoate, suggesting the involvement of sulfhydryl groups in the exchange reaction. Activity is unaffected by NaCl concentrations up to 0.1 M but declines gradually as the ionic strength is increased to 1.0 M NaCl. (Inhibition of 50% occurs at 0.3 M NaCl). Neither Pi nor inorganic sulfate are inhibitory at concentrations up to 40 mM; Pi in particular has a slight stimulatory effect on enzyme activity.

3. Mechanism of mRNA Capping-The Role of a Covalent Enzyme-Guanylute Intermediate The ability of the vaccinia capping enzyme to catalyze a GTP-PPi exchange reaction in the absence of an RNA cap acceptor suggests that transguanylylation occurs via a capping enzyme-guanylate intermediate. (The involvement of free GMP as an intermediate in GTP-PPi exchange was ruled out (22) by the failure of the enzyme to catalyze [3H]GMP-GTP exchange in the presence of PPi). The existence of a covalent guanyl intermediate has been demonstrated (33) and it has been shown that the

253

9. CAPPING ENZYME

capping reaction occurs by the following sequence of at least two partial reactions ( 5 and 6). G’ppp + E

E+G + PPRNA

* E-$3

f

PP,

e E + GPPP RNA

In reaction (51, the capping enzyme reacts with GTP to form an enzyme-guanylate intermediate, with concomitant release of PP,. The intermediate consists of a GMP residue covalently linked to the 95,000 dalton capping enzyme subunit. The intermediate was demonstrated by incubation of enzyme with [ (-u-32P]GTP and the subsequent detection of a 32P-labeled95,000 dalton polypeptide by SDS gel electrophoresis. The GMP moiety is linked to the enzyme via a phosphoamide bond, as judged by the acid-labile, alkali-stable nature of the bond and by the susceptibility of the linkage to cleavage by hydroxylamine at pH 4.75. Formation of the enzyme-guanylylate complex requires GTP and a divalent cation, but does not require the presence of an RNA cap acceptor. dGTP substitutes for GTP in formation of a stable enzyme-nucleotide complex, but UTP does not participate in this reaction. This suggests that the donor specificity of the capping enzyme resides at the level of formation of the enzymenucleotide intermediate. Either magnesium or manganese (but not calcium) satisfies the metal cofactor requirement in reaction (5). A similar cofactor specificity has been described in Section II,C, 1 for the complete capping reaction. In reaction (6), the guanylylated enzyme (in the absence of GTP) catalyzes transfer of the GMP moiety to the 5‘ terminus of RNA to form a GpppX cap structure. Guanylyl transfer to RNA requires MgC12. Both partial reactions in the transguanylylation pathway are readily reversible. In the reverse of reaction (3, the isolated enzyme-GMP intermediate reacts with PPi to regenerate GTP. Thus, the formation of the E-GMP complex, with rapid dissociation of the complex by PPi, accounts for the observed GTP-PPi exchange reaction in the absence of a cap acceptor. In the reversal of partial reaction (6), capping enzyme reacts with GpppAp(Ap)n (in the absence of PP,) to form the covalent enzyme[32P]GMPcomplex. The pyrophosphorolysis reaction (i.e., reversal of the complete RNA capping reaction) is therefore a two-stage reaction consisting of GMP transfer from capped RNA to capping enzyme to form E-pG, followed by dissociation of the intermediate by PP, to form GTP. We have shown that the inability of the vaccinia enzyme to pyrophosphorylize methylated capped RNA [m7GpppAp(Ap)n]is due to a failure of the enzyme to form an E-[32P]m7GMPcomplex in the presence of capped,

254

STEWART SHUMAN AND JERARD HURWITZ

methylated poly(A). This result rules out the alternative model in which E-m7GMP complex is generated, but cannot be dissociated by PPi to form 7-methyl GTP. We infer that the inability of 7-methyl GTP to act as a cap donor or to participate in PPi exchange is similarly due to a failure to form an E-m7GMP complex. From these studies, it seems that the association of guanylyltransferase and 7-methyltransferase activities has a functional purpose because the effect on the RNA capping reaction of concomitant methylation is to pull the reaction in the direction of cap formation. The applicability of the vaccinia transguanylylation mechanism to other capping systems is discussed in Section V. WITH VACCINIA CAPPING ASSOCIATED D. ACTIVITIES ENZYME

1. RNA (Gi4unine-7)-Metliyltr~insferu.~e u. Assay. Methyltransferase activity is assayed by the transfer of 3H methyl group from [3H-CH3]S-adenosylmethionineto an appropriate acceptor. Initial studies of this activity were performed using unmethylated, capped RNA (synthesized in vitro) as the acceptor. A far more convenient assay has been introduced (23) that utilizes GTP (at high concentrations) as the methyl acceptor. The methylated nucleotide or RNA is separated from S-adenosylmethionine by adsorption to DEAE-cellulose (DES 1) filters. b. Characteristics of the Methyltransferase Reaction. With capped RNA substrate, the enzyme methylates only the 5’-guanine nucleoside of the cap and does so exclusively at the N-7 position. The methyltransferase requires neither Mg2+nor GTP, and is optimally active over a broad pH range near neutrality. Enzyme activity is inhibited by S-adenosylhomocysteine, a product of the reaction. Steady-state kinetic analysis indicates that the methyltransferase reaction procedes via a sequential mechanism (18). In addition to capped RNAs, the enzyme can also methylate GpppX, GTP, dGTP, GDP, GMP, guanosine, and ITP. The methylation of these substrates requires much higher concentrations of acceptor (K,for GTP = 0.53 mM, K, for GpppG = 0.12 mM) than the methylation of capped RNA (K, for Gppp(A)n = 0.21 pM).The effect of incremental increases in RNA chain length on the methylation reaction has not been examined. Within the guanosine nucleotide family, methyl acceptor activity seems to depend on the number of 5’-phosphates, such that GTP > GDP > GMP > guanosine in acceptor activity. ATP, CTP, UTP, and xanthosine triphos-

9. CAPPING ENZYME

255

phate do not support methylation activity. The enzyme, although apparently specific for guanine-containing compounds, does not methylate 2'-GMP or 3'-GMP, thus accounting for the lack of internal 7-methyl guanine in vaccinia RNA (19).

2. RNA Triphosphatase a. Assay. RNA triphosphatase is assayed by the liberation of 32Pifrom y3'P-labeled triphosphate-terminated poly(A). 32Piis resolved from the labeled RNA by thin-layer chromatography on polyethyleneimine cellulose. b. Characteristics of the RNA Triphosphatase Reaction. The cleavage of the y-phosphate of triphosphate-terminated poly(A) requires MgC12. Mn2+ activates the enzyme, although only 12% as well as Mg2+. The RNA triphosphatase is reversibly inhibited by p- hydroxymercuribenzoate, suggesting a requirement for sulfhydryl groups. Activity is inhibited by PPi (98% inhibition at 2.5 mM Pi). Pi inhibits the enzyme at higher concentrations (50% inhibition at 20 mM Pi). The K , of the RNA triphosphatase for 5'-triphosphate-terminatedpoly(A) is approximately 6 x lo-' M with respect to 5' ends (22). The RNA triphosphatase associated with the capping enzyme is probably identical to the RNA triphosphatase purified from vaccinia cores by Tutas and Paoletti (35).These authors found that the enzyme is active on both 5'-ATP- and 5'-GTP-terminated RNAs, a result consistent with previous studies showing that capped vaccinia mRNA contains A and G exclusively at the 5' penultimate base. c. Cleavage of Nrrcleoside Triphosphates. The vaccinia capping enzyme complex catalyzes cleavage of ATP to ADP and Pi (22). This enzymatic activity requires a divalent cation (Mg, Mn, or Co) and has an alkaline pH optimum. ATP hydrolysis is unaffected by the presence of DNA or RNA, thus distinguishing this activity from the two nucleicacid-dependent nucleoside triphosphate phosphohydrolases from vaccinia described by Paoletti et al. (36, 37). The enzyme shows a strong preference for purine nucleotides. ATP, dATP, GTP, and dGTP are cleaved readily, whereas CTP, UTP, and dTTP are not. The K, for ATP is 0.8 mM, a value three orders of magnitude higher than the K , for triphosphate-terminated poly(A). It appears then that the y-phosphate cleaving activity of the capping enzyme is concerned primarily with the metabolism of 5'-RNA termini and not with the cleavage of free nucleotides. 35. Tutas, D., and Paoletti, E. (1977). JBC 252, 3092-3098. 36. Paoletti, E., Rosemond-Hornbeak, R., and Moss, B. (1974). JBC 249, 3273-3280. 37. Paoletti, E., and Moss, B. (1974). JBC 249, 3281-3286.

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111.

HeLa Cell Capping Enzyme

A.

A N D MOLECULAR PURIFICATION, ENZYME ASSAY, PROPERTIES

RNA guanylyltransferase has been isolated from HeLa cell nuclei by Venkatesan et cil. (28, 29). Nuclear extracts were found to contain sixfold more soluble capping enzyme activity than cytoplasmic extracts. A 1000fold enzyme purification from nuclear extracts was achieved by successive ion exchange and affinity chromatography procedures. Poly(A) was used as the cap acceptor, and [cI-~*P]GTPwas the cap donor in the enzyme assay. Activity was quantitated by formation of cap structures resistant to nuclease P-1 and alkaline phosphatase digestion. Product analysis of the resistant material by paper electrophoresis was performed for each assay. Although tedious, this procedure affords a highly specific capping assay, which was reportedly essential during early purification steps. The HeLa guanylyltransferase has a sedimentation coefficient of 3.5, determined by using 5-20% linear sucrose gradients that contain 50 mM NaCI. The estimated native molecular weight is 48,500, a value considerably smaller than the molecular weight of the vaccinia capping enzyme.

B. CHARACTERISTICS OF T H E HELACAPPING REACTION Like the vaccinia enzyme, the HeLa guanylyltransferase catalyzes the transfer of GMP from GTP to the 5' terminus of RNA to form GpppX- cap structures. The HeLa enzyme does not, however, have associated RNA triphosphatase and RNA (guanine-7-)-methyltransferase activities. The latter two enzymes, detectable in nuclear extracts, are separated from guanylyltransferase during purification (28). The HeLa cell RNA (guanine-7-)-methyltransferasehas been partially purified by Ensinger and Moss (25). The purified guanylytransferase requires a divalent cation. Optimal enzyme activity occurs in the presence of 2 mM MnCI2. MgClz is less effective (-25%) in supporting capping activity. The pH optimum is 7.5. Enzyme activity is inhibited by the sulfhydryl binding agent N-ethylmaleimide and by elevated ionic strength, but is unaffected by S-adenosylmethionine or S-adenosylhomocysteine.

c.

CAP DONORA N D ACCEPTOR SPECIFICITIES

The substrate requirements of the purified HeLa capping enzyme were determined by Venkatesan and Moss (29). GTP (K, = 1.1 p M ) and ITP were effective cap donors, yet ATP, CTP, UTP, GDP, dGTP, and 7-methyl GTP were not utilized by the enzyme. The failure to utilize dGTP further

257

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distinguishes the HeLa capping enzyme from the vaccinia enzyme, and indicates that the HeLa enzyme recognizes the sugar residue in addition to the base and the number of 5’ phosphates. Diphosphate-terminated RNAs (including RNAs with 5’-pyrimidine diphosphate termini), are effective cap acceptors. Unlike the vaccinia enzyme, which caps di- and triphosphate-terminated poly(A) with equal facility, the HeLa enzyme caps the diphosphate-terminated poly(A) at least fourfold more efficiently than triphosphate-terminated poly(A). The relative ability to cap triphosphate termini decreases with enzyme purification, concomitant with the elimination of RNA triphosphatase activity from the capping enzyme preparation. The apparent Km for diphosphateterminated poly(A) with an average chain length of 2000 nucleotides is 19 p M with respect to ends. 5’-Phosphate-terminated poly(A) is not capped by the enzyme. The HeLa enzyme is able to cap the diphosphate-terminated dinucleotides ppApGp and ppGpC, although the apparent Km for the dinucleotides (K, for ppApGp = 285 nM) is higher than that for poly(A). Triphosphate-terminated dinucleotides are capped 10-20% as well as their diphosphate-terminated counterparts at equal concentration. Capping of the triphosphate-terminated dinucleotides (like that of pppRNA) is apparently facilitated by residual RNA triphosphatase, which is present in even the most purified enzyme fraction. ADP is not an acceptor in the capping reaction.

D. PYROPHOSPHATE INHIBITION Pyrophosphate, a presumptive reaction product, inhibits capping by 50% at 1.5 p M PPi. The reaction is totally inhibited at 50 ~ L MPPi. Pi, on

the other hand, is not inhibitory at these low concentrations. It is presumed (although it has not been shown directly) that PPi is a product in the capping reaction mediated by the HeLa enzyme. Attempts to demonstrate pyrophosphorolysis of capped poly(A) by the HeLa enzyme were unsuccessful, suggesting that the capping reaction in HeLa cells is irreversible. This property is unique to the HeLa enzyme, since reversal of the capping reaction occurs with the purified vaccinia enzyme and with the reovirus core-associated guanylyltransferase ( 5 ) . IV.

Capping Enzyme from Rat liver Nuclei

A.

PURIFICATION A N D

MOLECULAR PROPERTIES

RNA guanylyltransferase activity has been isolated from rat liver nuclei by Mizumoto and Lipmann (30). Rat liver nuclear extracts contain both

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capping activity and 7-methyltransferase activity. The guanylyltransferase was purified 350-fold from the nuclear extract and was completely resolved from the 7-methyltransferase. Thus, the rat enzyme resembles the HeLa enzyme in the lack of a tight association between capping and methylating activities. A native molecular weight of 65,000 was estimated for the guanylyltransferase by Sephadex G-150 chromatography. (The 7-methyltransferase was estimated to be of 130,000 molecular weight.) OF T H E CAPPING REACTION B. CHARACTERISTICS

The rat liver enzyme, like that of vaccinia and HeLa cells, incorporated GMP from GTP into the RNA cap. The conclusion that the p- and y-phosphate residues of GTP are displaced as PPi is supported by the demonstration that the partially purified enzyme catalyzes a GTP-32PPi exchange reaction in the absence of an RNA cap acceptor. The RNA capping reaction requires dithiothreitol and a divalent cation. The reaction is twice as fast in the presence of Mn2+as in the presence of Mg2+at optimal concentrations. Only RNAs with 5'-di- or triphosphate termini are capped; 5'-monophosphate- or 5'-hydroxyl-terminated RNAs are inactive. The trinucleotides ppGpCpC and pppGpCpC are capped by the enzyme at equal rates at 3T, but at 30" the diphosphate-terminated derivative is more rapidly guanylylated. Mizumoto and Lipmann proposed that the capping of triphosphate-terminated substrates may be facilitated by a contaminating RNA triphosphatase activity, but no attempt was made to verify this hypothesis. V.

A.

Role of the Capping Enzyme System In Vivo

FUNCTION OF T H E RNA CAP

The synthesis of the modified RNA cap structure in civo is of great import insofar as the RNA cap itself plays a significant role in cellular metabolism. It is now well-established from in cdtro studies that the presence of the blocking 5 ' , 7-methylguanosine residue facilitates the translation of messenger RNA (38, 39, 1-3). The effect of the cap translation is manifest at the level of initiation, i.e., the cap stimulates the binding of mRNA to eukaryotic ribosomes (40). The preferential recognition of 38. Both, G., Banerjee, A , , and Shatkin, A. (1975). P N A S 72, 1189-1193. 39. Muthukrishnan, S . , Both, G . Furuichi, Y., and Shatkin, A. (1975). Nrrtrrre (London) 255, 33-37. 40. Both, G., Furuichi, Y., Muthukrishnan, S., and Shatkin, A. (1978). Cell 6, 185-195.

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capped RNA in initiation complex formation depends on certain structural features of the cap (41-43). In particular, the N-7 methyl group of guanine and the 5'-bridge phosphate residues appear to be involved. The effect of N-7 substitution is to introduce a positive charge in the base. Apparently it is the introduction of this charge, and not the presence of a methyl group per se, that contributes to cap function, since N-7 ethylation equally enhances the ability of capped RNA to form the initiation complex (44). Sonnenberg and co-workers (45-49) have recently identified a new class of protein(s)--called cap binding proteins (CBP)-that seem to play a role in cap-dependent translation. CBP specifically promote the translation of capped RNAs. This effect occurs at the level of ribosome binding of mRNA, apparently via interaction between CBP and the RNA cap. CBP seem not to correspond to any of the previously characterized protein synthesis initiation factors, although CBP activity may remain associated with certain initiation factors (eIF-4B and eIF-3) during purification of these factors. In addition to its effect on the initiation of protein synthesis, the RNA cap appears to confer increased stability on messenger RNA in vivo (50, 5 1 ) . This effect may be due to protection against a cellular 5'-exonuclease activity. A novel role for the mRNA cap in influenza virus transcription has been discovered by Krug and co-workers (52--55). They found that RNA synthesis by the influenza virion-associated transcription system is stimulated by exogeneous capped RNAs. Furthermore, the RNAs synthesized in 41. Canaani, D., Revel, M., and Groner, Y . (1976). FEES Left. 64, 326-331. 42. Hickey, E., Weber, L., Baglioni, C . , Kim, C . , and Sarma, R . (1977). J M B 109, 173-183. 43. Adarns, B . . Morgan, M . , Muthukrishnan, S . , Hecht, S . , and Shatkin, A. (1978).JBC 253, 2589-2595. 44. Furuichi, Y., Morgan, M . , and Shatkin, A. (1979).JBC 254, 6732-6738. 45. Sonenberg, N . , and Shatkin, A. (1977). PNAS 74, 4288-4292. 46. Sonenberg, N . , Morgan, M . , Merrick, W., and Shatkin, A . (1978). PNAS 75, 48434847. 47. Sonenberg, N . , Morgan, M . , Testa, D., Colonns, R., and Shatkin, A. (1979). Nircleic Acids Res. 7, 15-29. 48. Sonenberg, N . , Rupprecht, Hecht, S. , and Shatkin, A. (1979). PNAS 76,4345-4349. 49. Trachsel, H., Sonenberg, N . , Shatkin, A., Rose, J . , Leong, K . , Bergman, J . , Gordon, J . , and Baltimore, D. (1980). PNAS 77, 770-774. 50. Furuichi, Y . , LaFiandra, A . , and Shatkin, A. (1977). Ntrritre (London) 266, 235-239. 5 1 . Lockard, R . , and Lane, C. (1978). Nucleic Acids Res. 5, 3237-3247. 52. Bouloy, M . , Plotch, S . , and Krug, R . (1978). PNAS 75, 4886-4890. 53. Plotch, S., Bouloy, M . , and Krug, R. (1979). PNAS 76, 1618-1622. 54. Bouloy, M . , Morgan, M . , Shatkin, A., and Krug, R. (1979). J . Virol. 32, 895-904. 55. Robertson, H., Dickson, E., Plotch, S . , and Krug, R. (1980). Niicleic Acids Res. 8, 925-942.

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vitro contain (at the 5' end) the 5' cap and the first 12-15 nucleotides of the

exogenous RNA covalently linked to the newly synthesized plus- strand messenger RNA. Apparently, enzymes that can catalyze this unique pnming reaction are present in the virus. For this reaction to occur, the exogeneous RNA must be capped. Interestingly, not only must the RNA be guanylylated, it must be N-7 methylated at the blocking guanosine, and 2-0 methylated at the penultimate nucleotide sugar in order to function efficiently in the priming reaction (56). The influenza priming reaction is the first system in which the 2-0-methyl group has a distinct effect on RNA cap function. Note that the effector role of the cap in translational initiation in v i m is only slightly enhanced by the presence of a penultimate 2-0-methyl moiety (57, 58). B. RELATIONSHIP BETWEENRNA GUANYLYLATION AND GUANINE-7 METHYLATION RNA cap function in protein synthesis depends on both guanylylation and N-7 methylation of the RNA. It is not surprising then that capped mRNAs almost always contain a 7-methyl group in vivo (the exceptional case (59) is the developing oocyte of the tobacco hornworm, which contains capped, unmethylated mRNAs). This implies that eukaryotic cells and viruses have some <mechanismto ensure complete methylation of capped RNAs subsequent to guanylylation. Alternatively, capped but unmethylated RNAs may be rapidly degraded. Vaccinia virus achieves efficient methylation through the tight association of guanylyltransferase and methyltransferase activities in a rnultifunctional enzyme complex. From the properties of the purified enzyme, it can be seen that the association accomplishes two things: (a) It promotes efficient methylation of newly capped RNAs, since-90% of the cap product formed in a reaction containing [ CY-~*P]GTP, triphosphateterminated poly(A), and S-adenosylmethionine is m'GpppA; (b) the efficient methylation of newly capped RNAs prevents reversal (pyrophosphorolysis) of the guanylylation reaction, thereby improving the overall efficiency of the 5' modification process. The importance of this point is illustrated by the finding that RNA synthesized by vacciniain vitro in the 56. Bouloy, M., Plotch, S., and Krug, R. (1980). PNAS 77, 3952-3956. 57. Muthukrishnan, S . , Morgan, M., Banerjee, A., and Shatkin, A. (1976). Biochemistry 15, 5761-5768. 58. Muthukrishnan, S . , Moss, B . , Cooper, J., and Maxwell, E. (1978). JEC 253, 17101715. 59. Kastern, W., and Berry, S. (1976). EERC 71, 37-44.

9. CAPPING ENZYME

26 1

absence of S-adenosylmethionine contains predominantly diphosphate termini, with a lesser amount of GppX termini. However, in the presence of S-adenosylmethionine, a high proportion of the 5’-RNA termini are fully capped and methylated, while diphosphate termini are not present in significant amounts (10). Similar results have been reported for RNA synthesis directed by reovirus (60) and it has been shown that the reovirus core-associated capping enzyme catalyzes pyrophosphorolysis of GpppGpC, but not of m’GpppGpC ( 5 ) . In the two cellular systems (HeLa and rat liver) from which capping enzyme has been purified, there is no tight association of guanylyltransferase and 7-methyltransferase. However, the ability to dissociate the activities during purification does not rule out the existence of a “loosely” associated nuclear capping and methylating enzyme complex. Alternatively, the 5’ terminal-modifying enzymes may be only transiently associated at or about the time of initiation of mRNA transcription. Venkatesan and Moss (29) pointed out that the lack of intimate association between capping and methylating enzymes in HeLa cells may be correlated with the inability of the HeLa guanylyltransferase to catalyze pyrophosphorolysis of unmethylated capped RNAs. In essence, there is no “need” for methylation as a means of preventing reaction reversal, as there is in the case of vaccinia. The generality of this reasoning cannot be evaluated at this time since the reversal of the capping reaction in other cellular systems has not been examined.

c.

MECHANISM OF TRANSGUANYLYLATION

The mechanism of RNA capping by the vaccinia capping enzyme is clearly a two-stage process involving an enzyme-GMP intermediate (33). Although not demonstrated, it is likely that a similar mechanism is operative in all capping enzyme systems that catalyze transfer of GMP from GTP to RNA to form the cap. Consistent with this model, both purified rat liver guanylyltransferase (30) and reovirus cores (61) catalyze a GTP-PPi exchange reaction independent of RNA guanylylation. In all cases examined thus far (with one exception) the capping reaction involves transfer of GMP from GTP to a diphosphate terminated RNA. This acceptor specificity seemingly ensures capping of RNA termini that arise via initiation of RNA synthesis, and not of termini arising from internal RNA cleavage. Enzymes capable of phosphorylating 5’-hydroxyland 5’-monophosphate RNA termini have been purified from HeLa nuclei 60. Furuichi, Y . , and Shatkin, A. (1976). PNAS 73, 3448-3482. 61. Wachsrnan, J . , Levin, D., and Acs, G. (1976).J . Virol. 6, 563-565.

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(62) and vaccinia cores (63), respectively, but their in viva role in providing cap acceptors has not been evaluated. The exception to the predominant capping pathway is the capping reaction catalyzed by vesicular stomatitis virus (VSV, a member of the rhabdovirus group). In thein v i m VSV system, the a- and P-phosphates of the GTP cap donor are incorporated into the cap structure, indicating transfer of GDP (rather than GMP) from GTP to RNA (13). Banerjee and coworkers proposed that the cap acceptors in this system are 5’monophosphate-terminated RNAs generated either by internal RNA cleavage or by removal of the 5 ‘ 9 - and P-phosphates of newly initiated RNAs. Clarification of this interesting capping mechanism awaits further study.

D. RELATIONSHIP BETWEEN CAPPING A N D TRANSCRIPTION Studies of viral and cellular RNA synthesis suggest that modification of the 5‘ terminus of newly synthesized RNAs occurs after the initiation of transcription (i.e., after formation of the first phosphodiester bond by RNA polymerase). Capping does not depend on completion of synthesis of the transcript, but occurs on short, nascent RNA chains (5, 10, 15, 16). This temporal scheme is wholly consistent with the properties of the purified capping enzymes. Pretranscriptional capping, i.e., initiation of RNA synthesis with preformed cap fragment GpppX, may be ruled out in the case of vaccinia, HeLa, and reovirus, since these capping enzymes do not catalyze appreciable condensation of GTP with a nucleoside diphosphate to form GpppX. However in these cases, and in the case of the rat liver enzyme, there is significant capping of diphosphate-terminated dinucleotides or trinucleotides. The biochemical data are thus consistent with the capping of nascent RNA chains in viiv. The fact that newly initiated RNAs are rapidly capped and methylated suggests that RNA guanylyltransferase (and 7-methyltransferase) may be physically associated with RNA polymerase in viva, at least transiently around the time of transcriptional initiation. It is well-established that purified cellular RNA polymerase I1 (the enzyme responsible for mRNA synthesis) does not initiate accurately at promoter sites on the DNA template (64). Weil et a / . (65) have shown that RNA polymerase I1 re62. Shuman, S., and Hurwitz, J. (1979). JBC 254, 10396-10404. 63. Spencer, E., Loring, D., Hurwitz, J . , and Monroy, G . (1978). P N A S 75, 4793-4797. 64. Roeder, R. (1976). I n “RNA Polymerases” (R. Losick and M. Chamberlin, eds.), pp. 285-329. Cold Spring Harbor Laboratory, Cold Spring Harbor, New York. 65. We& P., Luse, D., Segall, J., and Roeder, R. (1979). Cdf 18, 469-484.

9. CAPPING ENZYME

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quires additional cellular factors (as yet undefined) for specific initiation in iirro. Interestingly, the accurately initiated RNAs synthesized in this and in another in virro system contain the 5'-terminal cap (65, 66). Conceiv-

ably, the capping enzyme may play a direct role in specifying proper initiation by interacting with the RNA polymerase 11, the DNA template, or both. At this time, there is no direct evidence to support a physical association of cellular capping enzyme(s) with RNA polymerase I1 in the nucleus, though this remains an attractive hypothesis. In the vaccinia system, the viral DNA-dependent RNA polymerase has been purified from vaccinia virions (67, 6 8 ) , and there is preliminary evidence suggesting that the RNA polymerase and the capping enzyme are physically associated in the virus core particle (22). The most compelling evidence for a physical and functional interaction between RNA polymerase and a 5'-modifying enzyme has arisen from studies of cytoplasmic polyhedrosis virus (CPV), a segmented doublestranded RNA virus of silkworm. Furuichi (69) has shown that in vitro transcription by CPV-associated RNA polymerase is dramatically stimulated by S-adenosylmethionine, and that the RNAs synthesized contain capped, methylated termini. This stimulation is not due to cap methylation per se since a similar effect occurs in the presence of S- adenosylhomocysteine, an inhibitor of cap methylation. The capped RNAs produced in this case contain unmethylated caps (70, 71). It appears that S-adenosylmethionine and a variety of S-adenosylmethionine analogs (72), by interacting with the CPV core-associated methyltransferase, can effect an allosteric change in the properties of the coreassociated polymerase. The K,,, of the polymerase for ATP (the initiating nucleotide in CPV RNA) is, in fact, lowered by the presence of S-adenosylmethionine (73). The functional interaction of the methyltransferase with the polymerase implies a physical association of the two enzymes as well. 66. Manley, J., Fire, A . , Cano, A., Sharp, P., and Gefter, M. (1980). PNAS 77, 38553859. 67. Spencer, E., Shuman, S . , and Hurwitz, J. (1980). JBC 255, 5388-5395. 68. Baroudy, B . and Moss, B. (1980). JBC 255, 4272-4380. 69. Furuichi, Y. (1974). N d e i c Acids Res. 1, 809-822. 70. Furuichi, Y. (1978). PNAS 75, 1086-1090. 71. Mertens, P. and Payne, C. (1978). J . Virol. 26, 832-835. 72. Wertheimer, A., Chen, S., Borchardt, R . , and Furuichi, Y. (1980). JBC 255, 59245930. 73. Furuichi, Y., and Shatkin, A. (1979).117 "Transmethylation" (E. Usdin, R. Borchardt, and C. Creveling, eds.), pp. 351-360. Elsevier/North-Holland, New York.

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OF CAPPING ENZYME ACTIVITY in Vivo E. CONTROL

No viral or cellular mutants that affect RNA guanylyltransferase activities have been isolated, making it difficult to assess the true role of the capping enzyme, and the regulatory phenomena which impinge on this enzyme. In addition, specific inhibitors of RNA guanylylation that do not affect other reactions have not been identified. Modulation of RNA guanylyltransferase activity in the cell as a function of cell growth conditions, cell cycle, or metabolic state (e.g., using inhibitors of macromolecular synthesis) have not been performed. All these areas need further invest igation. Evidence on in vivo regulation of reovirus guanylyltransferase activity has been obtained by Millward and co-workers (74-76). They found that viral mRNAs synthesized early in reovirus infection are capped, and that reovirus capping activity is detected in the infected cell. Late in infection however, viral RNAs are not capped, but contain pGpC at their 5’ ends. The absence of capped RNAs late in infection is correlated with “masking” of the guanylyltransferase and methyltransferase activities present in subviral particles. Other particle-associated activities (RNA polymerase or NTP phosphohydrolase) are not masked late in infection. The monophosphate RNA terminus present in late mRNA is generated by the action of a reovirus-induced enzyme activity that removes only the P-phosphate from ppG-terminated RNA. Significantly, these shifts in the RNA 5’-modificationpattern overlap with a reovirus-induced alteration in the cellular protein synthetic machinery that results in suppression of translation of capped RNAs while allowing efficient translation of uncapped RNA. Thus, during reovirus infection capping enzyme activity is regulated (i.e., turned off) as part of a complex strategy of host cell shutoff and viral takeover of cellular functions. VI.

Research Applications of Vaccinia Virus Capping Enzyme

Vaccinia capping enzyme is useful as a highly specific reagent for 5’ end-labeling of RNAs that contain (or can be converted into a form that contains) either 5’-triphosphate or 5’-diphosphate termini. The relatively low K , of the enzyme for GTP and RNA substrate, and the availability of [cI-~*P]GTPat high specific activity make the capping enzyme valuable in a 74. Skup, D., and Millward, S. (1980). P N A S 77, 152-156. 75. Skup, D., and Millward, S. (1980). J . Virol. 34, 490-496. 76. Zarbl, H . , Skup, D., and Millward, S. (1980). J . Virol. 34, 497-505.

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variety of analytical studies of RNA. These include studies of RNA 5’ terminal structure, sequence analysis, and transcriptional mapping. An added feature of the vaccinia enzyme is its ability to methylate the cap at the blocking guanosine with a [3H]CH3group in the presence of S- adenosylmethionine. Methylation at the 2 - 0 position of the penultimate nucleotide in addition to that of the blocking guanosine is easily achieved by using a less purified capping enzyme preparation (the DEAE-cellulose I1 enzyme fraction) that contains 2-0- methyltransferase activity. RNAs that originally contain capped termini may be decapped by periodate oxidation of the 2’, 3’-cis- diol on the blocking 7-methylguanosine followed by /3 elimination of the oxidized nucleoside with aniline (77). The product of this reaction is a triphosphate-terminated RNA that can then be recapped and methylated by the vaccinia capping enzyme. In this way, RNAs that contain cap 0, cap 1, or cap 2 structures may be radioactively labeled at the 5‘ cap and completely restored to their original capped terminal structure. By following this procedure, but omitting S-adenosylmethionine from the in vitro capping reaction, any methylated, capped RNA can be converted into its capped, unmethylated (at the N-7 position) counterpart. In another variation of this procedure, cap 0 structures can be converted to cap 1 structures using the crude vaccinia capping enzyme preparation. When this is done by prior decapping of the cap 0 RNA, the cap 1 derivative may be labeled with [w3*P]GMP in the bridge andor with [3H]methyl at both N-7 and 2 - 0 methyl positions. Alternatively, [3H]methyl labeling of only the 2 - 0 methyl position can be achieved by converting cap 0 RNA to cap 1 RNA without prior decapping. In vitro manipulation of the RNA cap structure using the vaccinia capping enzyme system has been applied to the study of cap function in initiation of protein synthesis (58, 78). The capping enzyme has been used by Krug and co-workers in determining the role of the methylated RNA cap in the priming of influenza virus transcription by exogenous RNAs (56). As the availability of the capping enzyme increases, so too should its use in future investigations of mRNA structure and function.

77. Fraenkel-Conrat, H . , and Steinschneider, A. (1967). “Methods in Enzymology,” Vol. 12, 243-246. 78. Paterson, M., and Rosenberg, M. (1979). Nature (London) 279, 692-696.

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Qp R e p l i c e THOMAS BLUMENTHAL

I. Introduction . . . . . . . . . . . . . . . 11. Purification and Properties . . . . . . . . . A. Enzyme Assay . . . . . . . . . . . . B. Purification . . . . . . . . . . . . . . C. Structure of the Enzyme . . . . . . . . 111. Catalytic Properties . . . . . . . . . . . . A. The Reactions Catalyzed . . . . . . . . B. Initiation with Heterologous Templates . C. Initiation with Homologous Template . . D. Functions of S1 and Host Factor . . . . E. Role of EF-Tu.Ts in Initiation . . . . . F. Inhibitors of Initiation . . . . . . . . . G . Elongation . . . . . . . . . . . . . . H. Termination . . . . . . . . . . . . .

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267 269 269 270 270 273 273 273 275 276 277 278 278 279

I. Introduction

The small plus-strand RNA phages that infect Escherichia coli (i.e., QP, R17, f2, MS2) contain single-stranded RNA genomes 3600-4500 nucleotides in length. Replication of viral RNA is catalyzed by enzymes called RNA replicases, assembled after infection. RNA replication is accomplished by production of a single-stranded minus strand, which is then copied by the replicase to produce replicas of the infecting viral RNA. All RNA synthesis is initiated with a 5'-GTP and proceeds in a 5' + 3' direction. The first RNA replicase to be described and studied was isolated from 267 THE ENZYMES. VOL. X V Copyright 0 1982 by Academic Press, Inc. All rights of reproduction in any form reserved. ISBN 0-1?-1?2715-4

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E. cofi infected with the Group I phage MS2 ( 1 , 2). However, the enzyme made after infection with the Group I11 phage, QP, was found to be more stable and hence was chosen for in-depth study ( 3 ) . Partially purified QP replicase preparations were found to have a strong preference for homologous RNA ( 4 ) , but they could also copy synthetic RNA templates rich in cytidylate ( 5 , 6). Furthermore, the enzyme was able to make infectious product RNA in great excess over added template (7, 8). Homogeneous QP replicase is composed of four nonidentical subunits of molecular weights 70,000, 65,000, 45,000, and 35,000 (9, 10). Only the 65,000 MW polypeptide is phage-coded. The other three, all host-coded, are taken from the protein biosynthesis apparatus. The largest is 30 S ribosomal protein S1 and the other two are the protein synthesis elongation factors Tu and Ts (1 1-13). An additional host-coded protein called host factor (HF), which is a hexamer of 12,500 MW subunits, is also required forin vitro QP RNA replication (14, 15). The replicase of a Group I RNA phage, f2,has also been purified. This enzyme contains the same three host polypeptides, although it apparently uses a different H F (16, 17). Since the host proteins were designed to function in protein biosynthesis, determination of the roles they play in phage RNA replication is obviously of great interest. Although QP replicase has been noted for its template specificity, several reports indicate that the enzyme can be used to make RNA copies of a great variety of RNAs under certain conditions (f8).Thus QP replicase is a potentially valuable tool for modern molecular biology. 1. Haruna, I . , Nozu, K., Ohtaka, Y., and Spiegelman, S. (1963). PNAS 50, 905. 2. Weissmann, C . , Simon, L . , and Ochoa, S. (1963). PNAS 49, 407. 3. Haruna, I., and Spiegelman, S. (1965). PNAS 54, 579. 4. Haruna, I . , and Spiegelman, S . (1965). PNAS 54, 1189. 5 . Eikhom, T. S., and Spiegelman, S. (1967). PNAS 57, 1833. 6. Hori, K., Eoyang, L., and Bannerjee, A. K. (1967). PNAS 57, 1790. 7. Pace, N. R . , and Spiegelman, S. (1%6). Science 153, 64. 8. Spiegelman, S., Haruna, I., Holland, I. B., Beaudreau, G . , and Mills, D. R. (1965). PNAS 54, 919. 9. Kamen, R . I. (1970). Nuture (London) 228, 527. 10. Kondo, M., Gallerani, R., and Weissmann, C. (1970). Nature (London) 228, 525. 1 1 . Inouye, H . , Pollack, Y., and Petre, J. (1974). EJB 45, 109. 12. Wahba, A. J . , Miller, M. J . , Niveleau, A . , Landers, T. A., Carmichael, G . G . , Weber, K., Hawley, D. A . , and Slobin, L. I. (1974). JBC 249, 3314. 13. Blumenthal, T., Landers, T. A . , and Weber, K. (1972). PNAS 69, 1313. 14. Franze de Fernandez, M. T., Eoyang, L., and August, J. T. (1968). Nutiire (London) 219, 588. 15. Franze de Fernandez, M. T., Hayward, W. S . , and August, J. T. (1972).JBC 247,824. 16. Federoff, N. V., and Zinder, N. (1971). PNAS 68, 1838. 17. Fedoroff, N . V., and Zinder, N. D. (1973). Nriture N e w Biol. 241, 105. 18. Blumenthal, T., and Carmichael, G. G . (1979). Annic. Rev. Biochern. 48, 525.

10.

QP REPLICASE

II.

Purification and Properties

269

A. ENZYME ASSAY

QP replicase is purified using a poly(C)-dependent poly(G) polymerase assay ( I 9). This assay measures incorporation of 3H- or 14C-labeledGTP into trichloroacetic acid-precipitable material. The assay mixture contains Mgz+(or MnZf),poly(C), GTP, and a source of QP replicase. EDTA, a sulfhydryl reducing agent, and glycerol are usually added as well. Rifampicin, DNase, and inorganic phosphate should be added during early stages in the enzyme purification to inhibit contaminating enzymatic activities. Although only three of the QP replicase subunits (11, Tu and Ts) are required for the poly(C)-dependent activity (20), enzyme containing equimolar amounts of all four subunits is in fact obtained from the purification (21). The HF is present only in small amounts in the purified enzyme. HF is thus purified separately by assay of stimulation of in vitro QP RNA replication (15). All of the subunits of QP replicase, in addition to HF, are required for in litro QP RNA replication. This activity can be monitored either by incorporation of 3H- or 14C-labelednucleotides into acid-precipitable material, or by the production of infectious RNA. The latter is measured by infection of E. coli spheroplasts. The Qp RNA-dependent assay is not used for routine purification of QP replicase because both excess QP RNA and excess HF inhibit the reaction (22). Thus the concentrations of both must be very carefully chosen and the enzyme preparation must be assayed at several concentrations to assure linearity. Furthermore, care must be taken to keep the ionic strength reasonably high to ensure that transcription of QP RNA rather than of contaminating 6 S RNAs (see below) is being measured. The number of active enzyme molecules in a preparation can be measured by incorporation of [Y-~’P]GTP.Aurintricarboxylic acid, an initiation inhibitor, can be added shortly after initiation to ensure that only first-round synthesis is assayed (23). The extinction coefficient (El,) of QP replicase has been reported to be 0.65 ( 1 3 , but results from this laboratory give a value of 1.0. If the latter value and a molecular weight of 215,000 are chosen to calculate enzyme concentration, it is found that 95% 19. 20. 21. 22. 23. 24.

Kamen, R. I. (1972). BEA 262, 88. Kamen, R . , Kondo, M . , Romer, W., and Weissmann, C. (1972). EJE 31, 44. Blumenthal, T. (1979). In “Methods in Enzymology,” Vol. 60,p. 628. Kondo, M . , and Weissmann, C. (1972). EJB 24, 530. Blumenthal, T., and Landers, T. A. (1973). BBRC 55, 680. Brown, S., and Blumenthal, T. (1976). P N A S 73, 1131.

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of the enzyme molecules initiate transcription of poly(C), whereas only 25% initiate in the in 1Jtt-o QP RNA replication assay (24). B. PURIFICATION

QP replicase is usually purified fromE. coli cells infected with QP phage containing an amber mutation late in the coat protein, QPam12 or QPamB86. These phage overproduce the enzyme because the coat protein normally serves as a repressor of translation of the replicase gene (15, 25). The standard purification utilizes a polyethylene glycol-dextran phase extraction procedure to remove nucleic acids, followed by chromatography on three successive ionic exchange columns: DEAEcellulose, phosphocellulose, and DEAE-Sephadex. If the enzyme is not sufficiently pure at that stage, sedimentation on glycerol gradients can serve as a final purification step. Upon analysis by SDS gel electrophoresis, purified enzyme preparations are found to contain equimolar amounts of the four subunits (21).

c.

STRUCTURE OF THE

ENZYME

1, Sirhirnit Ident$cation

The phage-coded polypeptide (subunit 11) has been identified as the product of the replicase (or synthetase) gene (9, 10). All of the other three subunits are present before infection. The largest polypeptide has been identified as ribosomal protein S1, while the two smaller ones are the protein synthesis elongation factors Tu and Ts. These identifications were based on identity of molecular properties, such as molecular weight and amino acid sequence, as well as on functional interchangeability with these polypeptides isolated from uninfected cells ( I 1 -13). The f 2 replicase contains three host-coded polypeptides with molecular weights identical to those of QP replicase (16);they are presumed to be the same polypeptides. A variety of experiments demonstrate that all four subunits are integral parts of QP replicase and not contaminants: (1) The four subunits co-electrophorese in nondenaturing solvents but separate into four bands when electrophoresed in the presence of urea (26); (2) all four polypeptides co-chromatograph with the RNA synthesis activity throughout the purification (21);( 3 ) S l is required for in vitro QP RNA replication (20);(4) the 25. Palmenberg, A . , and Kaesberg, P. (1973). J . Virol. 11, 603. 26. Karnen, R. I. (1975). In "RNA Phages" (N. Zinder, ed.), p. 203. Cold Spring Harbor Laboratory, Cold Spring Harbor, New York.

10. QP REPLICASE

27 1

elongation factors, acting as the EF-Tu.Ts complex (see below) are required to maintain the phage-coded polypeptide in an active conformation (27); ( 5 ) the rate of recovery of enzymatic activity during renaturation of QP replicase denatured in urea is controlled by the rate of renaturation of EF-Tu. Addition of native EF-Tu allows rapid recovery of activity (28); (6) replacement of EF-Tu in QP replicase with antibiotic-resistant EF-Tu from a mutant bacterial strain results in the formation of an unstable enzyme (29); (7) replacement of EF-Ts in QP replicase with EF-Ts from Bcrcillirs stecirotliermophilus results in the formation of QP replicase with several altered properties (30). The functions performed by the host polypeptides in RNA synthesis will be discussed in later sections. 2. Molecular Weight The sum of the molecular weights of the four subunits determined by SDS gel electrophoresis is 215,000. When the four polypeptides are covalently cross-linked by the bifunctional protein cross-linking reagent dimethylsuberimidate, a protein of approximately 215,000 molecular weight is seen on SDS gels (31). Furthermore the replicase activity elutes from sizing columns as if it had a molecular weight in this range (23).However, when QP replicase is analyzed by glycerol gradient sedimentation it behaves as if it had a molecular weight closer to 130,000. The low sedimentation value (approximately 7 S) is probably the result of an oblate shape conferred by ribosomal protein S1. An altered form of QP replicase lacking S1 also has a sedimentation constant of 7 S which is commensurate with the sum of its subunit molecular weights (145,000) (23).

3. Subunit Relationships Several lines of evidence suggest that QP replicase is composed of a relatively loose complex of two tighter subcomplexes, S1-I1 and EFTu.Ts. THe two subcomplexes appear to be bound to each other by nonionic interactions. If the enzyme is dialyzed into a low ionic strength buffer and then sedimented on glycerol gradients, it dissociates into the two subcomplexes (14). Furthermore, if QP replicase is treated with dimethyl suberimidate, covalent complexes of both the two subspecies, along with the complex of all four subunits, are found. As the ionic strength of the cross-linking mixture is increased, the amount of the large complex is increased at the expense of the two subcomplexes (31). 27. Landers, T. A., Blumenthal, T., and Weber, K. (1974). JBC 249, 5801. 28. Blumenthal, T., and Landers, T. A. (1975). Bicicliemistry 15, 422. 29. Blumenthal, T., Saari, B . , Van der Meide, P. H., and Bosch, L. (1980). JBC 255, 5300. 30. Stringfellow, L. E., and Blumenthal, T. (1982). To be submitted. 31. Young, R . A., and Blumenthal, T. (1975). JBC 250, 1829.

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The presence of RNA, particularly QP RNA, has been found to alter the relationship of the two subcomplexes. Sedimentation of QP replicase in the presence of RNA leads to release of the EF-Tu. Ts, whereas the S1-I1 complex remains bound to the RNA (10, 27). Also the presence of QP RNA completely prevents the covalent attachment of EF-Tu . Ts to S1-I1 by dimethyl suberimidate (31). However the EF-Tu Ts is not fully dissociated either in the presence of RNA (32) or during RNA synthesis, as indicated by continued sensitivity to antibodies to EF-Tu and EF-Ts following initiation (33). Thus, the following equation describes the subunit relationships of QP replicase as we currently understand them: 3 . 1 1 + Tu.Ts

Salt

S1.11.Tu.T~

RNA

where the S 1 * I1 + Tu . Ts represents a less tightly associated, rather than a fully dissociated, complex. A II*Tu.Ts complex can be isolated from QP replicase preparations. This enzyme will transcribe poly(C) but not QP RNA. Since the QP RNA replication activity can be regained by simply mixing S1 with II.Tu.Ts, S1 is not an important structural element of QP replicase (20). 4. Structurul Role of EF-Tu ' Ts

Neither S 1-11 nor EF-Tu .Ts alone has detectable RNA polymerizing activity. Furthermore if the two complexes are mixed together, only a small amount of QP replicase activity is recovered (14). However, substantial amounts of enzyme can be reconstituted if the S1-I1 is denatured in urea and then renatured in buffer containing EF-Tu-Ts (28). If the denatured S1-I1 is renatured in the absence of EF-Tu'Ts and then the EF-Tu.Ts is added, the activity is not regained (27). Thus the EF-TueTs is apparently involved in maintenance of enzyme structure. The rate of renaturation of denatured QP replicase is controlled by the rate of renaturation of EF-Tu. EF-Tu-dependent GDP binding and RNA synthetic activities are regained in parallel (28). If nondenatured EF-Tu or EF-Tu-Ts complex is added to renaturing QP replicase at the onset of renaturation, the enzymatic activity is recovered rapidly, even at low temperature, and all of the enzyme formed contains the exogenous EF-Tu (or EF-Tu-Ts) in place of the endogenous EF-Tu. This technique has been used to test the effects of a variety of alterations of the elongation factors on QP replicase reconstitution and activity. It has been found that when EF-Tu.Ts complexes, which are more stable than the endogenous 32. Blumenthal, T., Young, R. A . , and Brown, S. (1975). JBC 251, 2740. 33. Carmichael, G. G . , Landers, T. A . , and Weber, K. (1976). JBC 251, 2744.

10.

Qp REPLICASE

273

EF-Tu-Ts, are inserted by this technique large increases in both the rate and extent of reconstitution are observed. When covalently cross-linked EF-Tu-Ts or E. coli EF-Tu complexed with B. stearothermophilus EF-Ts (which form a very tight complex) are used, more than 50% of the initial QP replicase activity can be recovered (24, 30). Two additional experiments have also suggested that EF-Tu- Ts is important in stabilization of QP replicase structure. First, Sl-I1 forms large aggregates ( 2 1 1 S ) in the absence of EF-Tu . Ts (14). Second, if the EF-Tu is replaced by an antibiotic resistant mutant EF-Tu, the enzyme formed is extremely unstable even though the mutant EF-Tu is not itself more unstable than wild-type EF-Tu (29). 111.

Catalytic Properties

A. THEREACTIONS CATALYZED

QP replicase catalyzes synthesis of RNA in response to RNA templates. The enzyme initiates synthesis with GTP at or near the 3' end of the template and makes a complete complementary copy of the RNA by Watson-Crick base-pairing (34-36). An RNA primer can substitute for GTP to allow initiation (37). QP replicase has also been reported to be capable of autocatalytic RNA synthesis in the absence of template or primer, but since this reaction has not been studied in any detail it will not be considered further here (38). B. INITIATION WITH HETEROLOGOUS TEMPLATES Efficient in vitro QP RNA replication requires the presence of all four polypeptides of QP replicase as well as the host factor. However, the 11.Tu.T~complex by itself has the capability of transcribing QP RNA in addition to most other RNA species tested, but it does so at reduced efficiency (20, 39). With all templates the initiating nucleoside triphosphate is GTP. ITP, even at very high concentration, cannot substitute for GTP (37). Surprisingly, the amount of GTP required for a maximal initia34. August, J. T., Bannerjee, A. K . , Eoyang, L., Franze de Fernandez, M . T., Hori, K., Kuo, C. H., Rensing, U., and Shapiro, L. (1968). C S H S Q B 33, 73. 35. Billeter, M. A., Dahlberg, J. E., Goodman, H. M . , Hindley, J., and Weissman, C. (1969). C S H S Q E 34, 635. 36. Spiegelman, S . , Pace, N. R., Mills, D. R . , Levisohn, R., Eikhom, T. S . , Taylor, M. M . , Peterson, R . L . , and Bishop, D. H. L. (1968). C S H S Q E 33, 101. 37. Feix, G., and Hake, H. (1975). EERC 65, 503. 38. Sumper, M., and Luce, R. (1975). PNAS 72, 162. 39. Blumenthal, T., and Hill, D. (1980).JBC 255, 11713.

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THOMAS BLUMENTHAL

tion rate varies widely depending on the template (40). Templates that are selected against by QP replicase, such as f2 RNA and 16 S rRNA, require much higher GTP concentration for initiation than favored templates like synthetic polymers containing cytidylate. It has long been known that MnZ+ions reduce the template specificity of Qp replicase (4. 41). Since Mn2+also reduces the GTP initiation requirement for all templates (40), Mn2+ may reduce template specificity by allowing initiation under more stringent conditions. QP replicase may manifest a high template specificity by forming complexes with heterologous RNA species that fail to initiate because of failure to form an efficient initiation site for GTP. Mn’+ions might reduce the template specificity by forming a complex with GTP that is more efficiently incorporated than is Mg2+.GTP with unfavored templates. Monovalent cations produce an effect opposite to that produced by Mn‘+ ions. That is, they increase the GTP requirement for initiation with all templates (40). This effect is presumably a result of the tighter association between S1.11 and EF-lb.Ts at higher ionic strength as mentioned previously. Thus it appears that a looser complex between the two subcomplexes favors formation of an efficient initiation site by QP replicase and template. In the reactions discussed above it is presumed that initiation occurs at or near the 3‘ end of the template, but this has not been demonstrated. However, several experiments have shown that a stretch of C residues at (or one base away from) the 3’ end is necessary for transcription of both homologous and synthetic templates (42, 43). Transcription of oligo(C) is prevented if poly(A) is ligated to the 3’ end, but the presence of a single 3’ G residue is not inhibitory (42). Apparently any RNA molecule can be transcribed by QP replicase if oligo(C) is added to the 3’ end (42). Clearly the base sequence at the 3’ end of the template is critically important to the initiation reaction, but the precise nature of the sequence restrictions has not been elucidated. The GTP-dependent initiation reaction can be bypassed by addition of a primer as short as a dinucleotide complementary to the template ( 3 7 , 4 0 ) . In the presence of an appropriate primer, GTP can be entirely replaced by ITP, which will not substitute for GTP in initiation (37). It is not known whether this technique allows internal initiation. Since primer-dependent synthesis is very efficient with seemingly any RNA template, this is a particularly good method for producing cRNA using QP replicase. Nearly 40. 41. 42. 43.

Blumenthal, T. (1980). PNAS 77, 2601. Palmenberg, A., and Kaesberg, P. (1974). PNAS 71, 1371. Feix, G . , and Sano, H. (1975). EJB 58, 59. Kuppers, B . , and Sumper, M. (1975). PNAS 72, 2640.

10.

275

QP REPLICASE

full length copies of 9 S globin RNA have been prepared using oligo(U) or oligo(dT) as primers complementary to the 3‘ poly(A) (44, 45).

c.

INITIATION

WITH

HOMOLOGOUS TEMPLATE

QP replicase has been reported to bind tenfold more tightly to QP RNA than to nonhomologous RNA molecules (46). Furthermore, the enzyme has been shown to bind to internal regions on QP RNA, and there is some evidence that this internal binding is a prerequisite for RNA replication (47, 48). Weissmann (49) has hypothesized that the process of replication begins with binding of the replicase to two internal RNA binding sites, termed “M” and “ S . ” According to this scheme the secondary structure of the RNA causes the 3’ end of the template to be correctly positioned for initiation of RNA synthesis. Although both biochemical and electron microscopic evidence have been adduced in support of this model, it has not been unequivocally demonstrated that binding of QP replicase to the internal regions of the RNA is actually involved in replication [see Ref. (18)for a more detailed consideration of this problem]. It has been shown, however, that the penultimate C residue at the 3‘ end of QP RNA is required for efficient replication (50). In addition to QP RNA, QP replicase can replicate a variety of 6 S RNAs (51-5.3). These molecules, ranging in size from 91-220 nucleotides, are found in QP-infected cells and as contaminants of QP replicase preparations. They have unique sequences with no apparent homology to QP RNA or to each other, but all have a short stretch of C residues near the 3‘ end. These residues have been shown to be required for replication (54). Like QP RNA the 6 S RNAs contain extensive secondary structure. QP replicase binds to internal regions of these RNAs ( 5 3 , but there is no evidence that it must do so to initiate replication. 44. Feix, G. (1976). Nature (London) 259, 593. 45. Vournakis, J. N . , Carmichael, G. G., and Efstratiadis, A. (1976). BBRC 70, 774. 46. Silverman, P. M. (1973). ABB 157, 234. 47. Meyer, F., Weber, H . , and Weissmann, C. (1981). J M B , in press. 48. Vollenweider, H . J., Koller, T., Weber, H . , and Weissmann, C. (1976).JMB 101, 367. 49. Weissmann, C. (1974). FEES Leu. 43, 10. 50. Rensing, U . , and August, J. T. (1969). Nature (London) 224, 853. 51. Kacian, D. L., Mills, D. R . , Kramer, F., and Spiegelman, S. (1972). PNAS 69,3038. 52. Mills, D. R . , Kramer, F. R . , and Spiegelman, S. (1973). Science 180, 916. 53. Schaffner, W., Ruegg, K. J . , and Weissrnann, C. (1977). J M B 117, 877. 54. Mills, D. R . , Kramer, F. R., Dobkin, C . , Nishihara, T., and Cole, P. (1980). Bioclietnistry 19, 228. 55. Mills, D. R . , Kramer, F. R., Dobkin, C . , Nishihara, T., and Spiegelman, S. (1977).I n “Nucleic Acid-Protein Recognition” (H. J. Vogel, ed.), p. 533. Academic Press, New York.

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OF s1 A N D HOSTFACTOR D. FUNCTIONS

The requirement for both S1 and the host factor (HF) is specific for initiation of QP RNA replication (15, 20). 6 S RNA replication does not require these polypeptides, even though the replicase binds to internal regions of these RNA species (55). Thus it seems likely that S1 and H F perform sequence-specific functions in QP RNA replication. S1 in the absence of the other subunits has been shown to bind to the “S” site mentioned previously as well as another site close to the 3’ end of QP RNA (56,57). H F also binds to two sites on QP RNA, both A-rich. One is the site near the 3’ end to which S1 binds (56); the other is a site about 670 nucleotides from the 3‘ end, which does not correspond to any of the previously identified sites hypothesized to be important in QP RNA replication (56). However, none of the binding sites described has been directly implicated in RNA recognition by QP replicase. This apparently site-specific binding may be simply a reflection of the high affinities for homogeneous synthetic RNA polymers characteristic of these two proteins, S1 for polypyrimidines and HF for poly(A) (58, 59). The host factor is a hexamer of heat-stable 12,500 MW polypeptide (15), although recent evidence suggests it probably acts in QP RNA replication as a larger aggregate (60). It binds much more tightly to intact QP RNA than to QP RNA that has been fragmented with RNase TI. Furthermore the affinity for QP RNA is reduced in parallel with the loss of the tendency to aggregate as ionic strength is increased (60). Thus it seems likely that an H F aggregate binds to multiple sites in the folded structure of QP RNA. This binding could result in an alteration in the RNA secondary structure. Since H F causes a reduction in the requirement for GTP for initiation of transcription of QP RNA but not of other templates, it may do so by causing a specific change in the RNA secondary structure (39). These results suggest the possibility that template specificity may reside in the HF rather than in the replicase. Apparently, however, both components show specificity. It has recently been reported that the replicases produced by QP and by the closely related phage SP can efficiently replicate the 6 S RNAs from either QP- or SP-infected cells. However, in both cases, the replication of the heterologous 6 S RNA was much more easily inhibited by salt and low substrate concentration than was the replication of the homologous 6 S RNA (61). The most likely interpretation of these 56. Senear, A., and Steitz, J . A . (1975). JEC 251, 1902. 57. Goelz, S . , and Steitz, J. A. (1977). JBC 252, 5177. 58. Carmichael, G. G. (1975). JBC 250, 6160. 59. Carmichael, G. G . , Weber, K., Niveleau, A., and Wahba, A. (1975).JEC 250, 3607. 60. de Haseth, P L., and Uhlenbeck, 0. C. (1980). Biochemistry 19, 6146. 61. Fukemi, Y., and Haruna, I. (1979). Molec. Gen. Genet. 169, 173.

10.

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277

results is that the binding site for the initiating GTP is better with the homologous enzyme-template complexes than with the heterologous complexes. Since the replication of these 6 S RNA is an HF-independent reaction, these results indicate that template specificity can be expressed by the replicase alone. E. ROLEOF EF-Tu.Ts

IN

INITIATION

While it is not hard to imagine why polypeptides such as S1 and HF, which bind to single-stranded RNA, are found in an RNA replicase, it is more difficult to explain the presence of protein synthesis elongation factors in RNA synthesizing enzymes. Nevertheless two hypotheses have been advanced, both of which predict that the EF-Tu and EF-Ts would function in the initiation reaction. First, since EF-Tu binds aminoacyltRNA and since the 3' end of the phage RNA where replication initiates resembles the cloverleaf structure characteristic of tRNAs, the EF-Tu might be involved in binding the enzyme to the 3' end of the template. Second, since EF-Tu binds GTP tightly and specifically and since QP replicase uses only GTP to initiate RNA synthesis, the EF-Tu might supply the GTP binding site for initiation (13, 27). The fact that the phage-coded polypeptide is capable of elongating preinitiated polynucleotide chains at normal rates in the absence of EF-Tu and EF-Tu is consistent with an involvement of the protein synthesis elongation factors in initiation of RNA synthesis (27). The question of whether the RNA andor GTP binding functions of EF-Tu are actually utilized in the RNA replication process has been approached by using the denaturation-renaturation scheme described previously to insert modified EF-Tu in QP replicase. It has been demonstrated that contrary to expectations, EF-Tu modified by several different techniques such that it can no longer bind aminoacyl-tRNA, nevertheless functions apparently normally in QP replicase (62, 63). Furthermore EFTu * TS complex that has been cross-linked covalently with dimethyl suberimidate has been shown to substitute for EF-Tu and EF-Ts in QP replicase (24). Thus the elongation factors must act as a complex in QP replicase. Since the enzyme containing the cross-linked EF-Tu . Ts complex lacked detectable GTP binding activity characteristic of the EF-Tu in normal replicase, this experiment was interpreted as evidence that the EF-Tu GTP binding site was not used to supply the initiating GTP. However, the finding that EF-Tu .Ts from the thermophilic bacteria Thermirs thermophilus is able to bind GTP, but considerably more weakly than the 62. Brown, S . , and Blurnenthal, T. (1976). JBC 251, 2749. 63. Blumenthal, T., Douglas, J., and Smith, D. (1977). PNAS 74, 3264.

278

THOMAS BLUMENTHAL

EF-Tu ( 6 4 ) ,opens the possibility that the cross-linked E. coli EF-Tu . Ts in QP replicase could be binding the initiating GTP, but that the binding was not detected in the filter-binding assays used. Therefore the question of whether the initiation site on QP replicase is the hypothetical EF-Tu. Ts GTP binding site remains open. The fact that substitution of either the endogenous EF-Ts with EF-Ts from Bcrcilliis srerrrotherrnophilus or of the endogenous EF-Tu.Ts with cross-linked EF-Tu*Ts results in an altered Ki for competitive inhibition of initiation by GDP, circumstantially implicates the EF-Tu.Ts complex as at least a component of the initiation site (30, 65).

F. INHIBITORS O F INITIATION Polyanions such as polyethylene sulfonate (66), aurintricarboxylic acid (23) [in this case a polymeric contaminant is the actual inhibitor (67)], and poly(U) (22, 68, 6 9 ) inhibit QP replicase initiation by competing with the template for binding to the enzyme. These polymers do not inhibit the elongation of preinitiated RNA chains. GDP and ppGpp, which cannot substitute for GTP in the initiation reaction, act as competitive inhibitors of initiation but not of elongation (65). Other ligands that interact with EF-Tu, such as TPCK and kirromycin, do not inhibit QP replicase, although kirromycin does inhibit renaturation of denatured enzyme (62). G. ELONGATION In spite of the fact that QP replicase catalyzes the production of a product RNA strand wholly complementary to the template RNA strand, in the homologous reaction at least, the product behaves as a fully singlestranded molecule (70). If the enzyme is removed a double-stranded product-template molecule is formed. Thus during the elongation reaction the product and the template are maintained as single-stranded entities. Presumably both the enzyme and intrastrand hydrogen bonding serve as barriers to annealing of product and template. Multistranded structures are found both in QP-infected cells and in Arai, K . , Arai, N . , Nakamura, S., and Kaziro, Y. (1978). EJB 92, 521. Blumenthal, T. (1977). BBA 478, 201. Kondo, M . , and Weissmann, C. (1972). BBA 59, 41. Gonzalez, R . G., Blackburn, B. J., and Schleich, T. (1980). BBA 562, 534. 68. Hori, K. (1973). J B Tokyo 74, 273. 69. Kondo, M. (1976). BJ 155, 461. 70. Weissmann, C . , Feix, G . , and Slor, H . (1968). CSffSQB 33, 83.

64. 65. 66. 67.

10.

QP REPLICASE

279

replication reactions in vitro. These structures consist of a single plus or minus strand and several nascent strands (71, 72). However, these complex structures are not prerequisites for QP replicase-catalyzed RNA replication. Replication of a 6 S RNA molecule, MDV-1, can be carried out by a single enzyme molecule (73). The rate of MDV-1 elongation is variable. There are several discrete sites on the RNA where QP replicase pauses. These sites correspond to regions that have the ability to form 3’-terminal hairpin structures (74). Pausing could be due to formation of hairpin structures in either the nascent product or the portion of the template just copied, or both. QP replicase has a relatively high rate of base-pairing errors, about 1.6 transition mutations per doubling (75). This corresponds to a misreading frequency of 10-3-10-4. The lack of a proofreading function is presumably responsible for the high error rate. H. T E R M I N A T I O N Even though all RNA synthesis begins with GTP at the 5’ end, the 3’ end of all product strands in both QP RNA and 6 S RNA replication is an A residue (50, 76). This A must be added posttranscriptionally and is presumably integral to the termination process. The enzyme will not adenylate free completed plus or minus strands from which the terminal A has been removed (76).

71. 72. 73. 2038. 74. 75. 76.

Feix, G . , Slor, H., and Weissmann, C. (1967). P N A S 57, 1401. Hori, K. (1970). BBA 217, 394. Dobkin, C., Mills, D. R . , Kramer, F. R . , and Spiegelman, S . (1979). Biochemistry 18,

Mills, D. R . , Dobkin, C . , and Kramer, F. R . (1978). Cell 15, 541. Domingo, E., Sabo, D., Taniguchi, T., and Weissrnann, C. (1978). Cell 13, 735. Weber, H., and Weissmann, C. (1970). J M B 51, 215.

This Page Intentionally Left Blank

2 '.5 '.Oligoadeny late Synthetase L . ANDREW BALL I . Introd.uction . . . . . . . . . . . . . . . . . . . . . . . . . . I1 Purification and Properties . . . . . . . . . . . . . . . . . . . . A . Occurrence . . . . . . . . . . . . . . . . . . . . . . . . . B . Basal and Induced Enzyme Levels . . . . . . . . . . . . . C . Enzyme Purification . . . . . . . . . . . . . . . . . . . . . D . Molecular Weight Measurements and Polypeptide Composition . E . Properties . . . . . . . . . . . . . . . . . . . . . . . . . 111 Reactions Catalyzed . . . . . . . . . . . . . . . . . . . . . . A . Synthesis of 2' ,5 '-Oligoadenylates . . . . . . . . . . . . . . B. Nucleotidyl Donation by Nucleoside Triphosphates Other than ATP . . . . . . . . . . . . . . . . . . . . . . . C . 2'-Adenylylation of Acceptors Other than ATP and 2-5A . . . IV. Biological Role . . . . . . . . . . . . . . . . . . . . . . . . A . Nuclease Activation by 2-SA in Cell Extracts . . . . . . . . . B. Occurrence and Action of 2-SA in Whole Cells . . . . . . . . C . Effects of 2-5A Cores on Whole Cells . . . . . . . . . . . . D. Role of the 2-5A System in Interferon Action . . . . . . . . . E . Role of the 2-5A System in Non-Interferon-Treated Cells . . .

.

. .

.

.

1

. . . . . . .

281 284 284 286 287 289 289 290 290 303 303 304 305 307 309 309 312

Introduction

The series of experiments that led to the discovery of 2 ' 3 oligoadenylate synthetase originated in 1972. Studies were undertaken to investigate the mechanisms by which protein synthesis was inhibited in animal cells that had been exposed to the antiviral agent interferon . The 28 1 THE ENZYMES. VOL . XV Copyright 0 1982 by Academic Press. Inc All rights of reproduction in any form reserved . ISBN l%12-122715-4

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L. ANDREW BALL

approach was to prepare cell-free protein synthesizing systems from interferon-treated and untreated cells, and to compare their ability to translate viral messenger RNAs. The results from one research group consistently showed that interferon pretreatment alone was not sufficient to impair translational activity; it was necessary in addition to infect the cells with a virus shortly before preparation of the cell extract (I). Under these conditions, a clear interferon-mediated inhibition was observed that affected both the initiation and elongation of polypeptide chains. These results were not universal: other workers observed an interferonmediated inhibition of cell-free protein synthesis even without virus infection (2, 3 ) , but in this case too, infection accentuated the inhibition (2). How did virus infection trigger the development of the inhibition? It was proposed that the synthesis of double-stranded RNA (dsRNA) in the virus-infected cells was responsible (4). This was supported by the observation that translation catalyzed by extracts of interferon-treated cells was sensitive to inhibition by dsRNA, whereas that by extracts of untreated cells was largely insensitive (4, 5 ) . ATP was required for the inhibition to develop (6), and this observation led to the discovery of a latent protein phosphokinase, which was induced or enhanced by interferon treatment and required dsRNA for activation (7-9). This protein kinase phosphorylated a subunit of translation initiation factor 2 (eIF-2), and thus mediated at least part of the inhibition of ceII-free protein synthesis caused by dsRNA (10, 11). In addition to activation of the kinase, however, a heat-stable, low-molecular-weight inhibitor of translation was also formed when extracts of interferon-treated cells were incubated in the presence of dsRNA and ATP (7). Moreover, an enzyme preparation 1. Friedman, R. M., Metz, D. H., Esteban, R. M., Tovell, D. R., Ball, L. A,, and Kerr, I. M. (1972). J . Virol. 10, 1184- 1198. 2. Falcoff, E . , Falcoff, R., Lebleu, B., and Revel, M. (1973). J . Virol. 12, 421-430. 3. Gupta, S. L., Sopori, M. L., and Lengyel, P. (1973). BBRC 54, 777-783. 4. Kerr, 1. M., Brown, R. E., and Ball, L. A. (1974). Nature (London) 250, 57-59. 5 . Kerr, I. M., Brown, R. E., Clemens, M. J., and Gilbert, C. S. (1976). EJB 69,551-561. 6. Roberts, W. K., Clemens, M. J., and Kerr, I. M. (1976). PNAS 73, 3136-3140. 7. Roberts, W. K., Hovanessian, A,, Brown, R. E., Clemens, M. J., and Kerr, I. M. (1976). Nutiire (London) 264, 477-480. 8. Lebleu, B . , Sen, G. C., Shaila, S., Cabrer, B., and Lengyel, P. (1976). PNAS 73,

3107-31 1 1 . 9. Zilberstein, A., Federman, P., Shulman, L., and Revel, M. (1976). FEBS Lett. 68, 119- 124. 10. Kaempfer, R., Israeli, R., Rosen, H., Knoller, S., Zilberstein, A., Schmidt, A., and Revel, M. (1979). Virology 99, 170-173. 11. Revel, M., Gilboa, E., Kimchi, A., Schmidt, A., Shulman, L., Yakobson, E., and Zilberstein, A. (1977). I n “Proceedings of the Eleventh Meeting of the FEBS” (B. F. C. Clark e f ul., eds.), Vol. 43, pp. 47-48. Pergamon, New York.

I I . 2‘,5’-OLIGOADENYLATE SYNTHETASE

283

Nt12 I

Ntl,

,Q .O ;-

H

OH

0

i

.”x LN ‘i N>

OH

FIG. I .

OH

5’-O-Triphosphoryladenylyl(2‘ ,5’)adenylyl(2‘3’)adenosine.

from these extracts, immobilized by binding to a dsRNA-agarose column, was able to generate this inhibitor essentially indefinitely when perfused with ATP (12, 13). The inhibitor exerted its effects by activation of a latent endoribonuclease that degraded messenger RNA and thus arrested cellfree protein synthesis (14). Structural analysis showed that the inhibitor consisted of an oligomeric series of oligonucleotides composed of adenylate residues that were linked by 2’ ,5’-phosphodiester bonds. Their general structure is pppA2’(p5’A)n (15). The trimer and higher oligomers were active, so the prototype inhibitory oligonucleotide of this series is 5’-U-triphosphoryladenylyl(2’,5‘)adenylyl(2‘,5‘)adenosine (Fig. I). These are the first natural 12. Hovanessian, A. G . , Brown, R . E . , and Kerr, I. M. (1977). Nufitre (London) 268, 537-540. 13. K e n , I. M., Brown, R. E . , and Hovanessian, A. G. (1977). Nr~Iitre(London) 268, 540-542. 14. Clemens, M . J . , and Williams, B. R . G . (1978). Cell 13, 565-572. IS. Kerr, I. M . and Brown, R. E. (1978). P N A S 75, 256-260.

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L. ANDREW BALL

nucleotides with 2‘-5’ bonds to be discovered. In this chapter, the 2’3’oligoadenylates are referred to by the abbreviation 2-5A. Synonyms in current use in the literature are oligo-isoadenylates, 2‘ ,5’-oligo(A), (2’-5’)A, and 2’,5’A. Molecules that lack the 5’-terminal phosphates, A2’(p5’A),,are referred to here as cores. The enzyme activity responsible for 2-5A synthesis from ATP is referred to as 2‘,5’-oligoadenylate synthetase, or 2-5A synthetase. Synonyms in current use in the literature are oligo-isoadenylate synthetase, 2‘ ,5’-oligo(A) polymerase, and 2-5A polymerase. The protein kinase and 2-5A synthetase, both of which require dsRNA for activation, appear sufficient to account fully for the enhanced sensitivity of extracts of interferon-treated cells to the inhibition of protein synthesis by dsRNA. On the other hand, they are probably not sufficient to account for the multitude of biological changes that cells undergo in response to interferon treatment (16). Thus the biological role of 2-5A synthetase is an open question, and is discussed at the end of this chapter (Section IV). 11.

Purification and Properties

A. OCCURRENCE 2-5A Synthetase has been identified in several mammalian and avian cells grown in culture, and in a number of cells and tissues isolated directly from mammals and birds (Table I; see 12, 17-28). The enzyme has also been observed in two extracellular locations: the serum of interferon-treated or virus-infected mice, and in the purified virions of two enveloped viruses released from interferon-treated cells. The significance of these extracellular locations is not clear. 16. Stewart, W. E . , 11. (1979). “The Interferon System.” Springer-Verlag, Berlin and New York. 17. Stark, G. R., Dower, W. J., Schimke, R. T., Brown, R. E., and Kerr, I. M. (1979). Nature (London) 278, 471-473. 18. Ratner, L., Wiegand, R. C . , Farrell, P. J., Sen, G. C . , Cabrer, B . , and Lengyel, P. (1978). BBRC 81, 947-953. 19. Wood, J . N., and Hovanessian, A. G. (1979). Nature (London) 282, 74-76. 20. Jarvis, A . P.,White, C., Bail, A . , Gupta, S . L., Ratner, L., Sen, G. C., and Colby, C. (1978). Cell 14, 879-887. 21. Wallach, D. and Revel, M. (1980). Nature (London) 287, 68-70. 22. Baglioni, C., Maroney, P. A., and West, D. K. (1979). Biochemistry 18, 1765-1770. 23. Eppstein, D. A., Peterson, T. C . , and Samuel, C. E. (1979). Virology 98, 9-19. 24. Ball, L. A., and White, C. N . (1978). PNAS 75, 1167-1171. 25. Krishnan, I . , and Baglioni, C. (1980). Nature (London) 265, 485-488. 26. Shimizu, N., and Sokawa, Y. (1979). JBC 254, 12034-12037. 27. Sokawa, Y., Ando, T., and Ishihara, Y. (1980). Itifect. Immun. 28, 719-723. 28. Hovanessian, A. G., and Kerr, I. M. (1978). EJB 84, 149-159.

TABLE I OCCURRENCE OF

2-5A

SYNTHETASE

Source A. Cultured cells Mouse L cells Friend cells Ehrlich ascites tumor cells Embryonal carcinoma cells 3T6 fibroblasts 3T3 fibroblasts Human HeLa cells Namalva cells Diploid fibroblast (MRCS) cells Amnion U cells Monkey Kidney (Vero) cells BSC-I fibroblasts Chick Primary cultures of embryo cells B , Purified viruses Vesicular stomatitis virus, grown in BSC-1 fibroblasts Moloney murine leukemia virus grown in 3T3 fibroblasts C. Uncultured cells and animal tissues Mouse Serum and plasma Spleen and spleen lymphocytes Lungs Brain Liver Thy moc y te s Mesenteric lymph node cells Intestinal mucosa Bone marrow cells Trigeminal ganglia Rabbit Reticulocytes Guinea pig Mammary gland Chick Oviduct XenoprrA laevis Liver Pooled blood cells Drosophiln rnelonogaster Embryos Larvae Adult flies

Enzyme

+

+

+

+ + + + + +

+

+

+

+ + + + + + Very low Low

+ +

+ +

+ +

+ + -

Reference

286

L. ANDREW BALL

The biological distribution of 2-5A synthetase that has been reported overlaps exactly with the distribution of the interferon system itself (16). If this correlation is valid, it can be expected that mammals, birds, reptiles, and fish will have the enzyme, but that amphibians, invertebrates, plants, fungi, protozoa, and prokaryotes will probably not. On the other hand, the possibility exists that 2-5A synthetase plays an important role in cellular metabolism that is distinct from its role in the interferon system, and if this is the case the synthetase may predate interferon in evolution.

LEVELS B. BASALA N D INDUCED ENZYME Cell types differ greatly in their uninduced enzyme levels. For example, HeLa cells and rabbit reticulocytes are particularly rich sources of the enzyme in the absence of deliberate interferon treatment (22, 28). However, rabbit reticulocytes are routinely isolated following the induction of acute anemia by repeated injections of phenylhydrazine, and it remains a distinct possibility that the presence of the enzyme in these cells is the result of experimental manipulations rather than a natural consequence of erythropoiesis. Different lines of the same cell type can have widely different basal enzyme levels (17), and in one case elevated basal levels have been correlated with an enhanced resistance to virus infection in a variant cell line (20). Although in this instance the elevated synthetase level was apparently due to constitutive interferon production by the variant cells, this cannot be the explanation of all elevated basal levels. In a human endometrial cancer cell line, for example, the high basal level was unaffected by treating the cells with anti-interferon antiserum, indicating that in this case the spontaneous production of interferon was not responsible (29). In almost all instances that have been studied, the enzyme levels increase in response to treatment of the cells with homologous interferons. Treatment of human cells with any of the three known types of interferon, a (leukocyte), p (fibroblast), or y (immune) interferon, causes an increase in the levels of 2-5A synthetase (30, 31). Indeed, only two interferonresponsive cell types have been described in which the level of the enzyme is unaffected by interferon treatment, i.e., WBalb mouse fibroblasts 29. Verhaegen, M . , Divizia, M., Vandenbussche, P., Kuwata, T., and Content, J . (1980). P N A S 77, 4479-4483. 30. Hovanessian, A. G . , Meurs, E . , Aujean, O . , Vaquero, C . , Stefanos, S . , and Falcoff, E. (1980). Virology 104, 195-204. 3 1 . Baglioni, C.,and Maroney, P. A. (1980). Virology 101, 540-544.

11. 2’,.5’-OLIGOADENYLATE SYNTHETASE

287

(32) and human MRC5 cells (33).In other cells, the response is a function of the interferon dose, but at saturating interferon concentrations it can vary from a relatively modest 10-fold change in some cell types [e.g., HeLa cells (22)l to a dramatic 10,000-fold change in others [e.g., chick embryo cells (34)l. The reasons for these differences in the basal levels and in the inducibility of the enzyme are unclear, but they may be related to differences in the levels of the phosphodiesterase that degrades 2-5A (see Section III,A,7). Twelve to twenty-four hours are required for the 2-5A synthetase levels to respond fully to interferon treatment. The response is blocked by antiinterferon antiserum (35),and by inhibitors of cellular RNA and protein synthesis administered at the time of interferon treatment (22, 34, 35). Later administration of inhibitors of RNA synthesis can cause a modest superinduction of the enzyme (3.5, 36). Following removal of interferon from treated HeLa cells, the enzyme activity is stable for at least 3 days (22), the specific activity declining only because of cell growth and division. The synthetase level also increases with increasing confluency of cells in culture, and with decreasing estrogen levels in whole animals (17). The significance of these effects is a matter for speculation.

c.

ENZYME PURIFICATION

Procedures have been published for the purification of 2-5A synthetase from mouse L cells (37), Ehrlich ascites tumor cells (38), chick embryo cells (39, 40), and rabbit reticulocytes (41). The enzyme binds to dsRNA and can be isolated from crude cell extracts by passage through a column that contains immobilized dsRNA covalently bound to an inert support. Agarose or paper to which the synthetic double-stranded polynucleotide, 32. Hovanessian, A. G . , Meurs, E., and Montagnier, L. (1981). J . Inruf’eron Res. 1, 179- 190. 33. Meurs, E., Hovanessian, A . G., and Montagnier, L. (1981). J . 1nrer:feron Rev. 1, 2 19- 232. 34. Ball, L. A . (1979). Virology 94, 282-296. 35. Kimchi, A . , Shulman, Schmidt, A . , Chernajovsky, Y., Fraden, A , , and Revel, M . (1979). P N A S 76, 3208-3212. 36. Gupta, S. L., Rubin, B. Y., and Holmes, S. L. (1981). Virology 111, 331-340. 37. Hovanessian, A . G . , and Kerr, I . M. (1979). EJB 93, 515-526. 38. Dougherty, J. P., Samanta, H., Farrell, P. J . , and Lengyel, P. (1980). JBC 255, 3813-3816. 39. Ball, L. A. (1980). Ann. N . Y. Acrid. Sci. 350, 486-496. 40. Ball, L. A. and White, C. N . (1979). 111 “Regulation of Macromolecular Synthesis by Low Molecular Weight Mediators” (G. Koch and D. Richter, eds.), pp. 303-317. Academic Press, New York. 41. Justesen, J . , Ferbus, D., and Thang, M. N . (1980). Nucleic Acids Res. 8, 3073-3085.

288

L. ANDREW BALL

poly(riboinosinic) * poly(ribocytidy1ic acid) [poly(I) .poly(C)], has been bound has been widely used for this purpose. The enzyme can be eluted from these columns by buffer that contains KCI at concentrations greater than 0.2 M ; for the enzyme from interferon-treated mouse L cells or rabbit reticulocytes, such a procedure provides a purification of at least 1000-fold ( 2 8 , 3 7 ) .However, since the enzyme is very stable and active when bound to its affinity support, it has been widely used in the immobilized state, bound to either poly(1). poly(C)-agarose or poly(1). poly(C)-paper. The enzyme can also be immobilized by binding to 2',5'-ADP-agarose, but in this case it requires dsRNA for activation ( 4 2 ) . Hovanessian and Kerr ( 3 7 ) used repeated chromatography on DEAEcellulose columns to achieve further purification of the enzyme isolated from the postribosomal supernatant fraction of mouse L cells, and eluted from poly(1).poly(C)-agarose. Their recovery of activity was very low, but it could be significantly enhanced by recombining the synthetase with separated fractions that contained the interferon-induced kinase activity. This raised the possibility that 2-5A synthetase was a two-component enzyme and that the kinase was involved in its activation or activity. However, since no phosphorylation of polypeptides in the synthetase fraction was evident under these conditions, the significance of the second component and its apparent association with the kinase are unclear. Further doubt about the validity of the second component comes from the work of Dougherty et NI. (38) who purified 2-5A synthetase from the ribosomal wash fraction of interferon-treated Ehrlich ascites tumor (EAT) cells to a single homogeneous polypeptide. Unlike Hovanessian and Kerr, Dougherty et al. avoided the use of dsRNA-agarose affinity columns and relied instead on fractional ammonium sulfate precipitation and ion exchange chromatography to purify the enzyme. Their overall purification was 2500-fold, and the recovery of activity was 9%. The activity of the purified enzyme preparation cosedimented on a glycerol gradient and coeluted from CM-cellulose chromatography with the only polypeptide that could be detected by SDS-polyacrylamide gel electrophoresis of stained or radioiodinated enzyme. However, no evidence was presented to show that this polypeptide was synthesized in response to interferon treatment of cells. Indeed, none of the interferon-induced polypeptides that have been described in these cells has a molecular weight that matches that of the pure synthetase (43). Primary cultures of interferon-treated chick embryo cells contain 2-5A synthetase at about ten times the specific activity of cultures of 42. Johnston, M. I., Friedman, R . M . , and Torrence, P. F. (1980). Biochetnisrry 19, 5580-5585. 43. Farrell, P. J . , Broeze, R. J . , and Lengyel, P. (1979). Nofirre (London) 279, 523-525.

11. 2’3’-OLIGOADENYLATE SYNTHETASE

289

interferon-treated mouse L cells or Ehrlich ascites tumor cells (34). This facilitated attempts to identify the polypeptide(s) that constitute the chick enzyme by direct incorporation of radiolabeled amino acids during interferon treatment. Enzyme preparations purified 150- to 200-fold contained as their major labeled component a polypeptide of 56,000 apparent molecular weight that was undetectable in control preparations purified from non-interferon-treated cells (39). This polypeptide bound to dsRNAagarose affinity columns, and comigrated with the enzyme activity during gel filtration (34). Moreover, it was established that the 56,000 dalton polypeptide was synthesized in response to interferon treatment, and that its synthesis, like that of the enzyme, was prevented by inhibitors of transcription. Interestingly, polypeptide synthesis occurred transiently following interferon treatment, being turned on and then shut off despite the continued presence of active interferon in the medium (34). Similar transient synthesis of interferon-induced polypeptides has been observed in other systems (36. 44).

D. MOLECULAR WEIGHT MEASUREMENTS AND POLYPEPTIDE COMPOSITION The molecular weight of 2-5A synthetase has been estimated by gel filtration of unpurified extracts of interferon-treated chick embryo cells (341, and by glycerol gradient centrifugation of highly purified enzyme from Ehrlich ascites tumor cells (38). In the former case, the enzyme activity eluted between ovalbumin and bovine serum albumin, which indicated an apparent molecular weight of 50,000-60,000. In the latter case, the enzyme activity sedimented slightly faster than bovine serum albumin, indicating a molecular weight of about 85,000. Molecular weight estimates were also derived from polypeptide analyses performed under denaturing conditions by SDS-polyacrylamide gel electrophoresis. The activity of the chick enzyme appeared to be associated with an interferon-induced polypeptide of 56,000 apparent molecular weight (see Section 11,C). On the other hand, the purified murine enzyme contained a 105,000 dalton polypeptide as its only detectable component. In both instances, therefore, the enzyme appeared to behave as a monomer, at least when it was not bound to dsRNA. E. PROPERTIES

Few physical properties of the synthetase have been reported. However, its behavior during ion exchange chromatography at neutral pH 44. Rubin, B. Y.,and Gupta, S . L. (1980). J . Viroi. 34, 446-454.

290

L. ANDREW BALL

suggests that it is a somewhat basic molecule (38, 45). Both in the crude state and bound to poly(I)*poly(C)-paper the enzyme is stable at -70" essentially indefinitely. In unpurified cell extracts, 2-5A synthetase is absolutely dependent on dsRNA for activation, and this dependence is retained by the purified enzyme (38). The activation is fully reversible by removal of the dsRNA [by ribonuclease I11 digestion (46) or DEAEcellulose chromatography (37),for example], and thus does not appear to involve a covalent modification of the enzyme. That the enzyme has some unrecognized catalytic activity in the absence of dsRNA remains a formal possibility. Ill.

Reactions Catalyzed

2-5A Synthetase is a nucleotidyltransferase that transfers nucleoside 5'-monophosphate residues from appropriate nucleoside 5'-triphosphates to the 2' position of appropriate acceptors. The prototype reaction is the synthesis of 2'3-oligoadenylates [Eq. (l)], where the sole substrate is ATP. Adenosine 5'-monophosphate residues are transferred initially to the 2' position of ATP, and on subsequent occasions to the 2' position of the growing oligoadenylate chain. The other reaction product is inorganic pyrophosphate (47). n ATP + pppA2'(p"'A),-l + ( n

-

l)PP,

(1)

Other nucleotidyl donors and acceptors have been described, and the reactions catalyzed by 2-5A synthetase can be considered in three groups: (a) the synthesis of 2' ,5'-oligoadenylates, (b) nucleotidyl donation by nucleoside triphosphates other than ATP, and (c) 2'-adenylylation of acceptors other than ATP or 2'3-oligoadenylates. Although reactions in all three groups have been detected in cell extracts, only the synthesis of 2' ,S-oligoadenylates has been observed in whole cells, so the physiological significance of reactions in groups (b) and (c) remains questionable. A. SYNTHE5IS OF 2',5'-oLICOADENYLATES 1. Strirctrrres of the Reaction Prodircts

2-5A Synthetase activity was first detected as the synthesis (by extracts of interferon-treated mouse L cells) of a heat-stable, low-molecular45. Zilberstein, A,, Kimchi, A . , Schmidt, A . , and Revel, M. (1978). P N A S 75, 47344738. 46. Minks, M . A., Benvin, S . , and Baglioni, C . (1980). JBC 255, 5031-5035. 47. Samanta, H., Dougherty, J. P., and Lengyel, P. (1980). JBC 255, 9807-9813.

11.

2’,5’-OLIGOADENYLATE SYNTHETASE

29 1

weight inhibitor of cell-free protein synthesis (7). The inhibitor was apparently synthesized u‘e nova from ATP, and could be isotopically labeled by the incorporation of [-p3‘P]ATP, [U-~*P]ATP, or [3HlATP. The major inhibitory species had a net charge of -6, and contained radioactive label from these three precursors in the ratio 1 : 3 :3. Digestion with bacterial alkaline phosphatase yielded a resistant core that had a net charge of -2 and contained radioactivity from the above three precursors in the ratio 0 :2 : 3. The inhibitor and its phosphatase-resistant core were resistant to digestion with P1, T I , T2, U2, pancreatic, spleen, and micrococcal nucleases, but sensitive to digestion with either snake venom phosphodiesterase or alkali. Digestion of the core with the former yielded 5’-AMP and adenosine, and with the latter 2‘-AMP, 3‘-AMP, and adenosine. These data and others indicated that the structure of the major component of the inhibitor was probably pppApApA, but that the internucleotide linkages were not 3’-5‘. Direct evidence for the nature of the internucleotide linkages was derived from an analysis of the products formed on sequential degradation of the inhibitor by periodate oxidation and @elimination. One cycle of these reactions, followed by phosphatase digestion, yielded A2‘p5‘A which was identified by thin-layer chromatography. A second cycle of periodate oxidation and @-elimination yielded 2‘-AMP. These results identified the 5’-proximal internucleotide linkage as 2 ‘ - 5 ’ . The other internucleotide linkage of the trimer was shown to be 2’-5‘ also, because digestion of the putative pppA2‘p5’A2’p (derived from one cycle of periodate oxidation and P-elimination of the trimer) with high levels of snake venom phosphodiesterase yielded p5’A2‘pfrom the 3‘ terminus of the molecule. These data led to the conclusion that the structure of the major component of inhibitor preparations synthesized by the mouse enzyme was 5’-O-triphosphoryladenyly1(2’-5’)adenylyl(2’5‘) adenosine (Fig. 1) (15). Also present among the reaction products were the corresponding dimer, and the tetramer, pentamer, hexamer, and higher oligomers in decreasing amounts. Similar but less extensive structural analyses were performed on the inhibitor synthesized by the enzyme from rabbit reticulocytes (28) and from interferon-treated chick embryo cells (24). The results were fully consistent with the assigned structure and, in addition, revealed the presence of the corresponding 5’-diphosphorylated and 5’-monophosphorylated oligomeric series of products. Confirmation of the structure was obtained by comparing enzymatically synthesized inhibitor with material made chemically by the diphenyfphosphochloridate condensation reaction of Michelson (48), and then phosphorylated. The chromatographic properties, proton and phosphorus nu48. Michelson, A . M. (1959). J . Chrm. Soc.. pp. 1371-1394.

292

L. ANDREW BALL

clear magnetic resonance spectra, and the specific inhibitory activities of the two preparations were compared and provided conclusive proof of the chemical structure of the inhibitor (49). A rigorous chemical synthesis has also been performed (50). 2. Anrilyticnl Methods for 2’,5’-Oli~oaden.ylntes Oligomers of 2-5A can be separated from unreacted ATP by salt gradient elution from small columns of DEAE-cellulose ( 1 2 ) ,or by electrophoresis on thin layers of paper or cellulose (13, 15, 49). Separation by thin-layer chromatography, on the other hand, requires the use of specialized solvent systems (41 ). However, removal of the 5’-terminal triphosphate group from both substrate and product (13, 24), or conversion of the unreacted ATP to ADP (381, greatly facilitates their separation by chromatography on thin layers of PEI- or DEAE-cellulose. Chromatography on columns of DEAE-cellulose or DEAE-Sephadex in the presence of 7 M urea is particularly valuable for analyzing both 2-5A oligomers and their corresponding cores (15, 24) since the resolution afforded by this technique depends almost exclusively on net negative charge (51). The most powerful and versatile separation method, however, is highperformance liquid chromatography. Excellent resolution can be achieved using either amine (52) or reverse-phase C-18 columns (53).The latter columns are preferable for routine analyses because the amine columns were found to be unstable in certain solvent systems. 3 . Heterodispersity of 2’,5’-Oligoadenylates in Vivo

An unusual aspect of the 2-5A synthetase reaction was the heterodispersity of the products. Although the trimer was almost always the major species, significant amounts of the dimer and higher oligomers were usually detectable (15). The synthetase purified from Ehrlich ascites tumor cells synthesized a range of chain lengths up to at least 15 residues (38). Moreover, as discussed in Sections III,A,4,b and IV,A, all oligomers except the dimer appeared to be biologically active in most systems. That this heterodispersity of reaction products was not simply an artifact of the isolated enzyme was shown by the detection (by HPLC) of triphosphorylated dimer, trimer, tetramer, and pentamer in acid extracts of interferon49. Martin, E. M . , Birsdall, N. J. M . , Brown, R. E., and Kerr, I. M. (1979). EJB 95, 295-307. 50. Jones, S. S. and Reese, C. B. (1979). JACS 101, 7399-7401. 51. Tener, G. M. (1967). “Methods in Enzymology,” Vol. 12, pp. 398-404. 52. Williams, B. R. G . , Golgher, R. R., Brown, R. E . , Gilbert, C. S., and K e n , I. M. (1979). Notiwe (London) 282, 582-586. 53. Knight, M., Cayley, P. J., Sliilverman, R. H., Wreschner, D. H., Gilbert, C. S., Brown, R. E., and K e n , I. M. (1980). Natrite (London) 288, 189-192.

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293

treated mouse t cells 4 hr after infection with EMC virus (52-54). Some of the corresponding diphosphorylated species were also detected but may have arisen during extraction. Interestingly, substantial amounts of the trimer core (with no 5’-terminal phosphates) were also detected under these conditions (53). As discussed in Section IV,C, this material is inactive as an inhibitor of cell-free protein synthesis, but may have a separate physiological significance. 4. Assay Methods

The assay methods for 2-5A synthetase are all based on detection and quantitation of the enzymic products. They can be divided into three general categories: (a) Those that rely on direct radiochemical detection of the products, (b) those that measure the products by means of their biological activity, and (c) those based on the radiobinding and radioimmune assays. Since the synthetase requires ATP at about lOP3M,the sensitivity of direct radiochemical methods is severely restricted by the specific radioisotopic activities that can be achieved. Routine determination of oligoadenylates at concentrations below about lo-’ M is impractical by these methods. On the other hand, cell-free translation can be inhibited by concentrations of 2’,5’-oligoadenylates in the range from 3 to 6 x 10-’OM [adenylate residue concentrations (55)l. Therefore, assays that detect 2‘ ,5’-oligoadenylates by means of their biological activity are about 100 times more sensitive than direct radiochemical methods. However, not all 2’ ,5’-oligoadenylates have the same biological function, so different assays detect different synthetase products. The characteristics of the major current assays are summarized in Table 11. a. Direct Radiochemical Methods. 2-5A Synthetase, either in solution in the presence of dsRNA or immobilized on poly(1).poly(C)-agarose or poly(1) * poly(C)-paper, reacts with isotopically labeled ATP to form labeled oligoadenylates. The products can be separated from unreacted substrate and quantitated by chromatography on small columns of DEAE-cellulose (56). Alternatively, if [3H]ATPis the substrate, the reaction mixtures can be digested with bacterial alkaline phosphatase and the resulting oligoadenylate cores identified by thin-layer chromatography or 54. Golgher, R . R . , Williams, B. R. G . , Gilbert, C. S., Brown, R . E . , and Kerr, I. M. (1980). A n n . N . Y. Arad. Sci. 350, 448-458. 55. Since most inhibitor preparations are heterodisperse (see Section III,A,2), it is convenient to express inhibitor concentrations in terms of their molar content of adenylate residues. Indeed, it is not yet clear whether the specific inhibitory activities of different oligoadenylates are related to the concentration of oligomer or to the concentration of adenylate residues that they contain. 56. Minks, M . A . , Benvin, S. , Maroney, P. A . , and Baglioni, C. (1979). JEC 254, 50585064.

294

L. ANDREW BALL TABLE I1 CHARACTERISTICS OF 2-5A ASSAYS

Assay method

Sensitivity

2' ,5'-Oligoadenylate species detected

Direct radiochemical assay Protein synthesis inhibition or nuclease activation in lysates of mouse L cells, EAT cells, chick embryo cells, or HeLa cells Protein synthesis inhibition or nuclease activation in rabbit reticulocyte lysates Radiobinding assay

-lO-'M M

All species ( p ) p p A ( ~ Aand ) ~ higher oligomers

M

( p ) p ~ A ( p Aand ) ~ higher oligomers

M

Radioimmune assay

-lo-@ M

(p)ppA(pA)*and higher oligomers; significant reaction with monophosphorylated oligomers ApA and higher nonphosphorylated cores; significant reaction with monophosphorylated oligomers

by their ability to bind to DEAE-cellulose paper disks (34, 39). [3H]Adenosine, the digestion product of unincorporated [3H]ATP, is uncharged and fails to bind to anion exchangers. If [cx-~*P]ATP is the substrate, the reaction mixtures can be incubated with hexokinase and glucose, which facilitates separation by thin-layer chromatography of the oligoadenylate products (which are unaffected by hexokinase) from the residual substrate (which is converted to ADP by reaction with hexokinase) (38). When these radiochemical assays are performed with the enzyme in solution, incubations from which the dsRNA has been omitted should be included as controls. In this way the contributions from other enzyme activities [RNA polymerase, poly(A) polymerase] can be assessed. With the immobilized synthetase, such controls are not possible, but because of the greater purity of the enzyme, they are less necessary. The resistance of 2-5A to digestion with ribonucleases T2 or P1 can be used as a further identifying characteristic of the product. b. Oligoudenylcrte Activity Determination. To determine 2' ,5'oligoadenylate concentrations by measurement of their biological activity, it is necessary to assay a range of dilutions to establish a functional end point, which can then be related to a standard preparation of known concentration. This type of assay is laborious and cumbersome. However,

295

11. 2'5'-OLIGOADENYLATE SYNTHETASE

'Ot

0 4lo-ll

'

' lo-lo

I

I

c L ' 10-9

1p'

AMPResidue Concentration (MI

FIG. 2. Inhibition of cell-free protein synthesis by 2-5A. Protein synthesis was assayed in extracts of mouse L cells programmed with EMC virus RNA, and in extracts of chick embryo cells programmed with VSV messenger RNA. Protein synthesis activity is expressed as a percentage of the messenger RNA-dependent incorporation that occurs in the absence of inhibitor. A,Chick cell-free protein synthesis inhibited by mixed 2-5A oligomers made by 2-5A synthetase from interferon-treated chick cells. 0, Mouse cell-free protein synthesis inhibited by mixed 2-5A oligomers made by 2-SA synthetase from interferontreated chick cells. 0 , Chick cell-free protein synthesis inhibited by mixed 2-SA oligomers made by 2-5A synthetase from non-interferon-treated chick cells. 0 , Chick cell-free protein synthesis inhibited by mixed 2-5A oligomers made by 2-SA synthetase from interferontreated mouse cells. Chick cell-free protein synthesis inhibited by pppAZ'p"A2'p"A. A, Chick cell-free protein synthesis inhibited by pppA2'p5'A2'ps'A2'p5'A.From Ball and White ( 2 4 ) , with permission.

.,

the extreme sensitivity of such assays, and the fact that they measure the presumed physiological function of 2' ,5'-oligoadenylates, partially offset their disadvantages. Two aspects of oligoadenylate function can be monitored: The inhibition of messenger RNA-dependent protein synthesis in an appropriate cell-free system, andor the activation of the latent endoribonuclease that mediates the inhibition by degrading messenger RNA. Cell-free protein synthesis catalyzed by extracts of mouse L cells or primary chick embryo cells in response to exogenous mRNA is 50% inhibited by 2'3'oligoadenylates at concentrations of about 3 x lo-'' M adenylate residue concentration (Fig. 2) (13. 2 4 ) . The translation of viral and cellular exogenous messenger RNAs is equally sensitive to inhibition, but the translation of polyuridylic acid is substantially resistant (24, 57). This is probably 57. Vaquero, C. M., and Clemens, M. J. (1979). EJB 98, 245-252.

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L. ANDREW BALL

because the efficiency of translation of polyuridylic acid is largely independent of chain length. The translation of endogenous messenger RNA is also largely resistant to inhibition, but this is because the bulk of endogenous protein synthesis is usually completed by the time the oligoadenylatemediated inhibition becomes established. Not all cell-free translation systems are equally sensitive to inhibition. Those prepared from cells that lack the 2-SA-dependent nuclease, such as wheat germ, are totally insensitive (24). Systems prepared from some mouse cells lose their sensitivity during the routine preparative steps of preincubation and gel filtration, whereas those made from other cells are fully stable under these conditions (20, 58). Systems prepared from different lines of the same cell type can differ in this regard. The substantial variations that exist in the rate of degradation of the 2' ,5'-oligoadenylates do not appear to be major factors in determining the sensitivities of systems to inhibition (58). A different kind of atypical sensitivity is shown by lysates of rabbit reticulocytes, which are widely used as assay systems for 2',5'oligoadenylates. The translation of exogenous messenger RNA by such lysates is essentially insensitive to inhibition by the trimer, but fully sensitive to the tetramer and higher oligoadenylates (52). In this case, the insensitivity does not appear to be an artifact of the preparative procedures, but an intrinsic property of the 2-SA-dependent nuclease from rabbit reticulocytes, which, although it binds the trimer is not activated by it (52, 53). Whether this is a general property of the rabbit nuclease, or reticulocyte nucleases, or neither, remains to be determined. Alternative assays for the biological activity of 2-SA are based on the direct measurement of the 2-SA-dependent ribonuclease activity. This is a latent endonuclease that is present in both interferon-treated and untreated cells, and which apparently mediates the effects of 2-5A. It can be assayed directly by monitoring the oligoadenylate-stimulatedbreakdown of isotopically labeled RNA added to appropriate cell extracts (52). Although messenger RNA is thought to be the natural substrate for this nuclease, ribosomal RNA can also serve in the assay. (Also see Section IV,B.) The RNA degradation that occurs at several different oligoadenylate concentrations is measured by polyacrylamide gel electrophoresis, and related to the activity of a standard 2-SA preparation. Under optimum conditions, this assay is as sensitive, easier, and less prone to artifacts than measurement of the inhibition of cell-free protein synthesis. 58. Williams, B. R . G ., Kerr, I. M . , Gilbert, C. S . , White, C. N., and Ball, L. A. (1978). EJB 92, 455-462.

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c. Radiobinding rind Rndioinimirne Assciys. Probably the most versatile assays for 2’ 3’-oligoadenylates and their corresponding dephosphorylated cores are the radiobinding and radioimmune assays that have been developed (53). These assays are based on the high affinities of 2’,5’oligoadenylates and dephosphorylated cores for binding to the 2SA-dependent nuclease and to antibody against cores, respectively. Crude cell extracts were used as sources of the nuclease in the radiobinding assay; rabbit antisera raised against (AZ’p),Aconjugated to bovine serum albumin were used in the radioimmune assay. The assays measure binding competition between oligoadenylate (or core) samples of unknown concentration and “probe” material labeled to a very high specific radioactivity. Binding is determined by retention of the probe on nitrocellulose filters, and samples of known concentration are used to calibrate the assays. The sensitivity of such assays depends on the specific isotopic activity of the probe and, as previously mentioned, the properties of the synthetase limit what can be achieved by biosynthesis. Accordingly, T4 RNA ligase was used to add [5’-32P]pCpto the 3‘ end of preformed oligoadenylates and cores, achieving specific activities of 2-3 x lo6 Ci/mol of oligonucleotide. Probes of this sort enable the radiobinding and radioimmune assays, respectively, to detect oligoadenylates and cores at concentrations below M. The radioimmune assay is unique in its ability to detect cores in this concentration range. The specificities of the radiobinding and radioimmune assays are shown in Table 111. 5. Requirements of the Enzymatic Reaction 2-5A Synthetase has three requirements: dsRNA, ATP, and magnesium ions. It is clear from studies of both crude and purified enzyme preparations that complex quantitative interrelationships exist between these requirements, so a complete understanding of the situation must await more thorough studies of the pure enzyme. However, the present knowledge of the requirements is summarized below. a. dsRNA. No quantitative conclusions concerning the dsRNA requirement can be drawn from experiments in which the enzyme was studied in its immobilized state, bound to poly(1)-poly(C)-agarose or -paper. ConverseIy, studies of the properties of the enzyme in crude extracts may be subject to interference from contaminating activities that could compete for or degrade the dsRNA, or degrade the oligoadenylate products. Nevertheless, with these uncertainties in mind, the following properties of the crude or partially purified enzyme can be noted. The requirement for activation by dsRNA cannot be satisfied by dsDNA or by RNA-DNA hybrids (24). Some RNA preparations that are predominantly single stranded can activate the enzyme with low efficiency, presumably by

298

L. ANDREW BALL TABLE I11 SPECIFICITY O F THE

Compound

R A D I O B I N D IANNG D

RADIOIMMUNE

ASSAYS"

Rndiobinding nssay Concentration for 50% displacement of PPP(A"P)SA-[~*PIPCP

I x 10-5~ I x 10-9~ 1 x 10-9~ I x 10-9~ 3 x 10-7~ >3 x 10-"M > I x 10-3 M >i x 1 0 - 3 ~

pppA2'p5'A ppp(A2'p),A, ( n = 2 to 4) pp(A"p)A p(AZ'p)A (A"p)A (A3'p),A and (A3'p)3A 2'-, 3'-, and 5'-AMP, ADP, ATP 2'-, 3'-, and 5'-CMP, pCp Radioimmune cissny

Concentration for 50% displacement of (A"P)~A-[~PIPCP A "P" A (A2'p)A and (A"p)y4 P(A"P),A PP(A"P)zA PPP(A"P)ZA 2'- and 3'-AMP 5'-AMP (A3'p),A and (A3'p)y4 A"P"A Adenosine ATP, CTP, pCp, 2'-, 3'-, and 5'-CMP

1 1 5 I 5 5 1 I 5 5 >I

x 10-8M x 10-8M x 10-9~ x 10-7~ x 10-7 M x 10-6M x 10-jM x 10-5 M

x 10-5M x 10-'M x 1 0 - 3 ~

Reproduced from Knight rt NI. (53),with permission.

virtue of intramolecular regions of base-paired secondary structure (59). On the other hand, RNA molecules that contain both single-stranded and interchain double-stranded regions, such as the replicative intermediates of picomavirus replication, activate the enzyme efficiently (60, 6 1 ) . In rabbit reticulocyte lysates, the synthetase requires poly(1).poly(C) at lo-' g/ml for full activation. Concentrations that are tenfold lower and, 59. Revel, M., Kimchi, A., Schmidt, A., Shulman, L., and Chernajovsky, Y. (1979). I n "Regulation of Macromolecular Synthesis by Low Molecular Weight Mediators" ( G . Koch and D. Richter, eds.), pp. 341-359. Academic Press, New York. 60. Nilsen, T. W.,and Baglioni, C. (1979). PNAS 76, 2600-2604. 61. Nilsen, T. W., Maroney, P. A., and Baglioni, C. (1979). In "Regulation of Macromolecular Synthesis by Low Molecular Weight Mediators" ( G . Koch and D. Richter, eds.), pp. 329-339. Academic Press, New York.

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299

surprisingly, tenfold higher than this are less effective (62). The enzyme from interferon-treated HeLa cells is also saturated with poly(1).poly(C) at about g/ml but in this case there is no suggestion of decreased activation at supersaturating concentrations (56). The dsRNA requirement of the synthetase is thus distinctly different from that of the dsRNA-activated kinases from either rabbit reticulocytes or interferontreated cells, which require dsRNA at only lo-’ g/ml or less for full activation. For the enzyme purified to homogeneity from Ehrlich ascites tumor cells, the optimal concentration of dsRNA increases with the enzyme concentration (47). Optimal activity is achieved when the concentration of dsRNA is about half that of the enzyme (by weight), which corresponds to about 80 base-pairs per enzyme molecule ( 4 7 ) . At higher dsRNA concentrations, the enzyme activity decreases, but it is not known if this reflects the need for cooperative binding of enzyme molecules to the activating dsRNA. However, cooperative behavior is also suggested by the sigmoidal shape of the curve relating reaction rate to enzyme concentration (47). A spherical protein of the apparent molecular weight of the mouse synthetase would have a diameter that could cover only about 20 base-pairs. However, interstrand mismatches in poly(1) * poly(C) that are more frequent, on average, than one every 35 I-C base-pairs, or strand discontinuities that are more frequent than one every 65 bp, decrease the efficiency of activation (62). Poly(I).poly(C) in which one strand is fully 2‘-0-methylated is inactive in synthetase activation. However, 2’-0methylation of up to 40% of the bases in either strand does not significantly impair the activity. Minkser id. have suggested that these results can be reconciled if the separate processes of binding and activation have different structural requirements (63). Not all dsRNAs are equally effective in activating 2-5A synthetase. At all concentrations tested, poly(riboadeny1ic).poly(ribouridy1ic acid) and natural dsRNAs from Penicillium chrysogetzurn and reovirus are somewhat less effective than poly(I)*poly(C)in activating the enzyme in rabbit reticulocyte lysates (64). However, studies of the homogeneous synthetase from Ehrlich ascites tumor cells show that the relationships are complex, and depend on the enzyme concentration at which the assays are performed (47). 62. Minks, M. A., West, D. K., Benvin, S. , and Baglioni, C . (1979). JBC 254, 1018010183. 63. Minks, M . A . , West, D. K., Benvin, S . , Greene, J. J . , Ts’o, P. 0. P., and Baglioni, C. (1980). JEC 255, 6403-6407. 64. Williams, B. R. G . , Gilbert, C. S . , and Kerr, I . M. (1979). Ntrcleic Acids Res. 6, 1335-1350.

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L. ANDREW BALL

h. ATP. The K mfor ATP has not been determined. Most workers assay the synthetase, either in solution or in the immobilized state, at ATP concentrations of 1-5 mM. The enzymes from mouse L cells and rabbit reticulocytes show very low activity at ATP concentrations less than 0.10.2 mM (28,37). Minks et ul. (56)found that the activity of the HeLa cell enzyme was directly proportional to ATP-Mg2+concentration in the range 0.5-5.0 mM, but at a constant magnesium ion concentration, the ATP dependence was more complex. These data suggest that the Km for ATP-Mg2+is probably greater than 2 mM. Similarly, the ATP-dependence of the pure enzyme from Ehrlich ascites tumor cells suggests a Km for ATP in the region of 2 mM (47). Whether dimer synthesis, which is essentially a chain initiation reaction, has different characteristics in this regard from the subsequent steps that constitute chain elongation remains to be determined. However, the efficient use of low concentrations of nucleotidyl acceptors other than ATP (see Section II1,C) suggest that the enzyme may have distinct Km's for nucleotidyl donor and acceptor. A remarkable feature of the 2-5A synthetase reaction is that under optimal conditions essentially 100% of the ATP substrate can be converted to oligonucleotide product. This has been observed that the crude chick enzyme (L. A. Ball, unpublished results) and, more significantly, with the pure mouse enzyme (38, 47). This is remarkable because phosphodiester bond formation in polynucleotides usually occurs with an insignificant free energy change. This means that such reactions have an approximately central equilibrium position unless, for example, the presence of an inorganic pyrophosphatase displaces the equilibrium in favor of product formation. Samanta et ul. (47) have established that the equilibrium position of the reaction catalyzed by the pure mouse synthetase lies 96-98% toward 2-5A synthesis, despite the lack of hydrolysis of the inorganic pyrophosphate. Furthermore, the reverse reaction, i.e., the pyrophosphorolysis of 2-5A, and the exchange of [32P]inorganic pyrophosphate into ATP were undetectable under their conditions of assay. These observations indicate that the thermodynamic properties of 2'-5' bonds differ substantially from those of the corresponding 3'-5'linkages. It remains to be determined if this is a consequence of features of the three-dimensional structure of 2-5A that are relevant to its mechanism of action (see Section IV,A). c. Magnesium Ions. The synthetase is dependent for activity on the presence of magnesium ions. With the pure enzyme, the optimum magnesium ion concentration lies in the range from 8- 16 mM, the exact value being dependent on the ATP concentration (47). With the crude enzyme from HeLa cells (56) or chick embryo cells (39), on the other hand, the activity increases as a function of magnesium ion concentration up to at

11.

2',5'-OLIGOADENYLATE SYNTHETASE

30 1

least 30 mM or 50 mM, respectively. However, in both cases the bulk of the effect occurs below 20 mM. The possibility that other divalent cations-manganous or calcium ions, for example-could substitute for magnesium ions has not been explored. d . Other Fcictors. The distribution of product sizes is a function of the enzyme activity. Thus, highly active or concentrated enzyme preparations synthesize products that contain a greater proportion of higher oligomers. This is probably a consequence of the nonprocessive mechanism of chain elongation (see Section III,A,6). The pH optimum for the pure mouse enzyme is pH 7.8 (47), and most assays in other systems have been performed between pH 7.0 and 8.0. The enzyme has no apparent requirement for monovalent cations and, indeed, KCl concentrations greater than about 0.1 M severely inhibit its activity (64). Since the enzyme can be eluted from poly(I).poly(C)-agarose affinity columns by 0.2 M salt, it is likely that the inhibitory effect of KC1 is due to a weakening of the interactions between the enzyme and its dsRNA activator. Glycerol (10-20%), bovine serum albumin (0.1-1.0 mg/ml), and sulfhydryl reagents are often added to stabilize the partially purified enzyme. 6. Kinetics and Mechanism of Reaction The specific activity of the pure enzyme from Ehrlich ascites tumor cells is 2.4 pmol AMP polymerized/mg protein/hr when the enzyme is assayed at 30" in the immobilized state, and about 70 pmol AMP polymerized/mg protein/hr when it is assayed in solution (38, 47). This difference in the rates of reaction of the soluble and immobilized enzyme has not been observed by other workers (37). The immobilized enzyme is extremely stable at 30", and the reaction can continue for several hours until the substrate is exhausted. If the enzyme is regularly replenished with ATP, the reaction can continue essentially unabated for several days (49).Since the reaction products constitute an oligomeric series, the kinetics of the accumulation of any particular oligomer are not necessarily the same as the kinetics of substrate utilization (41, 46, 65). The effect of enzyme concentration on the distribution of product sizes (see Section III,A,S,d) first suggested a nonprocessive mechanism of elongation, and this was confirmed directly by the demonstration that purified, preformed 2-5A trimer could be further elongated by the activated chick enzyme (39). The nonprocessive nature of the reaction mechanism has been confirmed for HeLa cell synthetase (45). Analysis of the distribution of isotopic label in the product of a two-step synthesis indi65. Schmidt, A . , Zilberstein, A., Shulman, L., Federman, P., Berissi, H.. and Revel, M . (1978). FEBS Lett. 95, 257-264.

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cated that the direction of oligoadenylate chain growth was 5' --* 2' (40). No evidence for ligase activity (the joining of oligoadenylate molecules) has been found. 7 . Stability of the Reaction Products In most crude cell extracts, low concentrations of 2-5A lose biological activity with a half-life of a few minutes (58, 6 6 ) .A priori, this could be due to loss of the j3-and y-phosphate groups, which are necessary for nuclease activation, from the 5' terminus, or to cleavage of the 2'-5' phosphodiester bonds. However, in all cases where 2-5A degradation has been investigated, the latter reaction appears to be responsible for the loss of biological activity in cell extracts ( 2 3 , 5 8 , 6 6 ) .Breakdown occurs by progressive removal of 5'-AMP residues from the 2' terminus of 2-5A, and can be inhibited by 5'-AMP (39). Schmidt et at. (67) have purified, from mouse L cells, a phosphodiesterase that catalyzes 2-5A degradation. Although they find that this enzyme is induced by interferon, untreated cells nevertheless contain high basal levels. The action of this enzyme on 2-5A resembles that of snake venom phosphodiesterase, except that the former yields 5'-ATP, rather than 5'-AMP + inorganic pyrophosphate, from the 5'-terminus of 2-5A. Preparations of the synthetase can be freed of the phosphodiesterase activity by binding to immobilized dsRNA, or by passage through DEAE-cellulose (65). 8. SynthetLise Inhibitors 2-5A Synthetase is inhibited by removal of the activating dsRNA, by digesting it with ribonuclease 111, for example ( 4 6 ) . Some nucleoside triphosphates have been reported to inhibit ATP incorporation by the HeLa cell enzyme: 2'-dATP, 3'-dATP, GTP and P,y-methylene-ATP ( 4 6 ) . On the other hand, it has been reported that the rabbit reticulocyte synthetase can use all eight common ribo- and deoxyribonucleoside-5'triphosphates as nucleotidyl donors (41, 68) (see Section III,B), and 3'-dATP as both a nucleotidyl donor and acceptor (69). Whether these results reflect properties that are unique to the rabbit reticulocyte synthetase remains to be determined. 66. Minks, M . A . , Benvin, S . , Maroney, P. A . , and Baglioni, C. (1979). Nrrcleic Acids ReS. 6, 767-780. 67. Schmidt, A . , Chernajovsky, Y., Shulman, L . , Federman, P., Berissi, H . , and Revel, M . (1979). P N A S 16, 4788-4792. 68. Justesen, J . , Ferbus, D . , and Thang, M. N . (1980). P N A S 77, 4618-4622. 69. Doetsch, P., Wu, J. M., Sawada, Y . , and Suhadolnik, R. J. (1981). Nntrrre (London) 291, 355-358.

11.

2‘,5’-OLIGOADENYLATE SYNTHETASE

303

B. NUCLEOTIDYL DONATION B Y NUCLEOSIDE 5’-TRIPHOSPHATESOTHER T H A NATP Incubation of a partially purified synthetase preparation from rabbit reticulocytes with mixtures of ATP and any of the other seven common ribo- or deoxyribonucleoside 5‘-triphosphates results in the formation of co-oligomers with the general structure pppA2(p5’A),pN ( 4 / , 68). All the internucleotide linkages in these co-oligomers are presumed to be 2‘-5’ since they resist digestion with nuclease P1, but the nature of the 3’ proximal linkage has not been demonstrated directly. The heterologous nucleotides are apparently incorporated only at the 2‘ terminus of the products and hence function as chain terminators (68). Accordingly, similar co-oligonucleotides can be formed by heterologous nucleotidyl transfer to preformed 2-5A oligomers. The biological activities and physiological significance of these co-oligomers remain to be evaluated. The fluorescent derivative of ATP, 1,N-6-etheno-ATP, is a substrate for the HeLa cell synthetase, and the resulting 1,N-detheno-ZSA retains some biological activity in the nuclease activation assay (46). Similarly, it has been reported that 3’-dATP (cordycepin 5’-triphosphate) is a substrate for the synthetase from rabbit reticulocytes (69) and Ehrlich ascites tumor cells (47), although apparently not for that from HeLa cells (46). Interestingly, 3I-deoxy-2-5A has been reported to inhibit protein synthesis in rabbit reticulocyte lysates and to resist hydrolysis (69).

c.

2’-ADENYLYLATION OF ACCEPTORS OTHER THANATP 2-5A

AND

The observation that oligoadenylate synthesis occurred by a nonprocessive mechanism (40) (see Section 111,A,6) prompted an investigation of the structural requirements for nucleotidyl acceptor function. The sixteen 3’,5’-diribonucleoside monophosphates (X3’p5‘Y,where X and Y are any of the four naturally occurring ribonucleosides) were tested for their ability to serve as acceptors in adenylyl transfer reactions catalyzed by the chick embryo synthetase (40). Those that contained 3‘-adenylate residues (A3’p5’A,C3’p5’A,G3‘p5’A,and U3’p5’A)were found able to accept one or more further adenylate residues in 2’-5’ linkage, to form molecules of the general structure N3‘p5‘Az‘(p3’A)n. Those with other 3’-terminal residues were inactive. The structural requirement for 2’,5’-linked diribonucleoside monophosphates was the same, namely the presence of a 3‘-terminal adenylate residue. ADP-ribose, NAD’, NADH, and 5’3”diadenosine tetra- and pentaphosphate were also efficient adenylate accep-

304

L. ANDREW BALL

tors (38, 40, 4 6 ) . Conversely, none of the following molecules could function as adenylate acceptors under standard reaction conditions: poly(A), poly(A)-containing cellular messenger RNA, S -adenosylmethionine, coenzyme A, NADP’or adenosine (40).It is unclear which, if any, of the 2’-adenylylated acceptors can function as activators of the 2-SA-dependent nuclease. However, 2‘-adenylylated NAD’ is significantly less active as an electron acceptor than the nonadenylylated coenzyme (40). There is no evidence that adenylylation of these acceptors occurs in whole cells, or that these reactions have any physiological significance. a ,P- and @,y-Methylene-5‘-ATP can act as adenylate acceptors, but not as donors for the chick embryo cell enzyme. The products are oligomeric series of 2-5A molecules with methylene bridges between the a and p or P and y phosphates of the 5’-terminal triphosphate group. Despite the fact that the methylene bridges cannot be hydrolyzed, these analogs of 2-5A appear to retain biological activity, at least when assayed in a chick embryo cell-free system (39).These results indicate that nuclease activation by 2-5A does not involve cleavage between either the a- and @-phosphates, or the @- and y-phosphates at the 5’ end of 2-5A. IV.

Biological Role

It is clear from studies of cell extracts that the trimer and higher oligomers of 2-5A are potent inhibitors of protein synthesis, and that this inhibition is mediated by the activation of a latent endoribonuclease (14, 18,23, 57, 59, 65, 66, 70-77). The artificial introduction of 2-5A into intact cells causes similar effects and can partially inhibit virus replication (78-82). 70. Eppstein, D. A., and Samuel, C. E. (1978). Virology 89, 240-251. 71. Shaila, S . , Lebleu, B., Brown, G. E., Sen, G . C., and Lengyel, P. (1977). J . Gen. Virol. 37, 535-546. 72. Lewis, J. A., Falcoff, E., and Falcoff, R . (1978). EJB 86, 497-509. 73. Ball, L. A., and White, C. N. (1979). Virology 93, 348-356. 74. Farrell, P. J., Sen, G. C., Dubois, M. F., Ratner, L . , Slattery, E., and Lengyel, P. (1978). P N A S 75, 5893-5897. 75. Chernajovsky, Y., Kimchi, A . , Schmidt, A., Zilberstein, A., and Revel, M. (1979). EJB 96, 35-41. 76. Slattery, E., Ghosh, N . , Samanta, H . , and Lengyel, P.(1979). PNAS 76,4778-4782. 77. Baglioni, C., Minks, M. A., and Maroney, P. A. (1978). Ncrtrtre (London) 273,684687. 78. Hovanessian, A. G., Wood, J. N . , Meurs, E., and Montagnier, L. (1979). I n “Regulation of Macromolecular Synthesis by Low Molecular Weight Mediators” (G. Koch and D. Richter, eds.), pp. 319-327. Academic Press, New York. 79. Williams, B . R. G., Golgher, R. R., and Kerr, I. M. (1979). FEBS Left. 105, 47-52.

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Moreover, 2-5A oligomers have been detected in interferon-treated, virusinfected cells at concentrations that seem to be sufficient to activate the 2-5A-dependent endonuclease (52-54). However, there has been no clear demonstration in whole cells of enhanced RNA turnover that could be attributed to the combined action of the components of the 2-5A system. The presence of the 5‘-terminal triphosphate group on 2-5A oligorners renders them highly polar molecules, and presumably mitigates against their uptake by whole cells. Certainly, no effects of triphosphorylated 2-5A on whole cells have been observed without the use of artificial means to promote uptake (52). However, nonphosphorylated 2-5A cores can apparently inhibit cell growth and DNA synthesis in certain cells (83, 84). It is not clear if these effects are due to a distinct biological activity of the cores themselves, or to their uptake, intracellular phosphorylation, and consequent action as nuclease activators. These questions, and the role of the 2-5A system in interferon action and normal cellular metabolism, are discussed in the following sections. ACTIVATION B Y 2-5A A. NUCLEASE

IN

CELLEXTRACTS

The 2-5A-dependent endonuclease has two competing designations, RNase F (65) and RNase L (38) (for “latent”). To judge from the concentrations of 2-5A that are sufficient to fully activate the enzyme in crude cell extracts (and assuming a stoichiometric relationship), the nuclease may be a very minor cellular component. It has not been purified to homogeneity; the most extensive purification reported was from mouse L cells and resulted in a 1000-fold increase in specific activity (65, 75). The partially purified enzyme retained its dependence on 2-5A, and had detectable 2-5A-binding activity (75). (Nuclease binding of 2-5A is thought to be the basis of the radiobinding assay described in Section 111,A,4,c.) Upon gel filtration, the enzyme behaved as a protein of 60,000-80,000 molecular weight, but there were several polypeptides of this size in the partially purified preparation (59). On the other hand, a 100-fold purified enzyme preparation from Ehrlich ascites tumor cells had an apparent molecular weight of 185,000 on gel filtration (76). No gross changes were detected in the size or shape of the enzyme in response to activation. 80. Hovanessian, A. G.,Wood, J . , Mews, E . , and Montagnier, L. (1979). P N A S 76, 3261- 3265. 81. Hovanessian, A. G.,and Wood, J . N. (1980). Virology 101, 81-90. 82. Williams, B . R. G . , and Kerr, I. M. (1978). Nature (London) 276, 88-90. 83. Kimchi, A., Shure, H., and Revel, M. (1979). Nature (London) 282, 849-851. 84. Reisinger, D. M. and Martin, E. M. (1980). Proc. Intern. Workshop Interferons, held at the Memorial Sloan-Kettering Center, New York, April 22-24, 1979.

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In order to function as a nuclease activator, a 2-5A oligomer must contain at least three adenylate residues and at least two 5'-terminal phosphate groups. Hence, 2-5A dimers and monophosphorylated higher oligomers are essentially inactive (49). Any residual activity can probably be ascribed to contamination with other species. As previously discussed (Section III,A,4,b), the nuclease from rabbit reticulocytes fails to respond to activation by 2-5A trimer, but is fully responsive to higher oligomers (52). Other examples of different structural requirements may be discovered as more sources of latent nuclease are investigated. The latent nuclease depends on the continued presence of 2-5A for its activity; if the 2-5A is removed or degraded, the enzyme loses its catalytic activity, but it can be reactivated by addition of more 2-5A (58, 66, 76). These properties, and the 2-5A-binding activity of the partially purified enzyme, suggest that 2-5A exerts its effects by direct interaction with the nuclease. A consequence of this is that nuclease activation and protein synthesis inhibition in crude cell extracts is transient (58). The phosphodiesterase that is responsible for 2-5A hydrolysis (see Section III,A,7) copurifies with the latent nuclease through several (but not all) of the purification steps (65). This observation has prompted the suggestion that the two enzymes may be intimately associated with each other in the cell, and that the phosphodiesterase may modulate nuclease activity by controlling the levels of 2-5A. Various lines of evidence suggest that 2-5A does not act as a phosphate donor in its activation of the nuclease; for example, the comparable activities of di- and triphosphorylated oligomers as inhibitors of protein synthesis, as nuclease activators, and in the radiobinding assay (49, 53); and the activity shown by a,P- and P,y-methylene-bridged analogs of 2-5A (39). However, there has been one report of enhanced phosphorylation of a 110,000 dalton polypeptide in the presence of [y3'P]2-5A (23). The functional significance of this phosphorylation is unclear. The substrate specificity of the latent endoribonuclease has not been fully determined. It shows no activity against single- or double-stranded DNA, or against double-stranded RNA (85-87). Despite a journalistic report to the contrary (88),there is no evidence that the nuclease can degrade 2-5A. Like any endonuclease, it is more active on larger substrate 85. Brown, G. E . , Lebleu, B . , Kawakita, M . , Shaila, S., Sen, G . C., and Lengyel, P. (1976). BBRC 69, 114-121. 86. Ratner, L . , Sen, G . C . , Brown, G. E . , Lebleu, B . , Kawakita, M., Cabrer, B., Slattery, E. and Lengyel, P. (1977). EJB 79, 565-577. 87. Sen, G . C . , Lebleu, B., Brown, G . E., Kawakita, M . , Slattery, E . , and Lengyel, P. (1976). Natwc, (Lorzdtm) 264, 370-373. 88. Hunt, T. (1978). Nature (London),273, 97-98.

11.

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molecules than on smaller ones (86). It is not clear if this effect alone is responsible for the more rapid degradation of Mengovirus RNA than globin mRNA, for example, or whether some additional specificity for viral RNAs is involved. The action of the purified nuclease on both these RNAs is very limited and yields a continuum of RNA fragments of 50,000500,000 molecular weight (59, 65). This limited, semi-random cleavage suggests that the nuclease recognizes RNA features other than the primary nucleotide sequence, perhaps regions containing little secondary structure. However, analysis of the products of terminally labeled RNAs of known sequence indicates that the enzyme has some sequencespecificity, and cleaves with particular enthusiasm to the 3‘ side of pUpNp sequences, with a strong preference for pUpAp and pUpUp sequences. The products of cleavage carry 3’-terminal phosphate groups and 5‘terminal -OH groups (89). Interestingly, activation of the endonuclease in crude cell extracts leads to the enhanced degradation of messenger RNA to acid-soluble mono- and oligonucleotides (58, 73). This effect is presumably due to the combined actions of the 2-5A-dependent nuclease and other, nonlatent, endo-, and exonucleases.

B.

OCCURRENCE A N D ACTION OF 2-5A

IN

WHOLECELLS

Interferon-treated, EMC virus-infected mouse L cells contain levels of 2-5A oligomers that are detectable either by biological assays, or by the radiobinding and radioimmune assays (52, 5.3). The amounts recovered correspond to intracellular concentrations of 20-200 x lo-’ M (trimer equivalents), and HPLC analyses of this material showed that it contains di- and/or triphosphorylated dimer, trimer, tetrarner, and pentamer. Higher oligomers may be present at concentrations below the detection limits of the assays (about lo-’ M ) . Interestingly, the radioimmune assay also indicates the presence of trimer core at concentrations of 5-50 x lo-’ M under these conditions (53). Cells that have been either treated with interferon alone, or infected with EMC virus alone (rather than receiving both treatments) contain levels of oligoadenylates that are about one-tenth of the levels in interferon-treated, infected cells. These 2-5A concentrations, although at the limits of detection of the assays used, would appear to be in the range required to activate the 2-SA-dependent nuclease. The presence of 2-5A in interferon-treated, uninfected cells is particularly significant since it suggests that some component present in uninfected cells 89. Wreschner, D. H . , McCauley, J. W., Skehel, J . J . , and Kerr, I . M . (1981). Nafrrre 289, 414-4 17.

( Lo/7c/on)

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can satisfy the dsRNA requirement for synthetase activation. Moreover, the lower concentration of 2-5A under these conditions suggests that synthetase activity is normally limited by the supply of activator. The addition of triphosphorylated oligomers of 2-5A to intact cells has no detectable effects on protein synthesis. This is not surprising in view of the highly polar nature of the molecules, which presumably prevents them crossing the cell membrane. However, addition of the oligomers to cells using procedures that are designed to promote their uptake, such as hypertonic shock (79, 82) or calcium phosphate coprecipitation (78, 80, 81), results in the enhancement of a nuclease activity, the increased breakdown of messenger RNA, and a clear inhibition of protein synthesis. Active 2-5A can be recovered from the treated cells (79), so these effects appear to be mediated by 2-5A taken up from the medium. Viral RNA synthesis and replication can be inhibited in infected cells treated with 2-5A by these methods. However, there is no evidence that infected cells differ from uninfected cells in their sensitivity to 2-5A, either directly, or because of an infection-mediated change in membrane permeability. The magnitude of the effects of 2-5A depends on the concentration with which the cells were treated, in the range lo-'' to loe7M. At low concentrations, the inhibition of protein synthesis is transient, but at concentrations greater than about 2 x lo-' M, the effect persists, and secondary inhibitions of RNA and DNA synthesis and of cell multiplication are evident. Messenger RNAs are not the only RNA species to be affected by the introduction of 2-5A into cells. Surprisingly, 28 S ribosomal RNA is also partially degraded under these conditions, yielding two or more large fragments (78, 80). Similar cleavage of 28 S ribosomal RNA has been observed in cells infected with SV40 and subsequently treated with interferon (59, 90). It is particularly significant that in this case no extensive degradation of SV40 messenger RNA was detected. Although it is not certain that these effects on ribosomal RNA were mediated by the 2-5A-dependent nuclease, the results nevertheless raise the intriguing possibility that the natural substrate for the enzyme in intact cells is not (only) messenger RNA, but some other component of the protein synthesizing machinery. An effect of the 2-5A-dependent nuclease on ribosomal RNA is reminiscent of the mechanism of action of colicin E3, which cleaves the 16 S RNA in the small subunit of E. coli ribosomes (91). 90. Revel, M., Kimchi, A., Shulman, L . , Fradin, A . , Shuster, R., Yakobson, E . , Chernajovsky, Y., Schmidt, A . , Shure, H . , and Bendori, R . (1980). Ann. N . Y. Acad. Sci. 350, 459-472. 91. Bowman, C. M . , Sidikaro, J . , and Nomura, M. (1971). Nutitre New B i d . 234, 133137.

I I. 2',5'-OLIGOADENYLATESYNTHETASE

c.

EFFECTSOF 2-SA CORES

ON

309

WHOLE CELLS

Nonphosphorylated 2-5A cores are inactive as inhibitors of protein synthesis in cell-free extracts, and neither activate nor bind to the 2-5A dependent nuclease. However, chemically synthesized cores show significant activity when assayed as inhibitors of protein synthesis in permeabilized cells (82). It is assumed that this reflects the ability of whole cells to add a 5'-terminal di- or triphosphate group to the cores, an ability that appears to be lacking from cell-free extracts. Cores are much less polar molecules than the corresponding triphosphorylated species, so the possibility that they affect unpermeabilized cells was investigated. Surprisingly, the effects that have been reported are inhibitions of cellular DNA synthesis, both in lymphoblastoid (Daudi) cells (84), and in mitogen-stimulated mouse spleen lymphocytes (83). In both cases, chernically synthesized 2-5A cores were used, and the corresponding 3 ' 3 ' linked oligomers were inactive. The effects were maximal 24-48 hours after treatment of the cells, and occurred in the absence of a general inhibition of protein synthesis. It is difficult to explain these effects in terms of the known mechanism of the action of 2-5A, although it is conceivable that the RNA primers on which DNA synthesis occurs are substrates for the 2-5A-dependent nuclease. D. ROLEO F

THE

2-5A SYSTEM

IN

INTERFERON ACTION

As described in Section I, the 2-5A system was discovered as the result of experiments that were designed to elucidate the mechanism by which dsRNA inhibited protein synthesis in extracts of interferon-treated cells. However, it is not yet clear what role the 2-5A system plays in the effects that interferon has on the biology of whole cells, and some major questions remain unresolved. For example, there is little evidence that the breakdown of viral RNA occurs at an enhanced rate in most interferontreated infected cells, as would be predicted from the studies of cell extracts. The measurement of RNA breakdown is often technically difficult and the lack of data on this point may be largely due to the right experiments not having been done. However, there are some clear situations in which one would expect the 2-5A system to be operative, but where intact viral messenger RNA accumulates to levels that even exceed those in noninterferon-treated cells. One such example is that of interferon-treated cells infected with vaccinia virus (92). In these situations it seems unlikely 92. Metz, D. H.,and Esteban, M. (1972).Notiire (London) 238, 385-388.

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that the inhibition of viral replication can be attributed to enhanced breakdown of messenger RNA. Another concern is the question of whether the 2-5A system is specifically antiviral in its action. As discussed in Section IV,A, the 2-5A-dependent nuclease degrades both host and viral messenger RNAs, displaying little clear preference. Its activation might therefore be expected to result in a breakdown of both viral and cellular messenger RNAs. However, it is clear that in many situations interferon-treated cells can survive virus infection. Two suggestions have been made to resolve this paradox: First, that nuclease activation is transient and the cell (but not the virus) can therefore recover (58). The transient nature of the effects of 2-5A introduced into permeabilized cells (Section IV,B) supports this possibility. Second is that nuclease activation affects only limited regions of the cytoplasm, which constitute microenvironments surrounding each activating dsRNA molecule (60, 73). Since the dsRNA is thought to be provided by virus replication, these microenvironments might correspond to those regions of the cytoplasm where viral syntheses are taking place. Indeed, under certain conditions, it is possible to demonstrate that single-stranded RNAs that are covalently joined to doublestranded regions are preferentially degraded by activating the 2-5A system in cell extracts (60, 6 1 ) . Furthermore, 2-5A synthetase and the 2-SA-dependentnuclease copurify to some extent, suggesting that they may be functionally associated in the intact cell (45). This, together with the proximity of the 2’-phosphodiesterase (see Section IV, A) could serve to restrict the sphere of influence of the activated 2-5A system. Unfortunately, such ideas are very difficult to test experimentally. As noted in Section II,B, the basal levels of 2-5A synthetase vary widely in different cells, and bear no apparent relation to the ability of the cells to support virus replication. For example, HeLa cells, which are often used as laboratory host cells for a wide range of viruses, have a basal level of synthetase that is higher than the level in interferon-treated L cells, which are essentially nonpermissive for virus growth. Of course, the synthetase is only one component of the 2-5A system; differences in compartmentation or in the levels of dsRNA activator, 2-5A-dependent nuclease, or 2’-phosphodiesterase may be more important in determining the overall level of control that the system imposes. Some attempts have been made to assess the relative contributions of the interferon-induced kinase and synthetase to the virus-resistant state. For example, the induction of the kinase in HeLa cells can be prevented by actinomycin D under conditions where there is little inhibition of synthetase induction ( 2 2 ) . EMC viral RNA synthesis is inhibited under these

11. 2',5'-OLIGOADENYLATE SYNTHETASE

311

conditions, a result that is consistent with a role for the synthetase in the antiviral state. However, definitive indications of the role of the 2-5A system in interferon action must await the identification and characterization of variant or mutant cell lines that lack 2-5A synthetase. At present, no such cells are available, but some cell lines have been described that show basal synthetase levels that fail to respond to interferon treatment: For example, human endometrial cancer (HEC-1) cells (29) and WBalb mouse fibroblasts (32). In the former cells, the levels of dsRNAdependent phosphokinase also fail to respond to interferon treatment, and no resistance to infection by VSV or Sindbis virus develops. Like the interferon-resistant L1210R cells (93), HEC- 1 cells may lack interferon receptors. More revealing is the case of WBalb mouse fibroblasts, which respond to interferon by induction of the phosphokinase and the development of resistance to VSV infection, but show no synthetase induction (32). These results suggest that enhanced levels of 2-5A synthetase are not necessary for resistance to VSV. The properties of another cell type, mouse embryonal carcinoma cells, indicate further that 2-5A synthetase induction alone is insufficient to confer resistance to VSV infection (19). These latter cells show normal induction of the synthetase in response to interferon treatment, but no induction of the kinase or development of resistance to infection by VSV, Sindbis, or influenza virus. However, they do develop resistance to EMC virus and Mengovirus (94). While it is not clear that this resistance is mediated by the 2-5A system, no rival mechanisms are known with which to explain the results. It has been suggested that the interferon sensitivity of EMC virus and Mengovirus in these cells is due to the greater accumulation, during picornavirus infection, of viral dsRNA, which is required to activate the 2-5A synthetase (94, 95). The implication is that the resistance to VSV shown by other cells is mediated by mechanisms other than the 2-5A system. Indeed, in human MRC5 cells, interferon induces resistance to infection by VSV without an effect on the basal levels of either 2-5A synthetase or the protein kinase (33). Presumably some other mechanism, such as interference with virion maturation (%, 97), is responsible for viral resistance in this case. Thus the picture emerges of the 2-5A system as one independent element in a 93. Vandenbussche, P., Content, J., Lebleu, B . , and Werenne, J. (1978). J . Gen. Virol. 41, 161-166.

94. Nilsen, T. W., Wood, D. L . , and Baglioni, C. (1980). Nrrtrrre (London) 286, 178-180. 95. Nilsen, T. W., Wood, D. L., and Baglioni, C. (1981). Virology 109, 82-93. 96. Maheshwari, R . K . , and Friedman, R . M. (1980). Virology 101, 399-407. 97. Maheshwari, R . K . , Banejee, D. K . , Waechter, C. J., Olden, K . , and Friedman, R . M.(1980). Nnrirre (London) 287, 454-456.

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multicomponent defense mechanism by which interferon can protect cells against virus infection, The contribution of each element to the overall antiviral state appears to vary from one virus-cell system to another. Interferon has several effects on cells that are distinct from its ability to promote virus resistance (16). For example, interferon treatment enhances the sensitivity of certain cells to the cytotoxic effects of dsRNA (98), and it seems probable that 2-5A synthetase and/or the protein kinase are involved in this effect. In addition, interferon-treated cells usually divide less frequently than untreated cells (16). One variant line of human fibroblasts has been described that is resistant to this antiproliferative effect while retaining normal sensitivity to antiviral effects of interferon (99). Both 2-5A synthetase and the phosphokinase show normal inducibility in these cells, indicating that these enzymes are not sufficient to mediate the interferon’s antiproliferative effects.

E. ROLEOF CELLS

THE

2-5A SYSTEM

IN

NON-INTERFERON-TREATED

Various components of the 2-5A system can be detected in noninterferon-treated cells. The varied basal levels of 2-5A synthetase are described in Section I 1 , B . The 2-SA-dependent nuclease is also present and its level is affected little, if at all, by interferon treatment. Last, 2-5A itself has been detected in acid extracts of non-interferon-treated cells after infection by EMC virus. (52,5.3).These observations raise the possibility that the 2-5A system plays a role in the control of cellular metabolism that is distinct from its role(s) in interferon action. Indeed, teleological arguments would lead one to predict that this should be the case. Exactly what the role is, however, remains a matter for speculation. Messenger RNA degradation is the most obvious possibility, and the variations of 2-5A synthetase levels with cell growth and hormone status are consistent with this proposal. Moreover, the complexity of the system and its apparent potential for delicate control are attributes that would be expected in a process as crucial as messenger RNA degradation. However, the inactivation of a specific messenger RNA, that for human fibroblast interferon, was unaffected by 200-fold variations in the level of latent 98. Stewart, W. E. , De Clercq, E . , Billiau, A., Desrnyter, J . , and De Somer, P. (1972). P N A S 69, 1851-1854. 99. Vandenbussche, P., Divizia, M., Verhaegen-LewaHe, M., Fuse, A., Kuwata, T., DeClercq, E . , and Content, J. (1981). Virology 111, 11-22.

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313

2-5A synthetase activity (100). But as noted in section IV,B, synthetase activity in uninfected cells appears to be limited by the availability of dsRNA activator, rather than by the supply of latent enzyme, so these results do not exclude the 2-5A system from a role in messenger RNA degradation. As before, definitive answers must await the isolation of mutant cells that lack the various components of the system.

100. Sehgal, P. B.,and Gupta, S. L. (1980). PNAS 77, 3489-3493.

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Section 111

RNA Nucleases and Related Enzymes

This Page Intentionally Left Blank

Pancreatic R ibmuclease PETER BLACKBURN

STANFORD MOORE

I . Introduction . . . . . . . . . . . . . . . . . . . . . . . . . I1 . Preparation . . . . . . . . . . . . . . . . . . . . . . . . . 111. Chemical Properties . . . . . . . . . . . . . . . . . . . . . . A . Modification of Functional Groups . . . . . . . . . . . . . . B . Roles of Residues Near the NHZand COOH Termini . . . . . C . Chemical Synthesis . . . . . . . . . . . . . . . . . . . . D . Biosynthesis . . . . . . . . . . . . . . . . . . . . . . . E . Immunochemistry . . . . . . . . . . . . . . . . . . . . . IV. Physical Properties . . . . . . . . . . . . . . . . . . . . . . A . X-Ray Diffraction . . . . . . . . . . . . . . . . . . . . . B . Nuclear Magnetic Resonance . . . . . . . . . . . . . . . . C . Optical Properties . . . . . . . . . . . . . . . . . . . . . D . TheFoldingPathway . . . . . . . . . . . . . . . . . . . . V. Species Variations . . . . . . . . . . . . . . . . . . . . . . . A . Variations in Amino Acid Sequence . . . . . . . . . . . . . B . Variations in Carbohydrate Moieties . . . . . . . . . . . . . VI . Bovine Seminal Plasma RNase . . . . . . . . . . . . . . . . . VII . Cytoplasmic RNase Inhibitor . . . . . . . . . . . . . . . . . . A . Purification and Chemical Properties . . . . . . . . . . . . . B . Studies on in Virro Protein Synthesis . . . . . . . . . . . . VIII Catalytic Properties . . . . . . . . . . . . . . . . . . . . . . A.Assay . . . . . . . . . . . . . . . . . . . . . . . . . . B . Inhibitors and Activators . . . . . . . . . . . . . . . . . . C.Kinetics . . . . . . . . . . . . . . . . . . . . . . . . . D. Mechanism of Catalysis . . . . . . . . . . . . . . . . . . IX . Research Applications . . . . . . . . . . . . . . . . . . . . .

.

.

1

317 318 320 320 345 359 361 362 364 364 366 382 385 397 398 407 411 416 416 423 424 424 426 428 430 433

Introduction

Bovine pancreatic ribonuclease. the first enzyme for which the chemical structure could be written. has been the subject of extensive structure-function studies . The literature to 1970 was reviewed by 3 17 THE ENZYMES. VOL . XV Copyright @ I982 by Academic Press. Inc . All rights of reproduction in any form reserved . ISBN C-12-122715-4

3 18

PETER BLACKBURN AND STANFORD MOORE

Richards and Wyckoff ( I ) in Volume IV of this series. The present chapter covers some of the research on the enzyme in the subsequent decade. A characteristic of the research since 1970 is that it has broadened in scope. The subject has been reviewed within the context of mammalian nucleolytic enzymes by Sierakowska and Shugar ( 2 ) . When the structural work on RNase was begun in the 1950s, the enzyme was viewed as a catalyst of rather limited physiological interest; it was recognized as one of the enzymes of the digestive tract. Neutral RNases of similar molecular design are now known to be present in many tissues, and the specific cytoplasmic inhibitor of enzymes of this type has been characterized in considerable detail. The basic chemical information on the bovine pancreatic enzyme has facilitated studies of the biochemistry of a number of members of the class of catalysts (EC 3.1.27.5) defined as those which, at near neutral pH, cleave RNA endonucleolytically to yield 3’-phospho-, mono-, and oligonucleotides ending in Cp or Up, with 2’,3’-cyclic phosphate intermediates. In most of this review the term RNase refers to bovine pancreatic RNase A; in the extension of the discussion to other members of the series, the enzyme is defined in context. RNase S refers to the enzyme cleaved primarily between residues 20 and 21 by subtilisin ( I ) . RNase A, an unusually well-defined enzyme, has been a test protein in the study of a wide variety of chemical and physical methods of protein chemistry. The volume of the literature has necessitated a selection of topics for this chapter. II. Preparation

The starting product for the chromatographic purification of bovine pancreatic RNase has usually been the enzyme prepared as described by Kunitz and McDonald ( 3 ) . The previously reviewed ( 1 ) methods of ion exchange chromatography have proved effective in yielding homogeneous preparations of the protein, including the separation of RNases that differ in the extent of glycosylation. Such a fractionation is exemplified by the isolation of ovine RNases from pancreatic juice by Becker et a / . ( 4 ) . The introduction of affinity chromatography has simplified the isolation of pure RNases by providing a highly efficient method for separating active enzymes from molecules that do not have an affinity for the coupled substrate analog; the technique can be used as an early step in purificaI. 2. 3. 4.

Richards, F. M., and Wyckoff, H. W. (1971). “The Enzymes,” 3rd. ed., Vol. IV, p. 647. Sierakowska, H . , and Shugar, D . (1977). f r o g r . Nircleic Acid Rrs. M o l . B i d . 20, 59. Kunitz, M., and McDonald, M. R . (1953). Biocliem. f r e p . 3, 9. Becker, R. R., Halbrook, J. L., and Hirs, C. H. W. (1973). JBC 248, 7826.

12. PANCREATIC RIBONUCLEASES

3 19

tion or as a final step after ion exchange chromatography and gel filtration have been used to isolate an RNase fraction of given charge and size. Wilchek and Gorecki ( 5 ) introduced the use of 5’-(4-aminophenylphosphoryl)uridine-2’(3‘)-phosphate-Sepharose4B (pup-Sepharose) for this purpose; the purified enzyme was adsorbed at pH 5.2 in 0.02 M sodium acetate buffer and was eluted with 0.2 M acetic acid. Under these conditions, some subsequent users of the method have observed adsorption of proteins other than RNase and difficulty in the elution of the enzyme with acetic acid. Both of these problems have been overcome by maintaining a sufficient cation concentration in the loading buffer and in the eluent to nullify the properties of the acidic adsorbent as a nonspecific cation exchanger. Stewart and Stevenson ( 6 ) , in the course of preparing bison RNase, found that the positively charged heterocyclic base piperazine was preferable to Na’ for reduction of extraneous protein adsorption: they added the sample in 0.025 M piperazine-HC1 buffer at pH 5.3. Elution was with 0.25 M sodium phosphate buffer at pH 3: phosphate was chosen for the eluent since this anion has an appreciable affinity for RNase. In their isolations of pancreatic RNase from a variety of animal species, Beintema and his associates ( 7 - 1 0 ) have generally used acid extraction and (NH&S04 precipitation and applied the RNase-containing solution to pup-Sepharose in 0.23 M acetate buffer at pH 5.2; elution was with the same buffer, 4 M in NaC1. Alternatively, a linear salt gradient from 0.2 to 6 M NaCl was used for elution. In the isolation of human pancreatic RNase, Weickmann P t ul. (I I ) combined acetone precipitation, chromatography on phosphocellulose, and adsorption on pup-Sepharose. A long column (0.4 x 72 cm) of pup-Sepharose was used by Wang and Moore (12) to remove RNase from preparations of pancreatic DNase; the RNase content was reduced to less than 1 part per 10 million. Smith ef d. (13) used uridine 5’-triphosphate-hexane-agaroseas an affinity adsorbent with the same buffer systems as used by Beintema et ul. They found that a change to pH 5.5 for the eluent buffer gave slightly 5. Wilchek, M . , and Gorecki, M. (1969). EJB 11, 491. 6. Stewart, G. R., and Stevenson, K . J. (1973). BJ 135, 427. 7. Wierenga, R . K . , Huizinga, J. D . , Gaastra, W., Welling, G. W., and Beintema, J. J. (1973). FEES L e t f . 31, 181. 8. Gaastra, W., Groen, G., Welling, G . W., and Beintema, J. J . (1974). FEES L e u . 41, 227. 9 . Gaastra, W., Welling, G. W., and Beintema, J. J. (1978). EJB 86, 209. 10. Havinga, J . , and Beintema, J . J. (1980). EJB 110, 131. 1 1 . Weickmann, J. L., Elson, M., and Glitz, D. G. (1981). B i o c h e r n i s r ~ .20, 1272. 12. Wang, D., and Moore, S . (1978). JBC 253, 7216. 13. Smith, G. K., Schray, K. J., and Schaffer, S . W. (1978). A n d . Biochem. 84, 406.

320

PETER BLACKBURN AND STANFORD MOORE

sharper elution. The 5’-UTP derivative is less stable than pup-Sepharose, columns of which can be used repeatedly without loss of effectiveness ( 5 ) . Scofield et (11. (14) synthesized N4-(aminohexanoy1aminoheptyl)cytidine 2’(3‘)-phosphate-Sepharose4B by first using a bisulfite-induced transamination to introduce an alkyl diamine at the 4 position of cytidine. The product was compared with pup-Sepharose for the chromatography of RNase. The two adsorbents performed similarly and bound about 5 mg of RNase per milliliter of settled bed. 111.

Chemical Properties

Discussion of the results of chemical modification of bovine pancreatic RNase is made with reference to the sequence (15) in Fig. 1. The geometry is considered in reference to the three-dimensional structure derived by Richards and Wyckoff ( 1 , 16) for RNase S (Fig. 2,Ref. 160). A. MODIFICATION O F FUNCTIONAL GROUPS 1. Amino Croirps

Many of the reactions that modify the lysine residues of RNase A have been summarized in Table VI of Richards and Wyckoff’s review ( I ). Of the ten lysine residues, Lys-41 has been placed at the active center of the enzyme by both chemical and physical studies (1, 16-21); Lys-7 is nearby. Pyridoxal phosphate has been found to form a Schiff base with the c-NH2group of either Lys-41 or Lys-7 ( 2 2 , 2 3 )and with the a-NH2group of Lys-1 (24); reduction of the Schiff bases with borohydride yields stable adducts. The products of the reaction were separated and identified by 14. Scofield, R. E., Werner, R. P., and Wold, F. (1977). A n d . Biochem. 77, 152. 15. Smyth, D. G., Stein, W. H., and Moore, S. (1963). JBC 238, 227. 16. Richards, F. M., and Wyckoff, H. W. (1973). In “Atlas of Molecular Structures in Biology,” Vol. I , Ribonuclease S (D. C. Phillips and F. M. Richards, eds.), p. 1. Clarendon, Oxford. 16a. Cantor, C. R., and Schimmel, P. R. (1981). “Biophysical Chemistry,” p. 930. Freeman, San Francisco, California. 17. Glick, D. M., and Barnard, E. A. (1970). BBA 214, 326. 18. Brown, L. R., and Bradbury, J. H. (1975). EJB 54, 219. 19. Brown, L. R., and Bradbury, J . H. (1976). EJB 68, 227. 20. Jentoft, J . E . , Jentoft, N.,Gerken, T. A., and Dearborn, D. G. (1979).JBC 254,4366. 21. Wodak, S . Y., Liu, M. Y.,and Wyckoff, H. W. (1977). JMB 116, 855. 22. Means, G. E. , and Feeney, R. E. (1971). JBC 246, 5532. 23. Raetz, C. R. H., and Auld, D. S. (1972). Biochemistry 11, 2229. 24. Riquelme, P., Brown, W. E., and Marcus, F. (1975). Int. J . Peptide Protein Res. 7, 379.

4

3 2

fi

5

6

7

9

8

10

12

11

13

14

16

15

17

18

19

20

A l a - A l a ~ . y s ~ ~ e - G 1 ~ - * ~ ~ ~ & Z - H i s -Asp M e.ct ~er?fje

Ala t f

Tbr t Glu

r"b

+"pGlu

-

75

Ser- 9 r -

t JiYS

nnl -

55

-

115

l-Ala+ Cys-Glu-G+p

N n P-

116 117 118 119 120

~ - ? L P - ~ ~ 1 - P r o - i ~ - ~ - ~ ~ ~ ~ ~ - ~ ~

VII

100

48

47

46

45

44

43

42

41

40

94

39

38

37

36

35

34

33

11 FIG.1 . The sequence of amino acid residues in bovine pancreatic RNase A. From ( I S ) , based upon the combined researches of the several laboratories referred to therein.

FIG.2. Three-dimensional structure of RNase S, based upon the data of Richards and Wyckoff (I). From (160); illustratian caavrieht. Irvine Geiss.

12. PANCREATIC RIBONUCLEASES

323

peptide mapping and amino acid-analyses. They had 0, 17, and 58% of the activity of RNase, respectively. Pyridoxal phosphate is a competitive inhibitor with respect to the substrate cyclic 2‘ ,3’-CMP (23) whereas pyridoxal itself does not react with the enzyme (22). Circular dichroism measurements in the far-UV and thermal transition profiles measured by CD suggest that alkylation of Lys-7 or Lys-41 via pyridoxal phosphate does not significantly affect the conformation of the molecule (25). Results on the binding of 3’-CMP and the kinetics of the hydrolysis of cycIic 2‘,3‘-CMP agree with those of Riquelme et al. (24) and indicate that the eNH2 group of Lys-7, although located in the region of the active site, is not directly involved in catalysis. The inactive derivative formed by modification with this reagent at the e-NH2 group of Lys-41, however, no longer binds nucleotides (25); RNase modified at the +NH2 group of Lys-41 by dinitrofluorobenzene, although inactive, still binds substrate analogues. Pares et ul. (26) have shown that at pH 5.5, 6-chloropurine 5 ’ ribonucleotide monophosphate and 8-bromoadenosine 5’-monophosphate bind specifically to RNase with affinities similar to those of 5’-AMP and 5’-GMP. At pH 7.3 and 40°, specific alkylation of the m-NHz group of Lys-1 was observed with a 60-fold molar excess of the 6-chloro derivative (27). The modified enzyme is only slightly less active than RNase. The authors suggest (28) that their results are compatible with a third basebinding site on RNase A. From X-ray studies on RNase complexes with analogues of UpA ( I ) and CpA ( 2 I ) , the binding is considered to have at least 5 centers, Bl, R1,pl, Rz, B2 (B and R for base and ribose). Since p u p binds more strongly than 2’(3‘)pU, Irie and associates ( 2 9 , 2 9 0 )propose the existence of a po site to account for the influence of the 5’-phosphate. The binding of ApUp and GpCp is stronger than that of GpC (30); Pares et al. (28) conclude that they are observing an extension of the binding site to include pz, R3, and B3 in the vicinity of Lys-I. Reductive alkylation of RNase A with formaldehyde (31-33) has been 25. Dudkin, S. M., Karabachyan; L. V., Borisova, S. N., Shyiapnikov, S. V., Karpeisky, M. Ya., and Geidarov, T. G. (1975). BBA 386, 275. 26. Pares, X., Arus, C . , Llorens, R . , and Cuchillo, C. M. (1978). BJ 175, 21. 27. Pares, X., Puigdomenech, P., and Cuchillo, C. M. (1978).Int. J . Peptide Protein Res. 16, 241. 28. Pares, X . , Llorens, R., Arus. C., and Cuchillo, C. M. (1980). EJB 105, 571. 29. Sawada, F., and Irie, M. (1969). J . Biochem. (Tokyo) 66, 415. 29a. Mitsui, Y., Urata, Y., Torri, K . , and Irie, M. (1978). BBA 535, 299. 30. White, M. D., Bauer, S., and Lapidot, Y. (1977). Nucleic Acids R e s . 4, 3029. 31. Means, G. E., and Feeney, R. E. (I%@. Biochemisfry 7,2192. 32. Pa&, W. K . , and Kim, S. (1972). Biochemistry 11, 2589. 33. Means, G. E. (1977). “Methods in Enzymology,” Vol. 47, p. 469.

324

PETER BLACKBURN AND STANFORD MOORE

applied with substitution of borohydride by cyanoborohydride, which is more specific for the reduction of Schiff bases and improves the efficiency of alkylation (34).Borohydride will reduce disulfide bonds and can cleave peptide linkages (35);cyanoborohydride does neither, and can be used at physiological pH (34). Cyanoborohydride tends to favor dialkylation of the protein NH2 groups (36). Reductive methylation of proteins permits the lysine residues to be studied by NMR, either from the proton resonances of the N-methyl groups (18, 19) or from the 13Cresonances of the N-methyl groups when [ 13C]formaldehydeis used for the modification (20, 34). Such studies with RNase A indicate that the fully modified protein retains its native conformation, and that Lys-41 is the only lysine residue that the chemical shift alters on binding of phosphate (18, 19) or 3’-CMP (20). Ligand binding was found to perturb the p K, of dimethylated Lys-41 (37). Feeney and his associates (36) have demonstrated the reversible reductive alkylation of RNase A and other proteins by use of the a-hydroxyaldehyde or ketone compounds, glycolaldehyde and acetol. Reductive alkylation of the monosubstituted amine is reversed by periodate oxidation to yield the primary amine; the dialkyl derivative is not labile to periodate oxidation. Bello et al. (38, 39) have studied the reaction of the arylating reagent 2-carboxy-4,6-dinitrochlorobenzene(CDNCB) with a number of model compounds and RNase A. The reagent reacts with imino, amino, and sulfhydryl groups; at pH 8.2 sulfhydryl groups react much faster than amino groups. With RNase A, CDNCB reacts preferentially with the e N H 2group of Lys-41 at 450 times the rate it reacts with the e N H 2group of a-N-acetyllysine. The CDNP-derivatives have absorption spectra typical of nitroanilines with A,, at 368-370 nm at pH 7.0, and at 345-350 nm in 0.1 M HCl. With RNase A, there was a small amount of product modified only on the a-NH2 group (7%); this product was fully active toward yeast RNA. Reaction at e N H 2 groups other than that of Lys-41 was not observed, even with a 6-fold molar excess of reagent over the enzyme concentration. The product modified at Lys-41 had only 0.6% of the activity of native RNase A. Reaction of CDNCB with N- 1-carboxymethyl-His-119-RNase A was much slower than with the native enzyme. The anionic carboxymethyl 34. 35. 36. 37. 38. 39.

Jentoft, N., and Dearborn, D. G. (1979). JEC 254, 4359. Crestfield, A. M., and Moore, S. (1963). JEC 246, 831. Geoghegan, K. F., Ybarra, D. M., and Feeney, R. E. (1979). Biochemistry 18,5392. Jentoft, J. E., Gerken, T. A., Jentoft, N., and Dearborn, D. G. (1981). JEC 256,231. Bello, J., Iijima, H . , and Kartha, G. (1979). Int. J . Peptide Protein Res. 14, 199. I&na, H . , Patrzyc, H., and Bello, J . (1977). EBA 491, 305.

12. PANCREATIC RIBONUCLEASES

325

group on His-119 is presumed to inhibit binding and/or orientation of CDNCB at the active site. The reaction at Lys-41 is inhibited by 2'(3')UMP (38). CDNP-RNase. has been crystallized in the presence of phosphate. X-Ray diffraction data were collected at 3 A resolution. The difference electron density map indicates no overall change in the protein conformation (38). The CDNP group is situated in the active site but does not directly occupy the pyrimidine or ribose binding sites; it is situated in the wider space leading to the substrate binding region, in the same general region as the DNP group of DNP-Lys-41-RNase S (40). The a-NH2 group is a weaker nucleophile than the e N H 2 group of Lys-4 1. The low p K , of the a-NH2group results from the inductive effect of the peptide carbonyl group. The e N H 2group of Lys-41 has a lowered pK, as a result of neighboring positive charges; Carty and Hirs (41) suggested that a neighboring arginine residue was responsible for this shift. Similarly, Migchelsen and Beintema (42) propose that the higher p K , values, obtained by proton NMR studies, for the active-site histidines of rat pancreatic RNase, as compared to those of bovine RNase A, result from substitution of Arg-39 in the bovine enzyme by Ser-39 in the rat enzyme. These authors (42) also suggest that this substitution explains the much lower rate of reaction of fluorodinitrobenzene with the e N H 2group of Lys-41 of the rat enzyme (43)compared to that of the bovine enzyme. Modification of Arg-39 and Arg-85 by kethoxal (3-ethoxy-2-ketobutanal) reduces the reactivity of Lys-41 to CDNCB (39). Modification of the guanidino groups of these residues by kethoxal lowers the pK, of the group to about 6; the decreased reactivity of Lys-41 with CDNCB at pH 7 and 8 is compatible with an increase in the p K , of the c-NH2 group of Lys-41 by about 1 pH unit (38, 39). Other positively charged groups near the active site include the e N H 2 groups of lysine residues 7 and 66. Walter and Wold (44) have acetylated RNase in the presence of an RNA digest and 2'(3')-CMP substrate analogues. They found that no single lysine residue was protected by the substrate analogues from acylation, and suggested that the sum of the residual amounts of lysine residues 7, 41, and 66 correlated fairly well with the residual enzymatic activity. Their acylation reactions were performed at 4" with an excess of acetic anhydride at pH 8.7 in the presence of 1 M sodium acetate plus 0.5 M borate buffer. 40. 41. 42. 43. 44.

Allewell, N. M . , Mitsui, Y., and Wyckoff, H. W. (1973). J5C 248, 5291. Carty, R. P., and Hirs, C. H. W. (1968). JBC 243, 5254. Migchelsen, C., and Beintema, J. J. (1973). J M B 79, 25. Gold, M . H. (1971). Ph.D. Thesis, SUNY at Buffalo, Buffalo, New York. Walter, B., and Wold, F. (1976). 5iuchemisfr.~15, 304.

326

PETER BLACKBURN AND STANFORD MOORE TABLE I THEPROTECTIVE EFFECTOF PoLY(A) ON THE AMIDINATION OF LYSINE RESIDUES IN RNASEA

Lysine residue Protection afforded (%)"

1 <5

7 51

31 25

37 36

41 100

61 100

66 <5

91

40

98 <5

104 43

" Calculated from the relative recovery of individual peptides and their overlap peptides, the sum of which equals 100%. From Blackburn and Gavilanes ( 4 4 ~ ) . Blackburn and Gavilanes (440) have studied the protective action of poly(A) toward amidination of lysine residues by methyl acetimidate at pH 8.5. Tryptic hydrolysis of the amidinated and performic acid oxidized protein, and separation of the peptides by reversed phase high-pressure liquid chromatography ( 4 9 , permitted identification of the protected lysine residues (Table I). Only two lysine residues were completely protected from amidination, Lys-41 and Lys-61. Other lysine residues were protected to different degrees. In the study by Walter and Wold (44), it was not possible to distinguish between modification of Lys-61 and Lys-66. The results of amidination in the presence of poly(A) show Lys-66 to be fully available. Amidination of lysine residues 7 (49%) and 66 (>95%) occurred with no loss of enzymatic activity; this result demonstrates that a modification of these residues, which retains a positive charge, does not affect activity. Lys-41, however, has a more sensitive role at the active site, since amidination (45-47) or guanidination ( I ) leads to inactivation. RNase A is inactivated by methylthioinosinedicarboxyaldehyde,the periodate oxidation product of /3-~-ribosyl-6-methylthiopurine(48). Inactivation results from formation of a Schiff base, presumably with Lys-41. This reagent is a potent antitumor and immunosuppressive agent that is presumed to act via formation of Schiff bases with essential amino groups of proteins. Inactivation of RNase A after reaction with N-acetoxy-2fluorenylacetamide (N-acetoxy-2-FAA), an activated metabolite of the carcinogen, N-hydroxy-2-fluorenylacetamide, has been reported (49). N-Acetoxy-2-FAA modifies the e N H 2 groups of the protein primarily by acetylation rather than by arylamidation, and results in the inactivation of the enzyme. 44a. Blackburn, P., and Gavilanes, J. G. (1982). JBC 257, ,316. 45. Blackburn, P., and Gavilanes, J. G. (1980). JEC 255, 10959. 46. Reynolds, J. H. (1968). Biochemistry 7, 3131. 47. Blackburn, P., and Jailkhani, B. L. (1979). JEC 254, 12488. 48. Spoor, T. C,, Hodnett, J. L., and Kimball, A. P. (1973). Cancer Res. 33, 856. 49. Barry, E. J., and Gutmann, H. R. (1973). JBC 248, 2730.

12. PANCREATIC RIBONUCLEASES

327

The differential reactivities of the amino groups of RNase A have been demonstrated by reaction with phthalyl-4-isothiocyanate (50). At p H 7.2 the reaction is specific for the w N H 2 group of the enzyme. The adjacent carboxyl groups of the phthalyl group act as a strong binding site for lanthanides. Bradbury et al. (50) propose to use this derivative in NMR studies of the NH2 terminus of RNase A. Garel (51) has coupled fluorescein-isothiocyanate to the a-NHzgroup of RNase A by reaction at pH 6. The pK, of the fluorescein group, initially at 6.2, is sensitive to conformational changes in the protein. The molar absorption coefficient for the derivative at 495 nm, is 72,000 cm-' A ! - ' ; the maximum change in molar absorbance at 495 nm upon titration of the fluorescein group, k 4 9 5 is 50,000 cm-' M - I . The change in €495 due to change of the p K, of the fluorescein reporter group has been used to study the binding of 2'-CMP and the thermal unfolding of the protein. Reaction of RNase A with the heterobifunctional reagent ethyl bromoacetimidate has been reported (52). At pH 9.0 rapid amidination of the amino groups occurs with inactivation of the enzyme as a result of reaction with Lys-41. As the pH is lowered, alkylation of a single histidine residue occurs, with cross-linking between Lys-41 and, primarily, His119. The pH optimum for this cross-linking alkylation is at pH 5.6, the same as that exhibited by the reaction of RNase A with haloacetates (cf., I, 53). The dimeric structure of the ribonuclease of bovine seminal plasma (see Section VI) and the cytotoxic properties thereof prompted studies on the preparation of cross-linked dimers of RNase A. The reaction between a diimido ester and the NH2 groups of the protein was studied (54, 55) in terms of the yield of a cross-linked dimer with maximum activity toward poly(A). poly(U). The reaction had been examined earlier by Hartman and Wold (56), primarily in reference to intramolecular cross-linking. Dimer formation was favored (55) at pH 7.7-8.0 at 21" with 1.25 equiv of dimethyl suberimidate and a protein concentration of 6%; the product obtained in 20% yield had 19 unmodified NH2 groups out of a theoretical 20 for a dimeric molecule, in which 2 such groups are involved in the crosslinking. The activity of the cross-linked dimer (55) toward poly(A). poly(U) in 0.14 M salt was 8.5 times that of the monomeric enzyme toward

50. 51. 52. 53. 54. 55. 56.

Bradbury, J. H.. Howell, J. R., Johnson, R. N . , and Warren, B. (1978). EJB 84,503. Garel, J.-R.(1976). WE 70, 179. Diopoh, J . , and Olomucki,M. (1979). Hoppe-Seyler's Z . Physiol. C h e m . 360, 1257. Plapp, B. V. (1973). JBC 248, 4896. Bartholeyns, J., and Moore, S. (1974). Science 186, 444. Wang, D., Wilson, G . , and Moore, S. (1976). Biochemistry 15, 660. Hartman, F. C., and Wold, F. (1967). Biochemistry 6, 2439.

328

PETER BLACKBURN AND STANFORD MOORE

the same substrate. The dirneric derivative has been studied in terms of its tumorostatic properties (57, 58). The cross-linking reaction has been extended to the preparation of poly-sperrnine-RNase (59) (see Sections VI and VIILB). Wang (60) prepared the bifunctional enzyme RNase-DNase by a procedure similar to that introduced by King and Kochoumain (61). The initial step with each enzyme was thiolation by reaction of NH2 groups with N-acetylhornocysteine thiolactone. The SH group added to DNase was covered by preparing the derivative in the presence of 4,4’dithiodipyridine. The cross-linkage was accomplished by thiol-disulfide interchange. The hybrid enzyme (yield 25-33%, based upon the DNase used) contained one molecule each of DNase and RNase cross-linked by one - S - S bridge; the combination had 75 and 40% of the activities of the parent enzymes, respectively, toward DNA and RNA. Reductive glycosarnination of lysine residues can be used to obtain derivatives with carbohydrate moieties attached to NH2groups. The conditions described by Gray (62) for the coupling of oligosaccharides to proteins in the presence of cyanoborohydride have been used by Wilson (63, 64) to attach lactose or rnannobiose to RNase cross-linked dimer. In 24 hr at pH 7 and 37”, in the presence of phosphate to protect Lys-41, the disaccharide was attached to an average of five lysine residues per dimer without significant decrease in enzymic activity. The disaccharides were chosen in order to obtain derivatives [e.g., N‘- 1-(l-deoxylactitolyl-lysproteins] with terminal galactopyranose or mannopyranose rings to study the respective selective uptakes (63, 6 4 ) of injected synthetic glycoconjugates by the known receptors for the given sugars in hepatocytes or cells of the reticuloendothelial system. Baynes and Wold (65) conducted uptake studies with the naturally occurring bovine RNases B, C, and D, which differ in the carbohydrate side chains attached to Asn-34 (66, 67). Biondi 57. Tarnowski, G. S., Kassel, R. L., Mountain, I. M., Blackburn, P., Wilson, G., and Wang, D. (1976). Cuncer Res. 36, 4074. 58. Bartholeyns, J., and Zenebergh, A. (1977). B t r . J . Currcer 15, 85. 59. Wang, D., and Moore, S. (1977). Biochemistry 16, 2937. 60. Wang, D. (1979). Biochemistry 18, 4449. 61. King, T. P., Li, Y.,and Kochoumian, L. (1978). Biochemistry 17, 1499. 62. Gray, G. R. (1974).A B B 163, 426. 63. Wilson, G. (1978). JBC 253, 2070. 64. Wilson, G.(1979). J . Gen. Ptiysiol. 74, 495. 65. Baynes, J. W., and Wold, F. (1976). JBC 251, 6016. 66. Plummer, T. H., Jr., and Hirs, C. H. W. (1%4). JBC 239, 2530. 67. Plummer, T. H . , Jr. (1968). JBC 243, 5961.

12. PANCREATIC RIBONUCLEASES

329

et 01. (68)acylated the e N H 2groups of RNase with gluconyl-glycine azide to introduce polyhydroxyalkyl side chains. 2. Histidin e Residues The initial studies on modification of the imidazole side chains of histidine residues of RNase A have been reviewed by Richards and Wyckoff (I). In an extension of the studies on the reaction of bromoacetate with the enzyme, Lennette and Plapp (69, 70) showed that carboxymethylation of N-3 of His-12 and N-1 of His-119 occurs, respectively, 120 and 1000 times faster than with the corresponding imidazole nitrogens of histidine hydantoin (70). The increased rate of alkylation of the active-site histidines of RNase A by bromoacetate, compared to the carboxymethylation of histidine hydantoin, corresponds to a difference in the free energy of activation (AC) of 3.2 to 4.2 kcaYmo1. This results primarily from a decreased enthalpy of activation (AH of 5.5 kcal/mol) and not from the entropy of activation (AS). From crystallographic studies on RNase S, the imidazole side chain of His-119 can occupy at least four different positions in the crystal structure, depending on the nature of ligands bound at the active site ( I ) . Lennette and Plapp (70) suggest that bromoacetate binds to a particular conformation of the enzyme and stabilizes the position of the imidazole of His-119, permitting a hydrogen bond between the /3-carboxyl of Asp-121 and N-3 of His-1 19. In this way, the nucleophilicity of N-1 of His-1 19 would be greatly increased by the inductive effect resulting from the hydrogen bond with N-3 of His-1 19. Such a hydrogen bond was postulated to exist by Sacharovsky et ( I / . (71) to explain the increased pK, values of the active-site histidine residues in des-( 121- 124)-RNase A, and has similarly been invoked by Antonov et al. (72)to fix the positively charged imidazole of His-119 into the catalytically active position in the complex between RNase S and 3' ,5'-2-deoxy-2-fluoro-UpA (73). Santoro et nl. (74) found no evidence for a hydrogen bond between 68. Biondi, L . , Filira, F., Giorrnani, V., and Rocchi, R. (1980). f n t . J . Pepptide Protein Res. 15, 253.

69. Lennette, E . P., and Plapp, B. V. (1979). Biochrmis/r.v 18, 3833. 70. Lennette, E. P., and Plapp, B . V. (1979). Eiorlirmistry 18, 3938. 71. Sacharovsky, V. G., Chervin, I. I., Yakovlev, G. I., Dudkin, S. M . , Karpeisky, M. Ya., Shliapnikov, S . V., and Bystrov, V. F. (1973). FEES Lett. 33, 323. 72. Antonov, I . V., Gurevich, A. Z., Dudkin, S. M . , Karpeisky, Y. Ma., Sacharovsky, V. G . , and Yakovlev, G. I. (1978). EJB 87, 45. 73. Pavlovsky, A . G . , Borisova, S. N . , Borisov, V. V., Antonov, I . V., and Karpeisky, M . Ya. (1978). FEES Lett. 92, 258. 74. Santoro, J . , Juretschke, H. P., and Ruterjans, H . (1979). BEA 578, 346.

330

PETER BLACKBURN AND STANFORD MOORE

Asp-121 and His-1 19 in their NMR studies of the carboxyl groups of RNase A. Walters and Allerhand (75)found that only His-119 exists in the Nc2-H (i.e,, N-3-H) tautomeric form most commonly found for nonhydrogen-bonded histidine residues. Their 13C NMR studies were performed in 0.2 M acetate. It seems unlikely that bromoacetate or iodoacetate binds to the active site of RNase A much differently than acetate. Indeed, acetate competitively inhibits the alkylation of the active-site histidine residues (70). The proximal positioning of the imidazole side chain of His-1 19 near the phosphate of dinucleotide phosphates appears to result primarily from binding of the base at the purine binding site ( 2 0 . While it is tempting to invoke a hydrogen bond between N-3 of His-119 and the P-carboxyl of Asp-121 to explain the nucleophilicity of N-1 of His-119, no direct NMR evidence for such a hydrogen bond exists. (See neutron diffraction studies, Section IV,A.) Alkylation of the active-site histidine residues of RNase A with iodoacetamide occurs preferentially with His-12, although the rate of alkylation is 7 times slower than that with iodoacetic acid (76). The rate of inactivation by iodoacetamide shows a bell-shaped pH dependence, with midpoints at pH 3.8 and 6.2. It has been proposed from NMR studies that RNase A undergoes a pH-dependent conformational transition (77-79) involving hydrogen bonds between the P-carboxyl group of Asp-14 and Tyr-25 or His-48 (74, 80), with the midpoints of the transitions at pH values 4.2 and 6.2. Such a conformational change, which affects the locale of His- 12, might explain the pH-dependent alkylation with iodoacetamide. The preferential alkylation of N-3 of His-12 by the neutral iodoacetamide (76)is consistent with the respective p K, values of the active-site histidine residues, 5.8 for His-12 and 6.2 for His-119 (Table VIII, Section IV,B). Clearly, the negative charge on the carboxyl group of iodoacetate or bromoacetate is an important factor in determining both the rate and selectivity of the alkylation of the active-site histidine residues of RNase A. Orientation and facilitation of alkylation of the active-site histidine residues by bromoacetate or iodoacetate does not seem to depend upon the positively charged e N H 2 group of Lys-41. Modification of Lys-41 with iodoacetate or iodoacetamide does not substantially inhibit the alkylation 75. Walters, D. E., and Allerhand, A . (1980). JBC 255, 6200. 76. Fruchter, R. G., and Crestfield, A. M. (1967). JBC 242, 5807. 77. Ruterjans, H., and Witzel, H. (1969). EBJ 9, 118. 78. Patel, D. J . , Camel, L. L . , and Bovey, F. A. (1975). Biopdymers 14, 987. 79. Cohen, J. S., and Shindo, H. (1975). JBC 250, 8874. 80. Lenstra, J. A . , Bolscher, B. G . J. M., Stob, S., Beintema, J . J . , and Kaptein, R. (1979). EJB 98, 385.

12. PANCREATIC RIBONUCLEASES

33 1

of the active-site histidine residues even though it does inactivate the enzyme (81). The selectivity of the alkylation of RNase A with bromoacetate at pH 5.5 decreases with prolonged times of reaction (1 to 42 days) (82). Alkylation of residues proceeds in the following sequence: (1) N-1 of His-119; (2) Met-30; (3) N-3 of His-12; (4) N-3 of His-105 and N-3 of N1-carboxymethyl-His-119;and ( 5 ) a-NH2of Lys-1. Both His-12 and His119 of the same active site are reported to be ultimately alkylated. Pincus and Carty (83) showed that treatment of RNase A with 2’(3’)0-bromoacetyluridine results in carboxymethylation of the active site histidine residues. The ratio of N-3-carboxymethyl-His- 12 to NI-carboxymethyl-His- 119 products is approximately 6 : 1 (83, 84). The carboxymethyl derivatives arise from the rapid hydrolysis of the parent uridine carboxymethyl RNase A ester (83). Binding of the nucleoside portion of the alkylating ligand orients the bromoacetyl moiety in such a way that it lies close to N-3 of His-12. The small amount of alkylation of N-1 of His-1 19 of RNase by 2’(3’)-0-bromoacetyluridinemay arise from hydrolysis of the ester bond generating some bromoacetate, which then preferentially alkylates His- 119. With 2‘-bromoacetamido-2’-deoxyuridine at pH 5.5, RNase A reacts rapidly with absolute specificity for N-3 of His-12 (85). A comparison of the rate constants for the alkylation of RNase A and free L-histidine with 2’(3’)-0-bromoacetyluridine,2‘bromoacetamido-2’-deoxyuridine,bromoacetic acid and iodo- and bromoacetamide (86) are shown in Table 11. Taking into account the difference in the relative rates of reaction of bromoacetamide and 2’(3’)-0bromoacetyluridine with free L-histidine, the enhancement of the rate of alkylation that results from nucleoside binding to the active site is about a factor of 25. The overall rate of reaction of 2’(3’)-0-bromoacetyluridine with RNase A is 3100 times the rate with free L-histidine; thus, the nucleophilicity of the imidazole N-3 of His-12 is increased approximately 120-fold (86), a value similar to that obtained by Lennette and Plapp (70). Ferrate ion is a powerful oxidant and an analog of phosphate (87); ferrate has been demonstrated to inactivate RNase A as a result of reaction at His-119 (88). The specificity of the reaction is pH-dependent. At 81. 82. 83. 84. 85. 86. 87. 88.

Heinrikson, R. L. (1966). JBC 241, 1393. Bello, J . , and Nowoswiat, E. F. (1971). EJB 22, 225. Pincus, M . , and Carty, R. P. (1970). BBRC 38, 1049. Machuga, E . , and Klapper, M. H . (1975). JBC 250, 2319. Lan, L. T., and Carty, R . P. (1972). BBRC 48, 585. Pincus, M . , Thi, L. L . , and Carty, R. P. (1975). Biochemistry 14, 3653. Lee, Y. M . , and Benisek, W. F. (1976). JBC 251, 1553. Steczko, J . , Walker, D. E . , Hermodson, M., and Axelrod, B. (1979). JBC 254, 3254.

332

PETER BLACKBURN AND STANFORD MOORE TABLE I1 COMPARISON OF THE KINETIC CONSTANTS FOR T H E ALKYLATION OF RNASEA A N D FREEL-HISTIDINE’

Reaction

2’(3’)-O-Bromoacetyluridine+ RNase A 2’(3’)-O-Bromoacetyluridine+ L-histidine 2’-Bromoacetamido-2‘-deoxyuridine + RNase A 2’-Bromoacetamido-2’-deoxyuridine + L-histidine Bromoacetic acid + RNase A Bromoacetic acid + L-histidine Iodoacetamide + RNase A lodoacetamide + L-histidine Bromoacetamide + RNase A Bromoacetamide + r-histidine

(x

k3/ Ka or krpObs lo4M - ’ sec-’)

403 0.I29 518 0.113 89.0 0.037 0.48 0.0052 1.9 0.015

From Pincus et cil. (86) and references therein. [Reprinted with permission from Biochemistry 14, 3653-3661. Copyright (1975)American Chemical Society.]

pH 5.0 the reaction is specific for His-119; at pH 7.0 two tyrosine and two lysine residues were also modified. The enzyme is protected by active-site ligands from ferrate inactivation. Reaction of His-I 19 with ferrate was followed by specific titration of the histidine residues with diethylpyrocarbonate and measurement of the absorption of the product, ethoxyformylhistidine, at 240 nm (89-91 ). RNase A forms stable charge-transfer complexes between chloropentammineruthenium”’ dichloride and the imidazole moiety of histidine residues (92). Based on the kinetics of incorporation, three of the four histidine residues are reactive; the three derivatives that contain a single ruthenium-histidine complex were separable from other reaction products by ion exchange chromatography (93). These complexes have absorption bands arising from charge-transfer transitions between the imidazole and ruthenium ion, which depend upon the state of protonation of N-1 of the histidine; the protonated form has absorption maxima at 303 and 450 nm, the unprotonated form has absorption maxima at 365 and 600 nm, and the pK, of N-1 of the imidazole in the complex is 8.8 (94). His-105 89. Ovadi, J., Libor, S.,and Elodi, P. (1967).Artu Biochim.Biophys.Acad. Sri. Hung. 2, 455.

90. Melchior, W.B., Jr., and Fahrney, D. (1970).Biochemistry 9, 251. 91. Roosemont, J. R. (1978).Anal. Biochern. 88, 314. 92. Matthews, C. R., Erickson, P. M., Van Vliet, D. L., and Petersheim, M. (1978). JACS 100, 2260. 93. Matthews, C. R., Erickson, P. M., and Froebe, C. L. (1980).BBA 624, 499. 94. Sundberg, R. J., and Gupta, G. (1973).Biuinorg. Chem. 3, 39.

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is the predominant singly labeled derivative; the change in extinction at 370 nm of this derivative during thermal- and urea-induced unfolding of the protein has been followed (93). This derivative had 66% of the activity of native RNase A toward cyclic 2',3'-CMP; the other two singly modified derivatives had 89% and 53% activity, respectively (93). The effects of the paramagnetic ruthenium on the NMR spectrum of the histidine C-2 and C-4 protons have been used to confirm the assignments of these resonances in the derivative labeled at His-105 ( 9 5 ) , and have similarly confirmed His-105 and His-48 as the two histidine residues not fully exposed to solvent in thermally unfolded RNase A (96, 97). The paramagnetic hexacyanochromate [CI-(CN)~]~ion binds to the active site of RNase A and selectivity broadens the C-2 and C-4 proton resonances of His-12 and His-119 (98). 3. Arginine Residues Takahashi (99) reported that 2 to 3 arginine residues of RNase A were modified by reaction with phenylglyoxal at neutral pH and room temperature. The principal residues modified were Arg-39 and Arg-85; activity loss correlated closely with modification of Arg-39. Residues Arg-10 and Arg-33 were generally unreactive. The facilitated alkylation of the activesite histidine residues by iodoacetate was largely unaffected by the modification of the arginine residues. Yankeelov (100) reported that after more than three of the four arginine residues of RNase A were modified by oligomers of 2,3-butanedione, 45% of the activity of the native enzyme, measured at pH 5.2 with RNA as substrate, was retained. Patthy and Smith (101, 102) found that reaction of RNase A with 1,2-cyclohexanedione in borate buffer at pH 8 to 9 results in modification of 2 to 3 arginine residues and 90% loss of enzymatic activity, measured at pH 7.2 with RNA as substrate. Arg-39 reacts rapidly and its modification contributes mostly to the loss of enzymatic activity. Arg-85 also reacts rapidly, Arg-10 reacts slowly, and no reaction was observed with Arg-33. Removal of the blocking groups with hydroxylamine results in complete recovery of enzymatic activity, except when Arg-10 has also been modified, in which case recovery is 80%. Similar results on the loss of enzy95. Matthews, C. R . , Recchia, J . , and Froebe, C. L. (1981). A n d . Biochem. 112, 329. 96. Matthews, C. R . , and Westmoreland, D. G . (1975). Biochemistry 14, 4532. 97. Matthews, C. R., and Froebe, C. L. (1981). Mncromolecules 14, 452. 98. Inagaki, F., Watanabe, K., and Miyazawa, T. (1979). J . Biochem. 86, 591. 99. Takahashi, K. (1968).JBC 243,6171. 100. Yankeelov, J. A., Jr. (1970). Biochemistry 9, 2433. 101. Patthy, L., and Smith, E. L. (1975). JBC 250, 557. 102. Patthy, L., and Smith, E. L. (1975). JBC 250, 565.

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matic activity toward cyclic 2‘,3’-CMP at pH 6.5, and the number of arginine residues modified upon reaction of RNase A with 1,2cyclohexanedione were observed by Blackburn and Jailkhani (47);reaction with 2,3-butanedione, under conditions described by Riordan (103), modified four arginine residues resulting in 95% loss of enzymatic activity (47). Kethoxal (3-ethoxy-2-ketobutanal) reacts with the guanidino groups of arginine residues and to some extent with e N H 2groups of lysine residues (39). The predominant sites of modification were Arg-39, Arg-85, and 85% of Arg-10; no reaction occurred with Arg-33. With two arginine residues modified, Arg-39 and Arg-85, the derivative had 90% of the activity of native RNase A toward RNA at pH 5.0, but only 20 to 25% of the activity of native RNase A toward cyclic 2’,3‘-CMP at pH 7.0. Modification of Arg-10 results in further loss of activity. The pH dependence of the activity of RNase A modified with kethoxal relative to the activity of native RNase A exhibits a titration curve with a midpoint at pH 5.8 in 0.1 ionic strength buffer. Modification of the guanidino group of a-N-acetylarginine with kethoxal lowers the pKa of the modified guanidino group to about 6.0. Modification of the arginine residues by kethoxal had little effect on carboxymethylation of the active-site histidine residues with bromoacetate or with 2’-O-bromoacetyluridine. Reaction of the r-NH2 group of Lys-41 in kethoxal-modified RNase by the arylating reagent 2carboxy-4,6-dinitrochlorobenzeneoccurs at 25 and 2% of the rate with the native enzyme at pH 8.5 and 7.5, respectively, corresponding to an increase in pKa of the r-NH2 group of Lys-41 by 1 pH unit (38, 39). The differences observed in the effects of arginine modification on the activity of RNase, according to Iijima et al. (39),may be explained by the different pH values chosen for assay of the various derivatives. It is possible that the pH dependence of the relative activity of kethoxal-modified RNase A with respect to native RNase is related to the lowered pK, of the modified guanidino groups. Similarly, extensive modification of RNase A with phenylglyoxal yields a protein with a much lower isoionic point, with little or no migration exhibited at pH 6.5 (99). Also, the derivative produced by reaction of arginine with 2,3-butanedione elutes from a short column of sulfonated polystyrene ahead of lysine at pH 5.25. close to the breakthrough of this buffer (103). These results suggest that the pKa of the guanidino groups modified with phenylglyoxal and a,P-diketones is most likely similar to that obtained after modification with kethoxal, i.e., close to 6.0. Iijimaet al. (39) suggest that the decreased activity of kethoxal-modified 103. Riordan, J. P. (1973). Biochemistry 12, 3915.

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RNase A results from the modification of both Arg-39 and Arg-85, and not simply from modification of Arg-39. Arg-39 and Arg-85 are situated near the active site of the enzyme ( 1 ) . Arg-85 is invariant among the species of pancreatic RNases studied (Section V), with the exception of the mouse enzyme in which it is substituted by a histidine residue. Arg-39 is also generally conserved, but is substituted by serine in mouse and rat enzymes, by tyrosine in the muskrat and chinchilla, by lysine in bovine seminal plasma RNase, and is deleted in the RNases of the dromedary and bactrian camels and the horse. The activities of three of these enzymes toward cyclic 2',3'-CMP relative to that of the bovine enzyme (45) are: Mouse, 0.24; rat, 0.52; dromedary, 1.18. The enzyme that exhibits the lowest specific activity has substitutions at both Arg-39 and Arg-85. 4. Aspartate and Asparagine, Glutamate and Glirtamine Residues RNase A has 11 carboxyl groups: 5 aspartate, 5 glutamate, and the a-carboxyl of the terminal Val-124. Modification of these residues in general leads to decreased enzymatic activity, which appears to result from conformational changes that occur with progressive modification of the carboxyl groups ( I ) . Potential interactions involving carboxyl groups, the modification of which could lead to a loss of activity, include the hydrogen bonds between Asp-14 and His-48 or Tyr-25 (74, 7 7 4 0 ) , Glu-2 and Arg-10 (104-106), Val-124 and His-105 ( 7 3 , and possibly Asp-121 and His-119 (69, 71 -73). Glu-1 11 hydrogen bonds to N-1 of the purine base of dinucleotide substrates (2f). Esterification occurs most rapidly at Asp-53 ( 1 07) and Asp-49 (108, 109). The derivative esterified with methanol at both of these residues exhibits no gross conformational change, but the product has only 65% of the activity of the native enzyme toward cyclic 2' ,3'-CMP (109). Although the K, is unaltered, after limited digestion by subtilisin the dimethyl S-protein exhibits a fourfold weaker interaction with the S-peptide, which may explain the reduced activity (109). Modification of Asp-53 with diazoacetyl glycinamide has no effect on the enzymatic activity (107). In 104. Marchiori, F., Borin, G., Moroder, L., Rocchi, R . , and Scoffone, E. (1972). BBA 257, 210. 105. Hofmann, K . , Visser, J. P., and Finn, F. M. (1970). JACS 92, 2900. 106. Hofmann, K . , Andreatta, R., Finn, F. M., Montibeller, J . , Porcelli, G . , and Quattrone, A. J . (1971). Bioorg. Ckem. 1, 66. 107. Riehm, J . P., and Scheraga, H. A. (1965). Biorhemistry 4, 772. 108. Acharya, A. S . , and Vithayathil, P. J. (1975). I n t . J . Pepride Protein R e s . 7 , 207. 109. Acharya, A. S., Manjula, B . N . , and Vithayathil, P. J. (1978). EJ 173, 821.

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the presence of unmodified S-protein, inactive RNase methylated at five carboxyl groups (I 10) exhibits 30% of the activity of the native enzyme; esterification of the carboxyl groups leads to a weaker interaction between the S-peptide and S-protein segments of the RNase molecule and permits hybrid formation. The derivative esterified at Asp-49 and Asp-53 does not show increased activity in the presence of added S-protein (109). In strong acid, RNase A undergoes deamidation. A monodeamidated product accumulates, designated RNase Aa,; subsequent deamidation occurs more slowly (I If). The monodeamidated product has full enzymatic activity; further deamidation is associated with loss of enzymatic activity. Hydrolysis of the monodeamidated derivative with subtilisin gives only 50% conversion to monodeamidated RNase S (I I2). Tryptic hydrolysis and peptide mapping indicates that one of the four amides in the peptide, corresponding to residues 67 through 8 5 , is the primary site of deamidation (111). Spectroscopic and immunological techniques indicated that RNase Aal has a conformation close to that of native RNase A (I I1, 113). The conformation of RNase Aal is, however, less thermostable than that of RNase A, as judged by susceptibility to trypsin and reaction of methionine residues with o-benzoquinone (I 14). 5 . Methionine Residues

RNase A has four methionine residues at positions 13, 29, 30, and 79, which in general are invariant among pancreatic enzymes of different species, except for a few cases where they are conservatively substituted by valine, isoleucine, or leucine. They are buried in the interior of the protein and cannot usually be modified unless the molecule is denatured (I). Alkylation of RNase A with methyl iodide at low pH specifically modifies Met-29, with no effect on the enzymatic activity (115); in 8 M urea at low pH all four methionine residues are alkylated (I I 6 ) . Separation of the reaction products and identification of the sites of modification showed that alkylation at Met-29 or Met-79, or both Met-29 and Met-79 yield fully enzymatically active species upon removal of urea. Their re110. Acharya, A. S., Manjula, B. N., Murthy, G. S., and Vithayathil, P. J . (1977). Inr. J. Peptide Prorein Res. 9, 213. 1 1 1 . Manjula, B . N . , Acharya, S. A . , Vithayathil, P. J . (1976). Inr. J. Peptide Protein Res. 8, 275. 112. Manjula, B . N . , Acharya, A. S. , and Vithayathil, P. J. (1977). BJ 165, 337. 113. Das, M . K . , and Vithayathil, P. J . (1978). BBA 533, 43. 114. Das, M. K . , and Vithayathil, P. J . (1978). i n r . J . Peptide Protein Res. 12, 242. 115. Link, T. P., and Stark, G. R. (1968). JBC 243, 1082. 116. Stark, G. R . , and Link, T. P. (1975). Biochemistry 15, 3476.

12. PANCREATIC RIBONUCLEASES

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spective transition temperatures are 58", 43",and 36", compared to 63" for native enzyme. Methylation of Met-13 or Met-30 prevents refolding to an active conformation. Alkylation of methionine converts the residue to a positively charged sulfonium ion and introduces an alkyl group on the side chain, which can apparently be accommodated only at Met-29 and Met-79. Upon photoxidation of RNase A in the presence of hematoporphyrin in 10% acetic acid, only Met-29 is oxidized to the sulfoxide; at higher concentrations of acetic acid (10 to 50%) Met-13 is also oxidized (117). As with the alkylated derivatives, oxidation at Met-29 does not inactivate the enzyme; the derivative oxidized at both Met-29 and Met-13 had low activity. Modification of Met-13 in S-peptide by oxidation to the sulfone, or alkylation with iodoacetate or iodoacetamide to produce a sulfonium derivative, significantly reduces the strength of binding between S-peptide and S-protein; however, the complexes, when formed, are fully active (1, 118). Richards and Wyckoff suggested that rotation about the a-/3 carbon-carbon bond of Met-13 can allow the charged sulfur atom to have access to solvent without a change in the conformation of the S-peptide (1). In RNase A, the conformation of the S-peptide is more constrained because of the unbroken peptide bond between residues 20 and 21 (119), and it is less likely that the charged sulfur on Met-13 can be similarly accommodated (116). The reaction of o-benzoquinone with methionine at acidic pH produces the 3,4-dihydroxyphenylmethioninesulfonium salt (120).The accessibility of the methionine residues to o-benzoquinone has been used as a probe of protein conformation after esterification of the carboxyl groups of RNase A and RNase S (110). Modification with this reagent introduces a chromophore that enables the number of modified methionine residues to be determined spectrophotometrically (120-122). 6. Disuijide Bonds The four intramolecular disulfide bonds of RNase A are invariant features of the sequence among all of the mammalian pancreatic RNases; they are between residues 26 and 84,40 and 95,58 and 110, and 65 and 72. These four intrachain disulfide bonds contribute to the overall conforma117. Jori, G., Galiazzo, G., Tamburro, A. M., and Scoffone, E. (1970). JBC 245, 3375. 118. Vithayathil, P. J . , and Richards, F. M. (1960). JBC 235, 2343. 119. Carlisle, C . H . , Palmer, R. A . , Mazumdar, S . K., Gorinsky, B. A . , and Yeates, D. G. R. (1974). J M B 85, 1 . 120. Vithayathil, P. J . , and Murthy, G. S. (1972). Nature N e w B i d . 236, 101. 121. Gupta, M. N . , and Vithayathil, P. J. (1980). f n t . J . Peptide Protein Res. 15, 236. 122. Gupta, M. N., Murthy, G. S . , and Vithayathil, P. J. (1980). Znt. J . Peptide Protein Res. 15, 243.

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tional stability of the RNase molecule without determining the final overall conformation of the protein (123). Reduction of the disulfides between residues 65 and 72, and 58 and 110 (124-126) and protection of the sulfhydryls with phosphorothioate (126) yields a species with full activity toward RNA and enhanced activity toward cyclic 2’,3’-CMP. Spectrophotometric titrations (126) and CD measurements (127) show little difference from native conformation. Incubation of RNase A with 0.1 M mercaptoethanol at pH 8.5, upon exposure to the air over a period of days, yields a mixture of chromatographically separable, metastable products with enhanced activity toward cyclic 2’,3’-CMP (128). The fraction with the highest activity (fourfold) had no detectable free sulfhydryl groups; the protein has probably formed mixed disulfides with mercaptoethanol. Under nondenaturing conditions, at pH 8.7 and 25”, Creighton (129) found the half-life for the reduction of native RNase A by 10 mM dithiothreitol to be approximately 10 hours. The primary product was fully reduced RNase A, with no significant accumulation of species with 1,2, or 3 disulfide bonds. Sperling et al. (130) studied the reduction of the disulfide bonds with both dithiothreitol and dithioerythritol. After 1 hour at pH 8.0, with a fivefold molar excess of reducing agent over disulfide, partial reduction of RNase A occurs at the disulfide bond between residues 65 and 72 with no loss of enzymatic activity. The ability to selectively and partially reduce the disulfide bonds of RNase A depends upon the nature and concentration of the reducing agent, the pH, and the folded state of the protein. The ability to form linear mixed disulfides can be important for production of species with enhanced enzymatic activity. Dithiothreitol has little tendency to form mixed disulfides, since its greater reducing potential arises from its ability to form a thermodynamically stable intramolecular, disulfide-linked, six-member ring (131). In the presence of 6 M guanidinium chloride, at pH 8.5 and 25”, RNase 123. Scheraga, H. A. (1980). I n “Protein Folding” (R. Jaenicke, ed.), p. 261. Elsevier, Amsterdam. 124. Sela, M . , White, F. H., Jr., and Anfinsen, C. B. (1957). Science 125, 691. 125. Resnick, H., Carter, J . R . , and Kalnitsky, G. (1959). JBC 234, 1711. 126. Neumann, H., Steinberg, I. Z., Brown, J . R. Goldberger, R. F., and Sela, M. (1967). EJB 3, 171. 127. Tamburro, A. M., Boccu, E., and Celotti, L. (1970). Int. J . Peptide Protein Res. 2, 157. 128. Watkins, J. B., and Benz, F. W. (1978). Science 99, 1084. 129. Creighton, ‘r. E. (1979).J M E 129, 411. 130. Sperling, R., Burstein, Y., and Steinberg, I. 2. (1969). Biochemistry 8, 3810. 131. Cleland, W. W. (1964). Biochemistry 3, 480.

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A is completely reduced by 20 mM 0-mercaptoethanol. Lower concentrations of P-mercaptoethanol only bring about partial reduction of the disulfide bonds under these conditions (132). After net reduction of one disulfide bond and S-alkylation with N-ethylmaleimide, all enzymatic activity is lost. The sulfhydryl groups of reduced RNase react with selenium (133) and mercury (130, 1-34), which can be incorporated into (and elongate) the disulfide bonds. Elongation of the disulfide bond between residues 65 and 72 has no effect on the enzymatic activity (130). Sperling and Steinberg (134)introduced a single mercury atom into all four disulfides of RNase A; the derivative had 25% of the activity of native enzyme toward cyclic 2’,3’-CMP, and 5% toward RNA. The re-formation of RNase disulfides during refolding of the protein is dealt with in Section IV,D. 7. Tvrosine Residues RNase A has six tyrosine residues at positions 25, 73, 76, 92, 97, and 115 in the sequence (15). Early spectrophotometric titrations at alkaline pH suggested that three of the residues titrate with normal pK, values of about 10.2 and three titrate with abnormal ply, values above 12; these phenolic groups were described as “accessible” and “buried,” respectively (1). Subsequent proton (80) and 13C NMR (135) titration studies indicate only two tyrosine residues with abnormal p K, values. Moreover, the X-ray crystallographic data for RNase S ( 1 ) show that only two tyrosine residues are truly buried, residues 25 and 97. Iodination of RNase A permitted identification of the normal and abnormal tyrosine residues (1). Tyr-25 and Tyr-97 are not iodinated; Tyr-92 is iodinated at pH 9.4 (136, 137) but not at pH 6.4 (138), when only residues 73, 76, and 115 are iodinated. Tyr-92, although situated on the surface of the molecule (I), titrates with an abnormally high pK,; its hydroxyl is involved in a hydrogen bond with the carbonyl group of the amide bond between Lys-37 and Asp-38. A local conformational change in the protein may break this hydrogen bond and lead to “normalization” of the behavior of Tyr-92. For example, in horse pancreatic RNase, deletion of residue 39 introduces steric constraints that do not permit the 132. Garel, J.-R. (1977). FEBS Lett. 79, 135. 133. Ganther, H . E . , and Corcoran, C . (1969). Biochemistry 8, 2557. 134. Sperling, R . , and Steinberg, I. Z. (1971). JEC 246, 715. 135. Egan, W., Shindo, H . , and Cohen, J. S. (1978). JBC 253, 16. 136. Cha, C.-Y., and Scheraga, H . A. (1963). JEC 238, 2958. 137. Cha, C.-Y.,and Scheraga, H . A. (1963). JBC 238, 2965. 138. Woody, R . W., Friedman, M . E., and Scheraga, H. A. (1966). Eiodwmistry 5, 2034.

340

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hydrogen bond to involve Tyr-92; Tyr-92 of this protein has a normal pK, (139). Acetylation of the tyrosine residues at pH 7.5 with acetic anhydride or N-acetylimidazole prevents their cleavage by N-bromosuccinimide. By this approach Burstein and Patchornik (140) found that residues 73, 76, and 115 were available to acylation and residues 25, 92, and 97 were unavailable. The availability of the tyrosine residues to modification reactions depends upon the protein’s conformation. In des-( 121- 124)-RNase A, Fujioka and Scheraga (141) found Tyr-25 unavailable to iodination at pH 9.5. The peptide mapping data were not definite on the availability of Tyr-92 and Tyr-97. The CD meaururements of Taniuchi (142) and Puett (143) in the near-UV indicated only a small difference between the conformation of the aromatic residues in RNase A and des-(121-124)-RNase A. Ultraviolet absorption spectra on the native and denatured protein suggest between two and three “buried” tyrosine residues in des-(121124)-RNaseA and two in des-(119-124)-RNase A (144).Tyr-97 is the least accessible tyrosine residue of RNase A; exposure of this residue would require a substantial change in the protein conformation (1 ). Most probably, it is Tyr-92 that is exposed upon removal of the last 4 to 6 carboxylterminal residues of RNase A. Nitration of tyrosine residues with tetranitromethane (145, 146) has been applied to RNase A (146, 147). The modification reaction can be followed spectrophotometrically (146) and introduces a nitro group orrho to the phenolic hydroxyl, lowering its p K, from 10.2 to about 6.8. Reduction of the 3-nitro group to the amine with bisulfite raises the p K, of the hydroxyl to about 10; the pK, of the aromatic amine is about 5 (148). At pH 8, 3.1 tyrosine residues were reported to be nitrated, based on the extinction at 428 nm; 2.6 nitrated tyrosine residues were indicated by amino acid analysis (146).Beaven and Gratzer (147) reported that a maximum of three tyrosine residues could be nitrated; the product remained enzymatically active. Reaction of proteins with tetranitromethane can lead 139. 140. 141. 142. 143. 144. 145. 146. 147. 148.

Scheffer, A . J., and Beintema, J. J . (1974). E J E 46, 221. Burstein, Y.,and Patchornik, A. (1972). Biochemisrr.~11, 2939. Fujioka, H . , and Scheraga, H. A. (1965). Eiocliemistry 4, 2206. Taniuchi, H. (1970). JEC 245, 5459. Puett, D. (1972). Bioc/iemisrry 11, 1980. Puett, D. (1972). Biochemistry 11, 4304. Riordan, J . F., Sokolovsky, M., and Vallee, B. L. (1966). JACS 88, 4104. Sokolovsky, M . , Riordan, J. F.. and Vallee, B. L. (1966). Biochemistry 5, 3582. Beaven, G. B., and Gratzer, W. B. (1%8). EEA 168,456. Sokolovsky, M., Riordan, J. F., and Vallee, B. L. (1967). BBRC 27, 20.

12. PANCREATIC RIBONUCLEASES

34 1

to cross-linking (149, I N ) , resulting in poor correlation between spectrophotometric measurements and amino acid analyses; in that case, the sum of Tyr + 3-nitro-Tyr is less than the total tyrosine content of the native protein. The mechanism of the nitration reaction involves the phenoxide ion and proceeds via formation of free radicals that lead to cross-linking (151) and possibly other side reactions (152, 153). Thus, interpretation of the results of the reaction of proteins with tetranitromethane requires careful characterization of the reaction products. Garel and Baldwin (154)found that nitration of RNase followed first-order kinetics, indicating similar reactivities for the reacting groups. About 10% of cross-linked material was found using the procedure according to Sokolovsky er a / . (146). The monomeric fraction had 2.7-2.8 3-nitro-Tyr residues per molecule; the modified protein was similar to the native and kcat values toward cyclic 2',3'-CMP. enzyme with respect to T,, K,,,, Van der Zee r f 01. (155) have purified, by isoelectric focusing, two species from the reaction products of tetranitromethane with RNase A. One was nitrated at Tyr-115, the other at Tyr-76 and Tyr-115; both were fully active toward cyclic 2',3'-CMP. Seagle and Cowgill (156) reported on the nitration of RNase A and conversion of the nitro groups to the amine. The procedure described by Beaven and Gratzer (147) was used to modify three tyrosine residues under nondenaturing conditions. The same procedure under denaturing conditions permits modification of all six tyrosine residues. Prior acetylation (157) of the three normal tyrosine residues enabled only the three abnormal tyrosine residues to be modified (156). Seagle and Cowgill (156) report that after only one net tyrosine residue was nitrated and reduced to the amine, the product had 70% enzymatic activity toward cyclic 2',3'-CMP. The fluorescence emission maximum of this derivative was at 395 nm, higher than values observed for aminotyrosyl residues in peptides (I%),and suggests this aminotyrosyl residue is located in an unusual environment (156). The products of the nitration reaction were not analyzed for possible cross-linking; under the conditions of Beaven and Gratzer (147) as used by Seagle and Cowgill (156), this un149. 150. 151. 152. 153. 154. 155. 156. 157. 158.

Vincent, J. P., Lazdunski, M., and Delaage, M. (1970). EJE 12, 250. Hugli, T. E., and Stein, W. H . (1971). JBC 246, 7191. Bruice, T. C., Gregory, M . J., and Walters, B. L. (1968). JACS 90, 1612. Walters, S. L., and Bruice, T. C. (1971). JACS 93, 2269. Jewett, S. W., and Bruice, T. C. (1972). Biochemistry 11, 3338. Garel, J.-R., and Baldwin, R. L. (1975). J M B 94, 621. van der Zee, R.,Duisterwinkel, F. J . , and Welling, G . W. (1977). EJB 77, 125. Seagle, R. L . , and Cowgill, R . W. (1976). BBA 439, 470. Riordan, J. F., Wacker, W. E. C., and Vallee, B. L. (1965). Biochemistry 4, 1758. Seagle, R. L., and Cowgill, R. W. (1976). BBA 439, 461.

342

PETER BLACKBURN AND STANFORD MOORE

doubtedly occurred. Since van der Zee et al. (155) demonstrated Tyr-115 as a sole site of nitration in one of their reaction products, perhaps Tyr-115 is the 3-amino-Tyr residue reported by Seagle and Cowgill (156) to be in an unusual environment. Van der Zee et al. (155) suggest that this result may arise from the interaction between the neighboring side chains ( 1 ) of modified Tyr-115 and unmodified Tyr-73. These authors also observe that correct spectrophotometric quantitation of 3-nitro-Tyr can be best obtained at 381 nm between pH 4 to 10, where the absorption spectrum exhibits an isosbestic point (146). The diazonium salt of 5’-(4-aminophenylphosphoryl)uridine 2’(3’)phosphate reacts stoichiometrically with RNase A and modifies only Tyr-73 (159). The reaction is inhibited by the competitive inhibitor cytidine 2’(3’), 5‘-diphosphate. The modification does not affect the activity toward RNA but weakens 40-fold the binding constant toward cyclic 2‘ ,3’-CMP. By contrast, reaction withp-diazophenyl phosphate modifies 2 lysine, 1 histidine, and 3 tyrosine residues of RNase A (159). The peptide modified by reaction with 4-diazophenyl-pup is isolated in good yield from a tryptic digest of the reduced S-carboxymethylated RNase by affinity chromatograppy on Sepharose-RNase A. The K , values for 4-nitrophenyl-pup and 4-aminophenyl-pup determined by the inhibition of hydrolysis of cyclic 2’,3’-CMP at pH 7.0 and 22” are 43 and 70 p M , respectively (159). 8. Reactions wit17 Radicals (1. Radiolysis. In aqueous solutions RNase is inactivated by hydrogen atoms generated externally (160, 161), by steady (162) or pulse radiolysis (163, I @ ) , and by OH radicals and hydrated electrons generated by pulse radiolysis (165). The chemical evidence (160-162), and that from pulse radiolysis kinetic spectroscopy (163-165), indicate the occurrence of intramolecular radical chain reactions with cystine and methionine linked with modification of tyrosine and possibly phenylalanine residues (160166). Reaction with OH radicals leads to greater damage of surface residues and is associated with extensive cross-linking of the protein. Di-

159. Gorecki, M., and Wilchek, M. (1978). BBA 532, 81. 160. Holmes, B. E., Navon, G., and Stein, G. (1%7). Nature (London) 213, 1087. 161. Shapira, R., and Stein, G. (1968). Science 162, 1489. 162. Mee, L. K . , Adelstein, S. J., and Stein, G. (1971). Radiat. Res. 47, 349. 163. Mee, L. K., Adelstein, S. J., and Stein, G. (1972). Radiat. Res. 52, 588. 164. Lichtin, N . N . , Ogdan, J., and Stein, G. (1971). BBA 263, 14. 165. Lichtin, N. N . , Ogdan, J., and Stein, G. (1972). BBA 276, 124. 166. Stein, G. (1968). In “Energetics and Mechanisms in Radiation Biology” (G.0. Phillips, ed.), p. 407. Academic Press, London.

12. PANCREATIC RIBONUCLEASES

343

merization is essentially absent upon H atom attack. The cross-linking probably arises from the abstraction of H atoms from the saturated a-carbon atom of the polypeptide backbone, and accounts for 20% of the action of OH radicals with RNase (165). The kinetics of the transitions in the absorption spectra of RNase A and RNase S-protein upon reaction with H atoms differ both qualitatively and quantitatively at different wavelengths of the spectra. Thus, the intramolecular free radical chain reactions in the two proteins are different and reflect the different conformational flexibilities of the proteins. The reactions in RNase A differ from the sum of those observed spectrally in S-peptide and S-protein (167). b. UV Irradintion. Irradiation of anaerobic solutions of RNase A at pH 5.0 and 4" with UV light at 254 nm causes inactivation of the enzyme. Inactivation correlates with the destruction of disulfides (168-171); histidine is not affected (169). The loss of activity is associated with generation of free suIfiydry1, but always less than 2 moles per mole of disulfide disrupted (171); the remaining cysteine is lost via secondary reactions generating primarily H2S and aldehyde (160, 161, 166, 172, 173). The photodestruction of RNase A disulfides is nonrandom (172); rates differ by at least a factor of 10, and may involve interactions with neighboring aromatic groups (169, 172-174). When irradiation is at 280 nm, where 90% of the absorption is due to tyrosine residues, photoinactivation of RNase A still correlates with disulfide destruction (169). Amino acid analyses of photoinactivated RNase A, with 50% residual enzymatic activity, indicates that only the cystine content is significantly reduced (169-172). Spectrophotometric evidence suggests the possible modification of tyrosine to bityrosine (175). Since reduction of the disulfide bonds between residues 65-72 and 58- 110 does not inactivate the enzyme ( I ) , photoinactivation must arise from disruption of the disulfide between either residues 26-84, or 40-95, or both. Photoinactivation of RNase A at near 280 nm most likely occurs 167. Lichtin, N. N . , Ogdan, J . , and Stein, G . (1973). Radiat. Res. 55, 69. 168. Schultz, R. M., Immartino, A. J . , and Aktipis, S . (1975). BBA 386, 120. 169. Rathinasamy, T. K . , and Augenstein, L. G. (1968). Biophys. J . 8, 1275. 170. Grist, K . L., Taylor, T., and Augenstein, L. (1965). Radiat. Res. 26, 198. 171. Augenstein, L., and Riley, P. (1964). Phororhem. Photobiol. 3, 353. 172. Risi, S., Dose, K., Rathinasamy, T. K., and Augenstein, L. (1967). Photochem. Pirotobiof. 6 , 423. 173. Shafferman, A., and Stein, G. (1974). Photochern. Photobiol. 20, 399. 174. Arian, S . , Benjamini, M . , Feitelson, J., and Stein, G. (1970). Photochem. Phorobiol. 12, 481. 175. Aktipis, S . , and Iammartino, A . J. (1972). BBA 278, 239.

344

PETER BLACKBURN AND STANFORD MOORE

via activation of buried tyrosine residues, with subsequent sensitization of disulfide bonds (176, 177). Tyr-92 and Tyr-97 are both adjacent to the essential disulfide between residues 40-95. Tyr-97 is also adjacent to the essential disulfide between residues 26-84; Tyr-25 is in the vicinity of this disulfide. The exposed Tyr-73 and Tyr-115 are adjacent to the nonessential disulfide between residues 58- 110. The nonessential disulfide between residues 65-72 is not near any tyrosine residues; Tyr-76 is not close to any disulfide. The photoinactivation of RNase A has been shown by CD measurements to be associated with an altered and less stable conformation of the protein (168, 178). Photooxidation of NE-dinitrophenyl-Lys-41-RNase A by UV irradiation in the presence of molecular oxygen leads to specific modification of one methionine, one histidine, and one tyrosine residue (179) at positions 30, 12, and 97, respectively (180). After specific reduction of the disulfide bond between residues 65 and 72 according to Sperling ef al. (130), and S-dinitrophenylation, photooxidation of the protein upon irradiation led to modification of Tyr-73 and Tyr-115 (180). These reactions permit the topological and relative spacial arrangements of residues to be probed. In general, oxidation is normally restricted to Cys, Trp, Tyr, Met, and His residues (179, /81). The covalent attachment of 4-thiouridylic acid to RNase by irradiation with UV light at 334 and 365 nm has been reported (182). Irradiation of RNase A complexes with cytidine 2’(3’),5‘-diphosphate (pCp) or uridine 2‘(3’),5’-diphosphatewith ultraviolet light >300 nm resulted in covalent attachment of the pyrimidine nucleotides to the enzyme (183). Tryptic hydrolysis and peptide mapping showed attachment in the peptide segment from Asn-67 through Arg-85 of RNase A . Matheson et al. (184) studied the more generalized labeling of RNase A with the aryl nitrene, N-(4-nitreno-2-nitrophenyl)-2-aminoethylsulfonate, generated by Aash photolysis from N-(4-azido-2-nitrophenyl)-2aminoethyl sulfonate (NAP-taurine) (185), and have used this reaction to 176. 177. 178. 179. 180.

Volkert, W. A., and Grossweiner, L. I . (1973). Plzotochem. fhotohiol. 17, 81. Setlow, R., and Doyle, B. (1957). BBA 24, 27. Aktipis, S., and lammartino, A. J. (1971). BBRC 44, 918. Scoffone, E., Galiazzo, G . , and Jori, G. (1970). BBRC 38, 16. Jori, G . , Galiazzo, G . , Marchiori, F., and Scoffone, E. (1970).l a f .J . Peptide Protein Res. 2, 247. 181. Jori, G . , Gennari, G ., Galiazzo, G . , and Scoffone, E. (1970). FEBS Letr. 6, 267. 182. Sawada, F. (1975). BBRC 64, 311. 183. Sperling, J., and Havron, A. (1976). Biochemistry 15, 1489. 184. Matheson, R. R., Jr., Van Wart, H. E., Burgess, A. W., Weinstein, L. I., and Scheraga, H. A. (1977). Biochemistry 16, 3%. 185. Staros, J. V., and Richards, F. M . (1974). Biochemistry 13, 2720.

12. PANCREATIC RIBONUCLEASES

345

study steps in the thermal unfolding of RNase A (186). Nitrenes generated in the presence of the protein are capable of inserting into carbon-hydrogen bonds to form secondary amines that are stable to acid hydrolysis (187). Modification of exposed amino acids is not loo%, and varies in extent with the type of residue for reasons that are not altogether clear (188). With RNase, a degree of saturation of labeling occurred at a reagent to protein ratio of 212: 1 (184). With mixtures of amino acids free in solution, basic amino acids are labeled more than acidic amino acids, and nonpolar amino acids with larger side chains are labeled more than those with smaller side chains. However, the selectivity of the nitrene for the amino acids in a polypeptide chain like that of RNase A is influenced by the environment around these residues in the protein.

B. ROLESOF RESIDUES NEARTHE NH2A N D COOH TERMINI 1. S-PeptideS-Protein Interaction

The S-peptide-S-protein system discovered by Richards ( 1 , 189), which results from the controlled cleavage by subtilisin primarily between Ala-20 and Ser-21, has formed the basis for extensive studies of the roles of residues in the two sections of the enzyme. The first 25 residues of the sequence contain some of the most variable positions of the mammalian pancreatic RNase molecules (see Section V), yet hybrids prepared from S-peptides and S-proteins of different species have remarkably similar properties (190-192). The S-peptide segment has been subject to continued synthetic studies ( 1 93) to identify residues that have key roles in the binding to S-protein and in the activity of the complex. With the knowledge that residues 15 to 20 are dispensable ( I ) , substitutions in residues 1 through 15 have given a number of crystalline analogues that give X-ray diffraction patterns very similar to that of RNase S (194, 195). Substitution of Asn for Asp at 186. Matheson, R. R., Jr., and Scheraga, H. A. (1979). Biochemistry 18, 2437. 187. Knowles, J. R. (1972). Accounrs Chem. Res. 5, 155. 188. Bayley, H . , and Knowles, J. R. (1978). Biochemistry 17, 2414. 189. Richards, F. M. (1955). C.R. Trav. Lab. Carisberg 29, 322. 190. Welling, G. W., Lenstra, J. A., and Beintema, J . J. (1976). FEES Lerr. 63, 89. 191. Voskuyl-Holtkamp, I., Schattenkerk, C., and Havinga, E. (1976). Int. J . Pepride Protein Res. 8, 455. 192. Voskuyl-Holtkamp, I., and Schattenkerk, C. (1977). I n / . J . Peptide Prorein Res. 10, 60;ibid. 10, 153; ibid. 11, 218. 193. Chaiken, I. M. (1978). In “Semisynthetic Peptides and Proteins” (R.E. Offord and C. DiBello, eds.), p. 349. Academic Press, New York. 194. Chaiken, I. M., Taylor, H. C., and Amrnon, H. L. (1977). JBC 252, 5599. 195. Pandin, M., Padlan, E. A . , Di Bello, C., and Chaiken, I. M. (1976).PNAS 73, 1844.

346

PETER BLACKBURN AND STANFORD MOORE

position 14 reduces the affinity of S-peptide for S-protein about 20-fold but has no effect on the enzymatic activity in the presence of an excess of this analog (1%). In S-protein, Tyr-25 titrates spectrophotometrically with a normal pK, of about 10.2 (197); addition of S-peptide to form RNase S' restores the buried characteristics of Tyr-25 (197-200). Addition of lC,7€, 10S-triguanidino-(Orn-10, Asn- lrl)-S-peptide to S-protein restores 75% of the spectral characteristics of RNase S' (1%). This result indicates that Asn-14 can hydrogen bond to the phenolic hydroxyl of Tyr-25, as does Asp-14 in RNase A and RNase S. Finn and Hofmann (201) and Hearn et al. (202) proposed that a charge interaction exists between Asp-14 and Arg-33. Filippi et al. (1%) suggested that the weaker interaction arising from substitution of Asn for Asp-14 may result in large part from loss of this charge interaction. The binding of 1€,7€,10S-triguanidino-(Orn-10, Asn-14)-S-peptide to S-protein, as followed by the change in CD at 222 nm, exhibits a titration curve with a midpoint near pH 6.0 (1%). The binding of S-peptide or lC,7'-diguanidino-S-peptide does not exhibit this pH dependence. The titration behavior of the binding of the Asn-14 S-peptide analog may be a reflection of an interaction involving His-48 expressed in the absence of a strong electrostatic interaction with Arg-33. NMR titration studies on RNase A have suggested a close association between Asp-14 and His-48 (77-79), and between Asp-14 and Tyr-25 (74, 80). However, the 13C NMR studies of Niu et al. (203, 203a), with an S-peptide analog of residues 1-15 synthesized with 13C-enriched Asp14, Met- 13, and His-12, indicate that the proposed interaction between Asp-14 and His-48 is unlikely, and favor a hydrogen bond between Asp-14 and Tyr-25. (See Section IV,B.) Met-13 contributes to the binding of S-peptide to S-protein (I). Alkyla1%. Filippi, B., Moroder, L., Borin, G . , Samartsev, M., and Marchiori, F. (1975). EJB 52, 65. 197. Shenvood, L. M., and Potts, J. T., Jr. (1965). JBC 240, 3806. 198. Woodfin, B. M., and Massey, V. (1968). JBC 243, 889. 199. Fung, D. S., and Doscher, M. S. (1971). Biochemistry 10, 4099. 200. Rocchi, R., Borin, G . , Marchiori, F., Moroder, L., Peggion, E., Scoffone, E., Crescenzi, V., and Quadrifoglio, F. (1972). Biochemistry 11, 50. 201. Finn, F. M., and Hofmann, K. (1973). Accounts Chem. Res. 6 , 169. 202. Hearn, R . P., Richards, F. M. Sturtevant, J. M., and Watt, G . D. (1971).Biochemistry 10, 806. 203. Niu, C.-H., Matsuura, S., Shindo, H., and Cohen, J. S. (1979). JBC 254, 3788. 203a. Cohen, J. S . , Niu, C.-H., Matsuura, S., and Shindo, H. (1980). In "Frontiers in Protein Chemistry" (T.-Y. Liu, G . Mamiya, and K . T. Yasunobu, eds.), p. 3. ElseviedNorthHolland, New York.

12. PANCREATIC RIBONUCLEASES

347

tion or oxidation of Met-13 (204) or substitution by leucine (205, 206) decreases the affinity for S-protein, but has little effect on the activity of the complex once formed. In the three-dimensional structure of the RNase molecule, the methionine side chain fits into a hydrophobic pocket formed by Val-47, Leu-51, and Val-54 ( 1 ) . This interaction was observed in the I3C NMR spectrum of S-peptide-(l-15), synthesized with [13C]Met-13,as an upfield shift resulting from greater shielding by the hydrophobic pocket (203). Modifications of Met-13 that increase the hydrophilicity (e.g., formation of the sulfoxide or the sulfone) or introduce a charge by alkylation (116) alter the steric properties of this residue. Rotation about the a-P carbon-carbon bond of Met-13 permits a charged sulfur atom in the complex to have access to solvent without a change in the conformation of the S-peptide; enzymatic activity is retained but there is a decrease in binding stability through loss of the hydrophobic contacts. Substitution of His-12 by Ser in S-peptide-(1-14) (106) and by Om-12 in (Om-lO)-S-peptide-(l-20) (207) significantly decreases the affinity for S-protein, suggesting that the side chain of His-12 contributes to the binding between S-peptide and S-protein. Replacement of His-12 with P-(pyrazolyl-3)-~-alanine( 1 05) or 4-fluoro-~-histidine(208) has no effect on the binding of S-peptide to S-protein, but the complexes are inactive. The ring protons of these residues have pK, values near 2.6, and they cannot participate in acid-base catalysis at neutral pH. Competitive ligand (2’-CMP) elution affinity chromatography on Sepharose-Caminophenyl5’-phosphoryluridine 2’(3‘)-phosphate (209) has demonstrated that substitution of 4-F-His at position 12 does not affect substrate binding (210). Carboxymethylation of N-3 of the imidazole of His-12 in a peptide that contains residues 1 through 14 actually increases the affinity of this S-peptide analogue for S-protein ( 105). Substitution of His-12 by L-homohistidine lengthens the side chain of residue 12 by one methylene group, but retains the imidazole moiety with a pK, similar to that of histidine. This substitution in an S-peptide 204. Richards, F. M., and Vithayathil, P. J. (1980). Brookliuven Syrnp. B i d . 13, 115. 205. Hoes, C., van Batenburg, 0. D., Kerling, K. E. T.,and Havinga, E. (1977). BBRC 77, 1074. 206. van Batenburg, 0. D., Raap, J., Kerling, K. E. T., and Havinga, E. (1976). Rec. Truv. Chim. Pays-Bus 95, 278. 207. Borin, G., Toniolo, C., Moroder, L., Marchiori, F., Rocchi, R., and Scoffone, E. (1972). I n r . J. Peptide Protein Res. 4, 37. 208. Dunn, B. M.,Di Bello, C., Kirk, K. L., Cohen. L. A., and Chaiken, I. M. (1974). JBC 249, 6295. 209. Chaiken, I. M.,and Taylor, H. C. (1976). JBC 251, 2044. 210. Taylor, H . C., and Chaiken, I. M. (1977). JBC 252, 6991.

348

PETER BLACKBURN AND STANFORD MOORE TABLE 111 S-PROTEIN-ACTIVATING ABILITY (Amax)A N D B I N D I N G CAPACITY OF SOME RIBONUCLEASE S-PEPTIDE-(1-14) ANALOGS I N WHICH T H E ACTIVE-SITE IS REPLACED B Y OTHERRESIDUES" HISTIDINE-12 Binding capacity'

Residue 12

(%)

Substrate present

100

1000

80

10

Substrate absent

Ref.

1

H

I

0.01

(211)

Immeasurably low

(212)

0.01

(213)

nd'

(217)

H

Nhi

4IJ

80

0.01

20

0.1

<1

lood

34

100

H

(3-Pyd)Ala

- C H 2 G

-N

(4-Pyd)Ala

A H 2P

w

" From van Batenburg er

N

r r / . ( 2 1 1 - 2 / 3 ) and Hoes et rrl. ( 2 f 7 ) . with yeast RNA as a substrate at pH 5.0. A,,, is expressed relative to S-peptide-(1-14) (100%).

' Measured

12. PANCREATIC RIBONUCLEASES

349

analogue of residues 1 through 14 decreases the affinity for S-protein about 100-fold, but the complex retains 80% activity toward RNA (211); the kinetics of hydrolysis of cyclic 2’,3’-CMP exhibits a fourfold lower k,,, and a 9-fold higher K , than that exhibited by RNase S (206). Substitution of 4-imidazolylglycine for His-12 reduces the side chain of residue 12 by one methylene group; the p K , for dissociation of the imidazole protons is about 4.6 (212). The affinity for S-protein of this S-peptide analog, consisting of residues I through 14, is 10”-fold lower than that of native S-peptide residues 1 through 20; the complex has a maximum activity of 80% of that of RNase S toward RNA. Thus slight modifications in the side chain of His-12, while retaining the imidazole ring, can severely impair the ability of S-peptide to bind to S-protein (211, 212) and clearly demonstrate an important role for this residue in binding to S-protein. On the other hand, these modifications only slightly affect the activity of the complex and indicate that a degree of conformational flexibility is permissible at the active site of the complex. The effects of substitution of N-1-methylhistidine or N-3-methylhistidine at position 12 on the binding of S-peptide analogs of residues 1 through 14 (213) (Table 111) are consistent with the importance of N-3 of the imidazole for the catalytic activity, and that of N-1 of the imidazole in binding to S-protein; N-1 is thought to hydrogen bond t o the peptide carbonyl of Thr-45 ( I , 16, 214). Substitution of His-I2 of this S-peptide analogue by P-(3-pyridyl)-~-alanine introduces a side chain with a single titrating nitrogen atom with p K , for dissociation of the proton of about 5.5 (21.5). This derivative demonstrates a high affinity for S-protein, 211. van Batenburg, 0. D., Raap, J., Kerling, K. E . T., and Havinga, E. (1975). TctLett. 51, 4591. 212. van Batenburg, 0. D., Kerling, K. E. T., and Havinga, E. (1976). FEES Lett. 68, 228. 213. van Batenburg, 0. D., Voskuyl-Holtkamp, l . , Schattenkerk, C., Hoes, K., Kerling, K. E. T., and Havinga, E. (1977). BJ 163, 385. 214. Patel, D. J . , Canuel, L. L., Woodward, C., and Bovey, F. A. (1975). Biopolymers 14, 959. 215. Voskuyl-Holtkamp, I . , and Schattenkerk, C. (1979). h i . J . Pepride Protein R e f . 13, 185.

rrihedroti

‘ Binding capacity in the presence of substrate was determined directly from activity assays [Berger and Levit, Ref. (22211; values are expressed relative to Speptide-(l-14) + substrate = 1000, which corresponds to a binding constant of 5 x 10” M - ’ . Binding capacity in the absence of substrate was determined by UV-difference spectroscopy: values are expressed relative to S-peptide-(l-14) + substrate = 1000. Determined by competitive inhibition experiments by using ribonuclease S’ as a standard. ‘’ nd, Not determined.

‘

350

PETER BLACKBURN A N D STANFORD MOORE

but the complex is enzymatically inactive (213). Substitution of p(4-pyridyl)-~-alanineat position 12 (216) reduces the affinity for S-protein by about 10-fold; this complex exhibits 35% the activity of RNase S toward RNA but less than 4% toward cyclic 2',3'-CMP (217). The suggestion that both N-1 and N-3 imidazole nitrogens of His-12 are essential for activity (213) seems unlikely in view of the considerable activity toward RNA displayed by the complex of S-protein with p-(4-pyridyl)-~-Ala123-peptide-(1- 14) (217). Although Gln-11 can be substituted by Glu without major effect on activity, or significant reduction of the affinity for S-protein (218, 219), the adjacent Arg- 10 is important for binding to S-protein; substitution by ornithine significantly reduces the affinity of the S-peptide for the Sprotein (219, 220). Hofmann et cd. (105, 106) were first to suggest that an interaction between Arg- 10 and Glu-2 contributes to the conformation of S-peptide required for binding to S-protein. Evidence for this interaction has come from comparisons of the 250-MH, proton NMR spectra of N-3carboxymethyl-His-12-S-peptide with those of (Orn-lO,N-3-carboxymethyl-His- 12)-S-peptide upon addition of S-protein (221). Appropriate chemical shifts of the C6-H resonances of Arg-10 are observed upon addition of S-protein; the signals from the ornithine side chain remain unchanged. Marchiori er (11. (104) showed that substitution of Ala-6 by proline (thereby shortening the a-helical portion of the S-peptide) prevents the interaction between the side chains of Glu-2 and Arg-10. This substitution decreased the catalytic activity of the complex but was without serious effect on the K, for both cyclic 2',3'-substrates and RNA (104).

The binding properties and activities of S-peptide and des-1 through des-8 S-peptide complexes with S-protein have been studied by Berger and Levit (222, 223) and are shown in Table IV. Loss of residues Lys-1 through Thr-3 causes a 450-fold decrease in affinity for S-protein. The 216. Hoes, C., Raap, J., Bloemhoff, W., and Kerling, K. E. T. (1980). R r c . Tmv. Chim. Pays-Bris 99, 99. 217. Hoes, C., Hoogerhout, P., Bloemhoff, W., and Kerling, K. E . T., (1979). Rrc. Trav. Chim. Poys-Bns 98, 137. 218. Finn, F. M . , and Hofmann, K. (1965). JACS 87, 645. 219. Scoffone, E., Rocchi, R., Marchiori, F., Marzotto, A . , Scatturin, A . , Tamburro, A . , and Vidali, G. (1967). J . Clirm. Soc. ( C ) ,p. 606. 220. Moroder, L . , Marchiori, F., Rocchi, R., Fontana, A . , and Scoffone, E. (1969).JACS 91, 3921. 221. Finn, F. M . , Dadok, J . , and Bothner-By, A. A. (1972). Biochemistry 11, 455. 222. Berger, A . , and Levit, S. (1973). In "Peptides 1971" (H. Nesvadba, ed.), Roc. 11th Eur. Peptide Symp. Vienna, 1971, p. 373. North-Holland, Amsterdam. 223. Levit, S ., and Berger, A . (1976). JBC 251, 1333.

35 1

12. PANCREATIC RIBONUCLEASES TABLE IV B I N D I N PROPERTIES G A N D ACTIVITIES OF $PROTEIN : S-PEPTIDE SYSTEMS AT pH 5.0, 27""

(9-20) (8-20) (7-20) (6-20) (4-20) (3-20) (2-20) (1-20)

3.1 2.2 3.1 5.0 3.2 7.0

x lo-" x lo-"

x x 10-8 x lo-" x lo-'

3.2 x 4.6 x 3.2 x 2.0 x 3.1 x 1.4 x

lo4 104 10'

107 lo7 10"

6.3 6.5 7.7 10.2 10.5 11.4

Not measurable Not measurable 0.21 0.29 0.48 0.56 0.86 1.oo

" From Berger and Levit (222, 223 ). I,

Measures catalytic efficiency.

three alanine residues at positions 4 3 , and 6 contribute little to the binding with S-protein (220, 223) and may be substituted by hydrophilic serine residues with little effect on the interaction or on the activity of the complex ( 2 2 4 ) . The hydrophilic side chain of Glu-9 is exposed to solvent and forms part of the a-helical segment of S-peptide. Substitution by Leu-9 in Speptide-(l-15) causes only a 3-fold reduction in the K b for S-protein; substitution by Gly-9 in S-peptide-(I- 15) causes a 22-fold reduction in the K b (225). Thus, the negative charge and hydrophilicity at position 9 are not essential. The different abilities of leucine and glycine to substitute for Glu-9 reflect their different propensities to participate in a-helical structures (225). Acylation of the NH2 groups of S-peptide ( 1 , 222, 223) does not have a determining effect on the binding efficiency. Hofmann et al. (106) demonstrated that substitution of Nle for Lys-7 of S-peptide has only a small effect on the activity of the complex. Thus, Lys-7 is not essential for catalytic activity. The presence of substrate influences the interaction between S-protein and S-peptide (201). The complexes in general exhibit greater activity toward RNA than toward cyclic 2',3'-substrates (104, 218). The ultraviolet spectral changes that accompany binding of S-peptides to 224. Borin, G . , Marchiori, F., Moroder, L., Rocchi, R., and Scoffone, E . (1971). BBA 271, 77. 225. Dunn, B. M., and Chaiken, I . M. (1975). J M B 95, 497.

352

PETER BLACKBURN AND STANFORD MOORE

S-protein (226) are proportional to the concentration of the complex ( 1 98) and result from the exclusion of Tyr-25 from solvent. In general, the application of direct spectral techniques to follow binding between S-protein and S-peptide analogs produces qualitatively similar results to those obtained by activity determinations and competition assays between S-peptide and analogs that produce inactive complexes with S-protein (227). However, quantitatively, the presence of substrate has a significant effect on the binding constant for the association of S-peptide with S-protein (Table V ; including Refs. 228-230). Gawronski and Wold (231) studied the interaction between S-peptide coupled to CNBr-activated agarose and S-protein, and reported a K d at pH 7.5 and 23" of 2 x 10-6M.Covalent attachment of the S-peptide to the insoluble matrix had only a small (fourfold) negative influence on the association. Also, below 25" the dissociation constants for RNase S' and complexes with S-protein acetylated with 13H]aceticanhydride at 1 and 9 moles per mole of S-protein were essentially identical (232). With this system, saturating concentrations of a mixture of 2'(3')-CMP had no influence on K d . Studies of the thermodynamics of the interaction suggested that the association was entropically driven. Calorimetric studies on the association of S-peptide with S-protein, performed by Hearn et a/. (202) indicated that the process is enthalpically driven. Estimates of Kd were made between 30" and 45" from assays toward cyclic 2',3'-CMP. The van't Hoff plots were nonlinear in this range, as also reported by Gawronski and Wold (232);the higher temperature, in this case, lies close to the transition temperature of S-protein. The calorimetric measurements of AH were performed in the absence of substrate. The data in Table V and the studies of Schreier and Baldwin (233, 234), show the degree to which binding of substrate influences the association of S-peptide with S-protein. Schreier and Baldwin (233, 234), taking advantage of the different amide 3Hexchange rates of S-peptide when free and bound to S-protein, studied the interaction as a function of temperature, pH, ionic strength, and the presence of 2'-CMP. They distinguished two steps in the dissociation, a partial unfolding step and a separation 226. Richards, F. M., and Logue, A. D. (1962).JEC 237, 3693. 227. Finn, F. M. (1972). Biocltemistry 11, 1474. 228. van Batenburg, 0. D. (1977). Ph.D. Thesis, University of Leiden, The Netherlands. 229. Kenkare, U. W., and Richards, F. M. (1966). JEC 241, 3197. 230. Marzotto, A., Marchiori, F., Moroder, L., Boni, R., and Galzigna, L. (1967). EBA 147, 26. 231. Gawronski, T. H . , and Wold, E (1972). Eioehernistry 11, 442. 232. Gawronski, T. H., and Wold, F. (1972). Biochemistry 11, 449. 233. Schreier, A . A., and Baldwin, R. L. (1976). J M E 105, 409. 234. Schreier, A. A., and Baldwin, R . L. (1977). Biochemistry 16, 4203.

12. PANCREATIC RIBONUCLEASES

353

TABLE V

LITERATURE K b VALUES FOR T H E ASSOCIATION BETWEEN S-PEPTIDE-(I-20)A N D S-PROTEIN'

Substrate None

Ref.

(227) (208) (226) (198) (200) (223) (198) (223) (229) (202) (225)

1 x 10'

0.5 x 10' 1.4 x lo4 1 x lo5 0.3 x 10" 4 x 107 2.2 x lo6 1 x 10"

S-Protein conc. ( M ) 5.5 x 10-j 5 x lo-: 8.7 x 10-j 6.5 x lo-' 7 x lo-'

6.5 x 9.3 x 10-6 5 x 10-7 2.2 x

pH 5.0

7.1 4.5 -

6.8 5.0 5.4

6.5 7.0 7.0

1.2 x 107 2.8 X lo7

4.7 x 10-7

5 x 10-7

7.1

5 x 107

1.5 x 10-7

7.1

1.4 x 10"

5 x lo-'

(104)

1.9 x lo8 3.3 x loH

(191)

4.2 x 10'

5.2 x lo-' 8.5 x lo-' 2.5 x lo-"

5.0 5.0 6.0 5.0

(1%)

Yeast RNA

Kb(M-')*

(223) (230)

" From van Batenburg (228). 6Measured in the absence of substrate as determined by UVdifference spectroscopy, and in the presence of substrate as determined by activity recovery. ' Stoichiornetric binding; no calculation possible.

step. Unfolding was enhanced at lower pH and ionic strength, suggesting that it is induced by electrostatic repulsion, possibly between the positive charges of residues Lys-7, Arg-10, and His-12. The separation step was independent of ionic strength, indicating that nonionic interactions predominate. Binding of 2'-CMP had a large effect on the separation step, and the Kd became too small to measure. The equilibrium constants and thermodynamic parameters for the dissociation and partial unfolding of S-peptide bound to RNase S are shown in Table VI. Schreier and Baldwin indicate that residues 12-14 may serve as an "anchor" in the initial combination of S-peptide with S-protein; binding of 2'-CMP with His- 12 would then assist this association. Rosa and Richards (2342) have obtained increased structural resolution by combining HPLC of proteolytic 234a. Rosa, J. J., and Richards, F. M. (1979).JMB 133, 399; (1981).ihitl. 145, 835.

354

PETER BLACKBURN AND STANFORD MOORE TABLE VI

A. TEMPERATURE-DEPENDENCE OF T H E E Q U ~ L I B R CONSTANTS IUM FOR DISSOCIATION

(Kd) A N D PARTIAL UNFOLDING (K,) OF S-PEPTIDE BOUNDTO RNASEs"

4.25 4.25 4.25 6.9 6.9 6.9

B.

4.0 7.9 12.5 0 3.6 8.4

VAN'T

1.6 x 3.4 x 8.0 x 8.8 x 2.3 X 3.7 x

1.0 x 1.6 x 2.3 x 5.9 x 7.0 x 1.3 x

10-8 lo-' lo-'' lo-'"

HOFFENTHALPY A N D ENTROPY OF DISSOCIATION AND UNFOLDING OF S-PEPTIDE BOUNDTO RNASEs'

OF

10-3 10-3 10-3 lo+ lo-'

PARTIAL

PH

Temp. range ("C)

Reaction

AH (kcaYmol)

AS (euimol)

4.25 4.25 6.9 6.9

4- 12.5 4-12.5 0-8.4 0-8.4

Dissociation (Kd) Partial unfolding (K,) Dissociation ( K d ) Partial unfolding (K,)

29 14 28 13

71 39 57 24

' Reprinted with permission from Schreier and Baldwin (234), Biorhemistry 19, 42034209. Copyright (1977) American Society.

digests performed at pH 2.8 to identify residues undergoing tritium exchange in such experiments with S-peptide and S-protein. Niuet al. (235) used a synthetic S-peptide-(l- 15) labeled at His-12 with to obtain by NMR measurements a K d of 0.2 x lO-'M, which was greater by a factor of 5 than the Kd determined for the S-peptide-( 1-20) by competition experiments. On thermodynamic grounds they propose a hydrogen bond between Ser-16 and His-48 ( 1 ) to account for the difference in binding enthalpies between S-peptides-(1-20) and (1- 15). Schreier and Baldwin (234) suggest that the low K b values obtained by spectrophotometric techniques (Table 111), may reflect the presence of some molecules that are not completely native and dissociate more readily than RNase S. S-protein tends to aggregate at low pH values (236). At pH 4.5, Gawronski and Wold (231) reported anomalous binding data for the titration of agarose-S-peptide with S-protein. By attaching S-protein to the agarose matrix to prevent low pH aggregation of S-protein and titrating with 235. Niu, C.-H., Shindo, H., Matsuura, S . , and Cohen, J. S. (1980). JBC 255, 2036. 236. Allende, J . E . , and Richards, F. M. (1962). Biochemistry 1, 295.

12. PANCREATIC RIBONUCLEASES

355

S-peptide, normal titration behavior was observed. The dissociation constant of the aggregated S-protein dimer was reported to be near M at pH 7.5, but less than M at pH 4.5. They conclude that the pHdependent association of S-protein may affect the values obtained for K d of RNase S' derived by direct spectrophotometric titrations, especially those at lower pH values since relatively high concentrations of S-protein are required by this procedure (see Table V). Such an influence would tend to increase the values obtained for Kd.Indeed, Dunnet af. (208) were unable to derive a binding constant for the association of S-peptide with S-protein by spectrophotometric titration at pH 7, where the binding was apparently stoichiometric. Hearn et al. (202), however, found no effect of S-protein concentration on AH for the association of S-peptide with S-protein, and concluded that either the aggregation of S-protein does not involve significant enthalpy changes or that aggregation is insignificant in the concentration range 2 x 1O-j to 2 x lop3M. Gawronski and Wold (232) indicated that AH of dissociation of S-protein dimer is close to zero, but that A S of dissociation was - 18.5 ca1 deg-' mole-'. NMR studies have demonstrated that the active-site histidine residues of RNase A, S, and S' occupy the same chemical environment (237-239), and that the dominant conformation of free S-peptide in aqueous solution is that of a random coil (221, 240). Upon binding to S-protein, the S-peptide undergoes a coil-to-helix transition (221). By incorporating 13Clabeled glycine at position 6 and either I9F as p-fluorophenylalanine or 13C-labeledphenylalanine at position 8 of S-peptides of residues 1- 15, the coil-to-helix transition undergone by S-peptide upon binding to S-protein has been observed by NMR spectroscopy (241. 242). This transition in unfolded RNase A, with its native disulfide bonds intact, is thought to be an early event during refolding of the molecule (243). The interactions between S-peptide and S-protein help to stabilize intermediates during refolding of the molecule (244-246). These findings are in contrast to those 237. Meadows, D. H . , Jardetzky, O . , Epand, R. M., Riitejans, H. H., and Scheraga, H. A. (1968). P N A S 60,766. 238. Cohen, J. S., Griffin, J. H., and Schechter, A. N. (1973). JBC 248, 4305. 239. Shindo, H., and Cohen, J. S. (1976). JBC 251, 2648. 240. Silverman, D. N . , Kotelchuck, D., Taylor, G. T., and Scheraga, H . A. (1972). ABB 150,757. 241. Chaiken, I. M. (1974). JBC 249, 1247. 242. Chaiken, I. M . , Freedman, M. H . , Lyerla, J. R., Jr., and Cohen, J . S. (1973). JBC 248, 884. 243. Blum, A. D., Smallcombe, S. H . , and Baldwin, R. L. (1978). J M B 118, 305. 244. Schmid, F. X., and Baldwin, R. L. (1979). J M B 135, 199. 245. Labhardt, A. M., and Baldwin, R. L . (1979). J M B 135, 231. 246. Labhardt, A. M., and Baldwin, R. L. (1979). J M B 135, 245.

356

PETER BLACKBURN AND STANFORD MOORE

reported by Klee (247, 248) who estimated 10- 15% helical content of the peptide based on CD measurements. Scoffone et al. (249), on the other hand, concluded on the basis of CD and ORD measurements with S-peptide analogs that the dominant conformations were disordered. The contributions of S-peptide residues to the refolding of RNase A are discussed in Section IV,D. A new approach to the study of the S-peptide-S-protein interactions has been reported by Hoogerhout et a / . (250).Their approach is to prepare synthetic extensions at the NH2terminus of acetimidyl-blocked S-protein. This approach should reveal any influence that the covalent linkage between residues Ala-20 and Ser-21 exerts in native RNase A. Conformational restrictions associated with the covalent attachment of the S-peptide fragment to the S-protein portion have already been indicated (1, 119). Modifications to Met-13 such as, photooxidation (117, 118) and alkylation (1 16, 118),which inactivate RNase A, can be accommodated by RNase S’; even though the interactions of the modified S-peptides with S-protein are weaker, the complexes are active. Homandberg and Laskowski ( 2 5 0 ~have ) obtained enzymatic synthesis (yield 50%) of RNase A$ (a mixture of RNase A, des-Ser-21 RNase A, and possibly Ser-21A RNase A) from RNase S by the use of subtilisin in 90% (v/v) glycerol.

-

2. Modijication Near the COOH Terminus Anfinsen (251) showed that removal of the tetra peptide at the COOH terminus of RNase A by limited peptic hydrolysis resulted in a derivative with little or no activity. Lin (252) was able to demonstrate 0.5% of the activity of RNase A toward 2’,3’-cyclic CMP. The susceptibility of His-12 and His-119 to alkylation by iodoacetate is still present but reduced in rate; the alkylation favors His-12 rather than His-1 19, which is preferentially modified in the native enzyme. The transition temperature drops from 61” for RNase A to 44” for RNase-(1-120). Taniuchi (142) and Puett (143) found that the derivative maintains much of the original conformation, as judged by CD spectra. After the -S-Sbonds are split by 247. Klee, W. A. (1968). Biochemistry 7, 2731. 248. Brown, J. E., and Klee, W. A. (1971). Biochemistry 10, 470. 249. Scoffone, E., Marchiori, F., Moroder, L . , Rocchi, R., and Borin, G . (1973). In “Medicinal Chemistry 111” (P.Pratesi, ed.), p. 83. Buttenvorths, London. 250. Hoogerhout, P., Bloehmhoff, W., and Kerling, K. E. T. (1979). Rec. Trav. Chim. Pays-Bas 98, 515. 250a. Hornandberg, G . A . , and Laskowski, M . , Jr. (1979). Biochemistry 18, 586. 251. Anfinsen, C. B. (1956). JBC 221, 405. 252. Lin, M. C. (1970). JBC 245, 6726.

12. PANCREATIC RIBONUCLEASES

357

reduction (142) the reoxidized protein is largely disordered. In terms of the process of biosynthesis, the chain is thus not programmed for proper folding until the synthesis has proceeded beyond residue 120. When Phe- 120 is removed by controlled hydrolysis with carboxypeptidase A at pH 5 (2521, all evidences of catalytic activity and native structure are lost. The transition temperature of the des-( 120- 124) derivative is lowered to 34". The loosened structure is highly susceptible to proteolysis by trypsin at 25". CD spectral studies by Puett (144) show the decreased conformational stability of this derivative by unfolding experiments in guanidinium chloride solutions. The phenylalanine residue, which in the three-dimensional structure fits into a hydrophobic pocket, thus has an important role in maintenance of the native conformation of the molecule. The further removal of His-1 19 can be accomplished by carboxypeptidase A action at pH 7.6 to give RNase-( 1- 118). In a collaborative study initiated by Lin and Gutte (25.3) a synthetic peptide corresponding to the 14 amino acid residues at the COOH terminus, synthesized by the solid phase method, was added to RNase-( 1- 118);the peptide's presence led to regeneration of 90% of the activity toward 2',3'-cyclic CMP and 70% toward yeast RNA at a peptide-to-protein ratio of 3. If both the first 20 and the last 6 residues were removed from RNase, a three-component system consisting of S-peptide (3.7 equivalents), a 21-118 residue core, and residues 111-124 (3.4 equivalents) gave 30% activity toward the synthetic substrate. In this case both of the two histidine residues that are near the active center were supplied by adsorbed peptides. These studies were extended (254) to include synthetic COOH-terminal peptides of varying lengths. There was negligible reactivation until the chain length reached 9 residues, which gave 60% activity. The Kd values for the added peptides and RNase-(1-118) were 2.5 X M at 9 residues and 2 x lo-' M at 14 residues. The role of Phe-120 was examined by substituting other residues at this position in the synthetic peptides (25s). Leu, Ile, or Trp was inserted at position 120. The maximum regenerable activities, calculated according to Berger and Levit (222), were 98, 13, 12, and 0.5%, respectively, for the peptides with Phe, Leu, Ile, or Trp at position 120 (256). The order of the Kd values was 2 X 2.5 x 4x and 3.5 x 10-jM. The K , values for the complexes with 2',3'cyclic CMP (255) were all near the value for the native enzyme (about M ) . The lowered enzymatic activities appear to result from misalign253. Lin, M. C., Gutte. B., Moore, S., and Merrifield, R. B. (1970). JBC 245, 5169. 254, Gutte, B., Lin, M. C., Caldi, D. G., and Merrifield, R. B. (1972). JBC 247, 4763. 255. Lin, M. C., Gutte, B., Caldi, D. G., Moore, S., and Merrifield, R. B. (1972). JBC 247, 4768. 256. Hayashi, R., Moore, S . , and Merrifield, R. B. (1973). JBC 248, 3889.

358

PETER BLACKBURN A N D STANFORD MOORE

ment of the residues at the active site and not from changes in the binding affinity for the substrate. Hodges and Merrifield (257) synthesized the 14-residue peptide with Tyr or Ala in position 120. In combination with RNase-(1-118), the substitution of Tyr at this position gave a product that had the same activity as the Phe-containing peptide toward RNA and 2',3'-cyclic CMP, but twice the activity toward 2',3'-cyclic UMP. With the Ala substitution, the relative activity toward 2',3'-cyclic CMP was less than 1%. These results led the authors to predict that giraffe RNase, which has a Tyr residue at position 120, would have a higher relative activity toward the UMP substrate than toward cyclic CMP. The prediction was verified; giraffe RNase showed a 2.6-fold greater activity toward the former substrate, taking the k,/K,,, values with bovine RNase as 1 [cf., Ronda et al. (258)l. The X-ray data ( I ) indicate a hydrogen bond between the hydroxyl of Ser-123 and the C-4 carbonyl oxygen of uridine. Potts et al. (259) showed that removal of Val-124 and Ser-123 by carboxypeptidase leaves 45% activity toward RNA; thus it is known that Ser-123 is not crucial. Hodges and Merrifield (260) synthesized RNase-( 111- 124) with Ala in the place of Ser at position 123. When the synthetic peptide was mixed with RNase(1-1 18), the substitution caused no change in activity toward 2',3'-cyclic CMP or in the transphosphorylation step with poly(C); but with 2',3'cyclic UMP the analog was 4 times less active, and with poly(U) two times less active. The results indicate that a hydrogen bond between Ser123 and the C-4 oxygen of uridine may contribute to substrate binding and catalytic activity with the uridine-containing substrate. The Pro residue at position 117 prevents carboxypeptidase A from carrying the degradation from the carboxyl end beyond residue 118. Hayashi et al. (256) tried carboxypeptidase Y, an enzyme capable of releasing proline. Carboxypeptidase Y removed Val-118, Pro-1 17, and Val-116. Tyr- 115 could then be removed by carboxypeptidase A. With the RNase A chain minus the last ten residues RNase-( 1- 114), reconstitution experiments with the nonapeptide RNase-( 116-124), in a combination thus missing Tyr-115, gave a maximum regenerable activity of 54%, which is essentially the same as that observed when the protein moiety was RNase-(1- 118). Tyr-115 is thus not required for activity. The result correlates with the finding of Jackson and Hirs (261) that in porcine RNase a 257. 258. 259. 3781. 260. 261.

Hodges, R. S. , and Merriiield, R. B. (1974). Int. J . Peptide Protein Res. 6 , 397. Ronda, G. J . , Gaastra, W., and Beintema, J. J. (1976). BBA 429, 853. Potts, J. T., Jr., Young, D. M., Anfinsen, C. B., and Sandoval, A. (1%4). JBC 239, Hodges, R. S., and Merrifield, R. B. (1975). JBC 250, 1231. Jackson, R. L., and Hirs, C. H. W. (1970). JBC 245, 637.

12. PANCREATIC RIBONUCLEASES

359

proline residue occupies position 115. The hydrogen bond between Tyr115 and Tyr-73 suggested by X-ray data is not crucial. Consideration of the associations that could hold the nonapeptide RNase-( 116- 124) t o the main chain, by reference to Wyckoff rf rrl. (262), indicates that six hydrogen bonds are possible with RNase-( 1- 114). The stronger binding ( K d = 3.0 x M ) than that obtained with the same peptide and RNase-(1- 118)(256) indicates that when there are overlapping residues in the 115 and 116 positions, there is competition for binding sites between the added peptide and the residual tail of the main chain. The transition temperature curves for RNase-( 1- 114) and the protein moiety plus RNase-(116-124) (see Ref. 285) show a change from a melted state at 25" to a more structured complex with a T,,, of 38". Andria and Taniuchi (263) mixed a tryptic hydrolysate of performic acid-oxidized RNase with RNase-( 1- 118) and gel filtered the mixture. The 105- 124 segment, which represents the residues following the last trypsin-susceptible bond in the sequence (Lys- 104), adsorbed to the RNase moiety that lacked the normal COOH terminus. The combination had about 509% of the activity of RNase A. The presence of RNase-(105124) during the re-formation of -S-Sbonds from reduced RNase-(I118) increased the proportion of the species that exhibited the properties of RNase-( 1- 118) from zero to about 30%. C. CHEMICAL SYNTHESIS The Richards and Wyckoff review on RNase ( I ) recorded as dramatic events of 1969 the synthesis of RNase A (by Gutte and Merrifield) by the solid phase method, and the synthesis of RNase S-protein (by Denkewalter and Hirschmann and their associates) by solution methods. The details of the solid phase synthesis (264) document the isolation of 0.41 mg of purified synthetic enzyme with the substrate specificity of RNase A, and showing 78% of the activity of the native protein toward yeast RNA. A decade later, the synthesis of RNase A by solution methods was accomplished by Yajima and Fugii (265, 266). By including the use of affinity chromatography on pup-Sepharose (14% of the crude synthetic product was retained) they obtained 3 mg of a synthetic protein, which after fur262. Wyckoff, H. W., Tsernoglou, D., Hanson, A. W., Knox, J. R., Lee, B., and Richards, F. M. (1970). JRC 245, 305. 263. Andria, G., and Taniuchi, H. (1978). JBC 253, 2262. 264. Gutte. B., and Merrifield, R. B. (1971). JBC 246, 1922. 265. Yajirna, H., and Fujii, N. (1980). J . Clroni. S o c . Clrern. Couirn., p. 115. 266. Fujii, N . , and Yajima, H. (1981).J. Clwrn. Soc.. Perkin Trans. 1,789,797, 804, 811, 819. 831.

360

PETER BLACKBURN AND STANFORD MOORE

ther ion exchange chromatography was indistinguishable from the native enzyme by chemical and physical criteria and had 100% of the activities of RNase toward RNA and 2’,3’-cyclic cytidylic acid. The chemical synthesis of a 124-residue polypeptide chain represents atorrr de force by present methods, in which yields are in the 1% range. The challenge is stimulating innovations in some of the many steps of the process (267). Gutte (268) has undertaken the synthesis of shorter chains that might have ribonuclease action. From the study of the three-dimensional model of RNase A ( I ) , he aimed at the synthesis of a 70-residue chain that might have sufficient structure to form the central portion of the molecule that comprises the active site. The sequence selected was one in which the 104-residue S-protein structure was shortened by 34 residues; some surface loops and the 8 half-cystine residues were omitted, with Gly, Ala, or Leu inserted to fill seven of the gaps caused by the Cys deletions. The yield of a 70-residue analog of S-protein was, after ion exchange chromatography, about 1% of a product that in the monomeric form (with or without S-peptide) had 0.1% of the activity of RNase toward RNA. For reasons not growing from the original premise, a dimer of the product had 4% activity toward RNA, and the monomeric or dimeric analogs when added to S-protein gave 75 to 150% of the activity of RNase S in the hydrolysis of 2‘ ,3’-cyclic cytidylic acid. In a second approach, Gutte (269, 270) aimed for a 63-residue analog beginning with residue 26 and containing two of the four cystine bridges. The product was first purified by gel filtration. When affinity chromatography (265) was used to select molecules that had an affinity for a substrate analog, 98% of the preparation went straight through; the retarded 2%, isolated in pg quantities, represented an overall yield of about 0.25% of a product that had 7.8% of the activity of RNase A toward poly(C), without the addition of S-peptide. Refolding of the reduced 63residue preparation (or of RNase A) in the presence of mononucleotides (271) produced changes in activities toward synthetic substrates such as poly(C) and poly(U); the conformations thus obtained were thermolabile at 40”. Starting from theoretical considerations based upon secondary structure prediction rules and model building, Gutte et al. (272) designed a 267. Barany, G . , and Merrifieid, R. B. (1979). In “The Peptides” (E. Gross and J . Meienhofer, eds.), Vol. 2, p. 1. Academic Press, New York. 268. Gutte, B. (1975). JEC 250, 889. 269. Gutte, B . (1977). JEC 252, 663. 270. Gutte, B. (1978). JEC 253, 3837. 271. Gutte, B . (1978). EJE 92, 403. 272. Gutte, B., Daumigen, M., and Wittschieber, E. (1979). Nnture (London) 281, 650.

12. PANCREATIC RIBONUCLEASES

36 1

34-residue peptide structure with potential binding power for the anticodon of yeast tRNAPhe.The synthetic peptide (with two cysteine residues) forms an - S - S cross-linked dimer that binds tRNA and 2',3'-cyclic nucleotides. It also cleaves tRNAPheand poly(C); the activity toward yeast tRNA was 2.5% of that of RNase A. The product of the transphosphorylation step, the 2',3'-cycIic nucleotide, was not hydrolyzed. These experiments were designed to ascertain the feasibility of formulating polypeptides that would be artificial enzymes. The stimulating results demonstrate that structures can be built that fulfill predictions for ability to bind nucleotides, and that such a binding can labilize a phosphodiester bond to transphosphorylation. The data on the 70-residue and the 63-residue RNase A analogs have limited bearing on the specific features of pancreatic RNase per se. The 34-residue study shows most successfully that the ability to induce transphosphorylation in an RNA can be built into a polypeptide chain in a number of ways. When the yields of active product are as low as they are in the syntheses of the 70- and 63-residue peptides, it is difficult to assign definitive structures to the active fractions isolated in microgram amounts. Nature's design of a pancreatic enzyme also incorporates properties that are essential to the protein's function in vivo. The simplified analogs are readily hydrolyzed by proteolytic enzymes such as trypsin; the native structure of RNase A is uniquely constructed and cross-linked to render the catalyst resistant to proteolysis. The entire structure of RNase A is an entity that can function in the gastrointestinal tract. Semisynthetic studies to examine the roles of residues in the NHz- and COOH-terminal sections of RNase are summarized in Section II1,B.

D. BIOSYNTHESIS The signal hypothesis (273) for secreted proteins would predict that bovine pancreatic RNase should be synthesized as a presecretory protein. Demonstration of this fact has awaited a method for preparing intact mRNA from bovine pancreas by the employment of conditions that inactivate the RNase that otherwise destroys the message in a homogenate of the tissue. The method of Chirgwin ef crl. (274), using 5 M guanidinium thiocyanate plus 0.1 M mercaptoethanol in the extraction medium, opened the way to the in vi?ro translation experiment. Haugen and Heath (275) prepared mRNA by that procedure and conducted the translation with the 273. Blobel, G . , and Dobberstein, B. (1975). J . Cell B i d . 67, 852. 274. Chirgwin, J. M . , Rzybyla, A. E., MacDonald, R. J., and Rutter, W. J. (1979). Biochemi.str.v 18, 5294. 275. Haugen, T. H . , and Heath, E. C . (1979). P N A S 76, 2689.

362

PETER BLACKBURN AND STANFORD MOORE

rabbit reticulocyte lysate system. The pre-RNase was isolated by precipitation with antibody to bovine pancreatic RNase. The product showed a MW of 16,500 by SDS gel electrophoresis. Sequence analysis of the [14C]Ala-labeledprotein showed the presence of a 25-residue segment ahead of the normal NH2 terminus. When incubated with dog pancreas microsomal membranes, the product yielded a protein that comigrated with RNase in gel electrophoresis; a band corresponding to RNase B was shown to be the glycosylated enzyme. Although assays for enzymatic activity on the picogram quantities of protein synthesized are difficult, the results indicate that pre-RNase A can fold into an enzymatically active form. Since RNase is a pancreatic enzyme that does not have a zymogen form, it is understandable that the presecretory (the signaled form) could have an active conformation. The authors reflect on the potentially damaging effect of any RNase activity that might reach the cytoplasm, and suggest that the cytoplasmic RNase inhibitor may have a function in this regard (275). E . IMMu N oc HE M ISTRY

Experiments on antibody-mediated modification of RNase activity have been reviewed by Cinader (276). Rabbit sera drawn on the 255th day after the start of immunization were chromatographed on DEAE-Sephadex A-50 (277). One fraction contained a small quantity of antibody that increased the activity (up to twofold) of the enzyme toward 2’,3‘-cyclic CMP (the activity toward RNA remained unchanged), other fractions contained antibodies that inhibited the enzyme’s actions up to 80% with the cyclic substrate and 98% with RNA. The effects leveled off at high antibody concentrations. The activating antibody gave a similar response with RNase S (278). In Section IV,D, on the refolding of reduced RNase, we refer to the ways in which Chavez and Scheraga used their findings on the antigenic sites of the enzyme to follow the folding process. They reviewed (279) the previous literature on the location of antigenic determinants of the native enzyme and applied a variety of immunochemical techniques to the definition of four antigenic segments of the chain. The coupling of peptides from RNase to Sephadex for the isolation of antibodies to given segments was a 276. Cinader, B. (1977). I n “Methods in Immunology and Immunochemistry” (C. A. Williams and M. W. Chase, eds.), Vol. IV, p. 313. Academic Press, New York. 277. Suzuki, T., Pelichova, H . , and Cinader, B. (1969). J . Immunol. 103, 1366. 278. Cinader, B., Suzuki, T., and Pelichova, H. (1971). J . Immunol. 106, 1381. 279. Chavez, L. G . , Jr., and Scheraga, H. A. (1979). Biochemistry 18, 4386.

12. PANCREATIC RIBONUCLEASES

363

key step in their analysis. Working with late hyperimmune antisera, four sites in bovine RNase A were localized within residues 1-10, 40-61, 63-75, and 87-104; the data do not provide information on the possible antigenicity of the region 30-39. Welling et a / . (280) conducted an immunological comparison of pancreatic RNases from nine species with antisera toward four of the species. Cross-reactivities by Ouchterlony double-immunodiffusion tests ranged from identity, through partial identity, to no identity. The complement fixation titers varied over a 200-fold range. Lee ef al. (281) found that sheep antibodies to purified rabbit spleen RNase gave only one precipitin band with rabbit pancreatic RNase, thus providing immunological evidence on the similarity of the enzymes from the two rabbit tissues. However, the antibody to the rabbit enzyme did not react with bovine o r rat pancreatic RNases. Working with antisera to bovine RNase A, Welling and Groen (282) focused on the relative antigenicities of six pancreatic RNases rather closely related in sequence; the tests were made by competition experiments employing the modified phage technique of Haimovich el al. (283). Through consideration of both the immunological results and the known sequence substitutions, they concluded that residues 34, 35, 103, and 50 and/or 99 are parts of antigenically reactive regions in bovine RNase A. These results are consistent with those of Chavez and Scheraga (279). Bovine RNase B, which has a carbohydrate side chain on Asn-34, was measurably less competitive than RNase A in the tests by Welling and Groen (282), confirming that this region of the enzyme has an input into the reaction with antiserum. A synthetic peptide comprising residues 1-14, at a molar excess over RNase up to lo6, did not inhibit phage inactivation (282). Chavez and Scheraga (279) suggest that their positive result for this region may depend upon the use of late hyperimmune sera. The experiments of Brown and associates on the antigenic regions of performic acid-oxidized RNase have been summarized (284). There is minimal immuno-cross-reactivity between the oxidized chain and the active enzyme; one section, residues 38 (or 40)-61, is antigenic in both the native and oxidized molecules. 280. Welling, G . W., Groen, G . , Beintema, J. J . , Emmens, M., and Schroder, F. P. (1976). Imrnrrnach~mistrv 13, 653. 281. Lee, W. Y., Gyenes L.. and Sehon, A. H. (1971). ~ ~ ~ u n a ~ h ~8,m751. i . ~ t ~ y 282. Welling, G. W., and Groen, G. (1976). BBA 446, 331. 283. Haimovich, J., Hurwitz, E., Novik, N., and Sela, M. (1970). BBA 207, 115; ;bid, 125.

284. Liu, S., Johnsen, E . , and Brown, R. K. (1974). lmmurzochemistry 11, 55.

364

PETER BLACKBURN AND STANFORD MOORE

Reports on the immunosuppressive action of polyribonucleases prepared by cross-linkage with bisdiazobenzidine ( 2 8 5 , 2 8 5 ~have ) not been confirmed (286).

IV.

Physical Properties

The following sections cover studies subsequent to the definition of the physical param2ters of RNase summarized in a chapter in this series (1). A. X-RAYDIFFRACTION Richards, Wyckoff and colleagues (1, 16, 286n) have thoroughly reported and interpreted the extensive data on the three-dimensional structure of RNase S . Subsequent results include those of Wodak et al. (21) on the binding of 2',5'-CpA, a substrate analog that binds to the enzyme but is not cleaved. Their results reinforce the evidence obtained with other dinucleotides for a hydrogen bond between the N-3 of cytosine and the 7-OH of Thr-45. The orientation of the cytosine ribose in 2',5'-CpA in the binding site ;s such that the 3'-OH is away from the active center and not in a position to be rendered more nucleophilic to facilitate a transphosphorylation. The results of White et a/. (287) indicated little interaction of 2',5'-UpA or 2',5'-4-thio-UpA with RNase A, in solution in 0.2 M imidazole buffer at pH 7.0, both in terms of inhibition of activity and direct binding by CD measurements. The combination of RNase S and the analog of 3'3'-UpA with a fluorine atom replacing the 2'-OH has been studied crystallographically by Pavlovsky et (11. (73). The data indicate that the distance from F-2' to N-1 of His-12 is 3 A, a result that is consistent with the positioning of a normal substrate with a hydrogen bond between the 2'-OH and His-12 (1). Allewell et al. (40) have examined the crystallographic structure of edinitrophenyl-Lys-41-RNase S. The derivative and the crystals were prepared by Fung and Doscher (199). They first prepared eDNP-Lys-41RNase A in a chromatographically purified form that had less than 0.001% of the activity of RNase A. That derivative was cleaved by subtilopepti285. Mowbray, J . F., and Scholand, J . (1966). Imrnrrnolngv 11, 421. 285a. Mowbray, J. F. (1967).Svmp. Tissue Org. Trnnsplnnt ( S u p p l . , J . Clin. Paihol.) u), 499. 286. Chakrabarty, A . K., and Friedman, H . (1970). Clin. Exp. Immunol. 6, 619. 286a. Richards, F. M., Wyckoff, H. W., Carlson, W. D., Allewell, N. M., Lee, B . , and Mitsui, Y. (1971). Cold Spring Harbor Svmp. Qunnt. B i d . 36, 35. 287. White, M. D., Keren-Zur, M., and Lapidot, Y. (1977). N i d e i c Acids Res. 4, 843.

12. PANCREATIC RIBONUCLEASES

365

dase A to give the RNase S analog. In solution, the inactive DNPsubstituted protein binds nucleotides such as 3'-CMP about one-tenth as strongly as does the parent enzyme; crystals of the RNase A and S derivatives show the same affinity for 3'-CMP. The presence of the DNP group displaces the E-N of Lys-41 by about 3 A; His-12 and His-119 are not moved. The loss of activity appears to result primarily from perturbation of the molecule in the region of Lys-41. H. C. Taylor has advised us of results (2870) on the crystal structure of the semisynthetic RNase S in which 4-fluoro-His replaces His- 12 in the synthetic segment of residues 1-15 (288). The overall structure, including the positioning of His-12, does not differ detectably from that of normal RNase S; the data support the view that the 4-F-His analog is inactive as a result of the lowered pK, of His-12 rather than distortion of the active site. The inhibition of RNase S by Cu'+ has been studied by Allewell and Wyckoff (289). They identified 7 binding sites, 3 of which were intermolecular. Two of the intramolecular sites are close to the active-site histidine residues 12 and 119 and could be expected to cause the observed inhibition. The effect of the pH of crystallization on the conformation of RNase has been examined by Martin et cd. (290). Crystals of RNase A and of RNase S formed at pH 9 have been found to be isomorphous with those of RNase S crystallized at pH 6.6. X-Ray diffraction shows small but significant intensity differences comparable in magnitude to those observed between RNase A and RNase S. The results of the crystallographic studies on RNase A by Carlisle and associates (119) are in general agreement with the data previously available for RNase A and RNase S ( I ) in terms of the relationships of His-12, His-119, and Lys-41 at the active center of the catalyst. However, the peptide bond between residues 20 and 21 restricts the conformational freedom of the S-peptide segment compared to that found with RNase S . Timchenko et a / . (291) have studied RNase A in solution by large-angle X-ray scattering, which is sensitive to the internal structure of globular proteins in solution. The observed scattering curve was compared with the curve calculated from the atomic coordinates available from X-ray 287a. Taylor, H . C., Richardson, D. C., Richardson, J. S., Wlodawer, A , , Komoriya, A,, and Chaiken, I . M . (1981). J M B . 149, 313. 288. Dunn, B . M . , DiBello, C . , Kirk, K . L., Cohen, L. A . , and Chaiken, I . M . (1974). JBC 249, 6295. 289. Allewell, N . M . , and Wyckoff, H. W. (1971). JBC 246, 4657. 290. Martin, P. D . , Petsko, G . A . , and Tsernoglou, D . (1976). J M B 108, 265. 291. Timchenko, A. A . , Ptitsyn, 0. B., Dolgikh, D. A , , and Fedorov, B. A. (1978). FEBS Lett. 88, 105.

366

PETER BLACKBURN AND STANFORD MOORE

analysis. The conclusion was that the structure in solution does not differ significantly from that in the crystal. Wlodawer (292) has undertaken a computational refinement of the available crystal structure coordinates af RNase A for comparison with results of neutron diffraction on deuterated crystals. Neutron diffraction can provide detailed information on the orientation of the imidazole rings of histidine residues. Large crystals (1 x 5 x 6 mm), successfully prepared in about 50% rerr-butyl alcohol at pH 5.3, were used. The results (Wlodawer and Sjolin, 292a) give evidence for a hydrogen bond between His-119 and Asp-121 in crystals suspended in deuterated tert-butyl alcohol-D20. B. NUCLEAR MAGNETIC RESONANCE In the early NMR work on RNase A, Meadows et al. (237,293,294)and Ruterjans and Witzel (77) differentiated between the C-2 proton resonances (HI, H2, H3, and H4) of the four histidine residues based upon their titration curves before and after carboxymethylation and upon cytidine monophosphate binding. Meadows et al. (237) differentiated between the His-12 and His-119 C-2 proton resonances by a comparison of the titration curves of RNase S and RNase S’ reconstituted with S-peptide deuterated at the C-2 proton of His-12, and assigned the resonance with the lower p K, value to His- 119. They were supported in their assignment by results from the deuterium exchange studies of Bradbury and Chapman (295). However, subsequent studies have shown that these original assignments of the active site histidine proton resonances were incorrect; current assignments are included in Table VII (see also Refs. 296-303). 292. Wlodawer, A. (1980). Acto Crystallogr. 836. 1826. 292a. Wlodawer, A., and Sjolin, L. (1981). PNAS 78, 2853. 293. Meadows, D . H., Markley, J. L., Cohen, J. S., and Jardetzky, 0. (1967). PNAS 58, 1307. 294. Meadows, D. H., and Jardetzky, 0. (1968). PNAS 61, 406. 295. Bradbury, J. H . , and Chapman, B. E. (1972). BBRC 49, 891. 296. Markley, J. L. (1975). Biochemisfn’ 14, 3546. 297. Shindo, H . , Hayes, M. B . , and Cohen, J . S. (1976). JBC 251, 2644. 298. Bradbury, J. H., Crompton, M. W., and Teh, J. S. (1977). EJB 81, 411. 299. King, N . L. R., and Bradbury, J. H. (1971). Nature (London) 229, 404. 300. Meadows, D. H., Roberts, G. C. K., and Jardetzky, 0. (1969). JMB 45, 491. 301. Kaptein, R . , Dijkstra, K . , Muller, F., van Schagen, C. G . , and Visser, A . J . W. G. (1978). J . Mugneric Resonanr‘e 31, 171. 302. Kaptein, R., Dijkstra, K., and Nicolay, K. (1978). Nature (London) 274, 293. 303. Kaptein, R. (1978). In “Nuclear Magnetic Resonance Spectroscopy in Molecular Biology” (B. Pullman, ed.), p. 21 1 . Reidel, Dordrecht, Netherlands.

367

12. PANCREATIC RlBONUCLEASES TABLE VII SUMMARY O F

Resonanceb) H1 H2 H3 H4' HI' H2' H3'

Yl" y2d Y3"

THE

ASSIGNMENTS O F AROMATIC PROTON RESONANCES OF BOVINERIBONUCLEASE A"

Chemical shiftb (ppm)

8.08 7.91 7.77 7.18 6.70 6.43

( ( 7.19 6.76 { ;:I

Assignment His-105, C-2H His-119, C-2H His-12, C-2H His-48, C-2H His-105, C-4H His-1 19, C-4H His-12. C-4H

I

Tyr-76* C"H Tyr-76, Cf-H Tyr-115, Cd-H Tyr-115, Cf-H

Y5

6.90 6.70

Tyr-25, C'-H Tyr-92, Cs-H + Cf-H Tyr-73, CS-H or Cf-H

F3 F4

6.83 6.61

Phe-120, ring protons Phe-46, ring protons

Y4

I

6.3-6.5

Phe-120, ring protons near active-site inhibitor

Reference or evidence (78. 237, 296-298) (237, 296-298) (237, 296-298) (237, 296. 298) (237, 296, 299) (299); CIDNP spectrum' (299); Comparison with RNase S ; pH midpoint; effect of inhibitors Comparison with nitrated RNase Comparison with nitrated RNase Comparison with RNase S; pH titration Alkaline titration pH dependence in nitrated RNases Effect of active-site inhibitors Comparison with RNase S; pH titration curve (300)

From Lenstra et r i l . (80). The last five assignments are tentative. "The chemical shifts were measured at pH 7.0, 38", downfield from sodium 2,2dimethyl-2-silapentane 5-sulfonate. ' Not visible at pH 7.0. The upfield doublets are assigned to tyrosine C' protons on the basis of the polarizations in the CIDNP (chemically induced dynamic nuclear polarization) spectra. In the CIDNP technique (301-303), the appearance of resonances from histidine, tyrosine, and tryptophan residues in these spectra depends upon the accessibility of the aromatic ring systems to photo-excited flavin, resulting in nuclear spin polarization through reversible hydrogen or electron transfer (302. 303 ).

"

1. Histidine Assignments

A resonance, designated n , downfield of the C-2 proton resonances of histidine residues was observed by Patel et NI. (304) and was assigned to a 304. Patel, D. J., Woodward, C. K., and Bovey, F. A. (1972). PNAS 69, 599.

368

PETERBLACKBURN ANDSTANFORDMOORE

slowly exchangeable, possibly buried or hydrogen bonded imidazole N-H whose chemical shift and linewidth are governed by ionization of the other imidazole ring nitrogen. In the presence of 3'-CMP the p K , values of the active-site histidine residues are increased by > 1 pH unit (see Table VIII) (see also refs. 305-307), whereas the pK, values of His-48 and His-I05 were found to increase by only 0.1 pH unit (214). The p K , of resonance a increases from 6.3 to 7.2 in the presence of 3'-CMP (214), and this rules out its assignment to His-48 or His-105. In N-l-carboxymethyl-His119-RNase A, des-(121- 124)-RNase A, and DNP-Lys-41-RNase A, resonance (i titrates with constant band area and a pK, -6.8, which rules out its assignment to N-3 of His-119 and the e N H 2 group of Lys-41. In N-3-carboxymethyl-His-12-RNase A, resonance CI is observed with the chemical shift of a protonated histidine residue; it does not tirate, but disappears at alkaline pH as the proton exchanges with solvent. Resonance n was thus assigned to the N-1 proton of His-12 (214), which is proposed to hydrogen bond to the peptidyl carbonyl of Thr-45 ( 1 , 203). Based upon the titration behavior of resonance (I in the presence and absence of phosphopyrimidine nucleotides, by correlation with the previous histidine C-2 proton assignments (237), resonance a had been incorrectly assigned to the N-H of His-119 (308). The ionization characteristics of resonance fi were associated with the active-site histidine C-2 proton resonance that exhibits the lower p K, (H3) on the basis of Gun, Ag', and spin labeling data (309) and active-site inhibitor binding (214,308), suggesting that the histidine C-2 proton resonance with the lower pK, (H3) is from His-12 (78),and not from His-119, as was previously indicated (237, 294). The assignments of the C-2 proton resonances of His-12 and His-119 were thus shown to require reinvestigation. Patel et a/. (78) demonstrated in RNase S and RNase S', reconstituted with S-peptide deuterated at the C-2 proton of His-12, and by Cu" and 3'-CMP binding, that the resonance with the lower pK, (H3) should be assigned to His-12. Markley (296) from a comparison of the deuterium exchange kinetics of the histidine C-2 protons of RNase A, measured by NMR, with the order of tritium exchange rates into the individual his305. Griffin, J . H., Schechter, A. N., and Cohen, J. S . (1973).Awi. N. Y. Actrtl. Sci. 222, 693. 306. Haar, W., Maurer, W., and Riiterjans, H. H. (1974). EJB 44, 281. 307. Haffner, P. H., and Wang, J. H. (1973). Biochenrisfry 12, 1608. 308. Griffin, J. H., Cohen, J. S., and Schechter, A. N. (1973). Biochemistry 12, 2096. 309. Patel, D.J . , Woodward, C., Canuel, L. L., and Bovey, F. A. (1975). Biopdymers 14, 975.

12. PANCREATIC RIBONUCLEASES

369

tidine residues ( 3 / U ) , confirmed that resonances H4 and H1 could be assigned to His-48 and His-105 (237, 294) but that H3 and H2 should be reassigned to His-12 and His-1 19, respectively. Cohen et a / . (238) demonstrated that the C-2 proton resonances of the active-site histidine residues are essentially identical in RNase A and RNase S. Meadowset a/. (237) had concluded that these resonances were different. These differences were shown to be most probably a result of phosphate in the sample of RNase S (238). Moreover, all four C-2 proton resonances are resolved at pH >5.5 in RNase S. By a comparison of the rate of tritium exchange of the C-2 proton of His-12 of S-peptide with the rates of deuterium exchange of individual C-2 protons of the histidine residues of RNase S, Shindo et 01. (297) confirmed the reassignment of resonance H3 to His-12. A comparison of the titration of the imidazole C-2 protons of N- 1-carboxymethyl-His-119-RNase A with those of the model compounds N-1-carboxymethyl- and N-3-carboxymethylhistidine (p K, values of 6.5 and 5.9, respectively) and N- l,N-3-dicarboxymethylhistidine, permitted reassignment of both resonances H3 and H2 to His-12 and His-1 19 respectively. A, the p K, of His-12 is elevated In N-1-carboxymethyl-His-119-RNase by 1.1 pH units to 7.17 as a result of interaction with the carboxyl group on the modified His-119 side chain. The C-2 proton resonance of the imidazole moiety of carboxymethyl-His-119 was undetected; this possibly indicates that its motion is restricted upon carboxymethylation (297). From the data of Meadows et a/. (237), in the light of the new assignments, it can be concluded that His-12 has a pK, of 6.9 inN-1-carboxymethyl-His119-RNase A, and His-1 19 has a pK, of 6.7 inN-3-carboxymethyl-His-12RNase A (a change of 0.5 units upon derivatization). The pK, values and chemical shifts of the resonances of His-48 and His-105 were not affected in these derivatives. The increased pK, values of the active-site histidine residues in the carboxymethylated derivatives arise from local electrostatic interactions. The reassignment of the H2 and H3 C-2 proton resonances of the active-site histidine residues requires that work (18, 42, 71, 77, 300, 307, 311,312,313) based on the assignments of Meadows l’f d . (237) should be interpreted in this light. Accordingly, where referred to in this review, the appropriate reassignments of the H2 and H3 resonances have been made to the C-2 protons of His-I19 and His-12, respectively. 310. Ohe, M . , Matsuo, H . , Sakiyama, F., and Narita, K . (1974). J . Biochewz. ( T o k y o ) , 75, 1197. 311. Westmoreland, D. G . , and Matthews, C. R. (1973). P N A S 70, 914. 312. Benz, F. M . , and Roberts, G. C. K . (1975). J M B 91, 345. 313. Benz, F. M., and Roberts, G . C. K . (1975). J M B 91, 367.

370

PETER BLACKBURN AND STANFORD MOORE

“1

t

RNase A.

I

7-0

I

I

I

1

9

8

7

6

6 (Ppm) FIG.3. Aromatic region of the 360 MHz proton NMR spectrum of RNase A. The prefixes H, Y, and F denote assignments to histidine, tyrosine, and phenylalanine residues, respectively. From Lenstra ef c d . (80).

The high resolution, 360 MHz proton NMR spectra of RNase A have been reported by Lenstraer a/. (80). Most of the resonances in the aromatic region have been assigned to specific residues (Fig. 3). The assignments, where identified, and their chemical shifts are presented in Table VII. The two spectra shown in Fig. 3, recorded at pH 7.0 and pH 3.4, illustrate resonances that undergo a pH-dependent chemical shift, presented in more detail in Fig. 4. Four singlets H1, H2, H3, and H1’ are identified (80) as histidine resonances on the basis of their titration curves (78, 237, 296-299, 314). The singlet H4 is not observed between pH 5.2 and pH 7.9 (237, 315, 316) except in acetate buffer (79,315,316) and is assigned to the C-2 proton of His-48 (237, 296, 298). H2’ was assigned to the C-4 proton of His-119 based on results from laser-induced photo-CIDNP spectra (see footnote to Table VII) and the effect of the active-site inhibitor 2’-CMP, associating it with the H2 resonance of the C-2 proton of His-119 assigned previously (214, 2%-298). 314. Meadows, D. H. (1972). “Methods in Enzymology” Vol. 26, p. 638. 315. Roberts, G . C. K., Meadows, D. H., and Jardetzky, 0 . (1969). Biochc~niisrry8,2053. 316. Markley, J. L. (1975). Biorhemi.sfry 14, 3554.

12.

37 1

PANCREATIC RIBONUCLEASES 90

85

\'2

-44

.\

80

7s 72

71

g 70 Q

1

og

i

\

69

68

61

66

65

64

3

4

S

6

7

e

PH FIG.4. The pH dependencies between pH 3.4 and 8.0 of the chemical shifts of aromatic proton resonances of RNase A. The prefixes are those defined for Fig. 3. Crosses, triangular crosses, and triangles denote singlets representing one proton; squares denote doublets, and circles denote nonresolved multiplets. From Lenstra et rrl. (80).

372

PETERBLACKBURN ANDSTANFORDMOORE

Unlike RNase A at neutral pH, RNase S exhibits the H4 resonance of the C-2 proton of His-48 (77, 294, 315). Comparison of the NMR spectra of RNase A and RNase S permitted assignment of resonance H4’ to the C-4 proton of His-48 and thus the remaining resonance, H3’, was assigned to the C-4 proton of His-12 (80). The assignments of H2’ and H3’, as shown here, to the C-4 protons of His-119 and His-12, respectively, as reported by Lenstraet al. (80),agree with the assignments of King and Bradbury (299) and the assignment of H2‘ by Markley (296). However, Markley had assigned the resonance reported here as H3’ (80) to the C-4 proton of His-48. 2. Tvrosine Assigrimerits The proton resonances of tyrosine rings that rotate rapidly about their CB- Cybonds are represented by two mutually coupled doublets, both . such pairs were revealed by corresponding to two protons ( 3 1 6 ~ )Three double resonance experiments (80); they are Y 1, Y2, and Y3. Two other resonances that do not appear as mutually coupled doublets are Y4 and Y5. Y1, Y2, Y4, and Y5 exhibit alkaline shifts between pH 9 and 11, and therefore titrate as normal tyrosine residues. Earlier, Egan et ul. (135) had also found four normally titrating tyrosine residues in 13C NMR studies. This conclusion is contrary to the data obtained by spectrophotometric titration of the tyrosine residues of RNase A [reviewed by Richards and Wyckoff (1) and in Section III,A,7], which indicate three tyrosine residues with normal pK, values and three with abnormally high pK, values. The resonances Y1 and Y2 are identified as being due to exposed tyrosine residues from photo-CIDNP spectra of RNase A (80).They were assigned to Tyr-76 and Tyr-115, respectively, after their elimination from the NMR spectrum following selective nitration of Tyr-115 and Tyr-115 plus Tyr-76 of RNase A with tetranitromethane according to van der Zee et a / . (155).The titration behavior of resonance Y5 between pH 6 and pH 7 in RNase A nitrated at either Tyr-115 or both Tyr-115 and Tyr-76 indicated that Y5 is due to a normally titrating tyrosine residue close to the 3-nitro-Tyr-115 phenolic hydroxyl, which has a pK, of 6.3, and thus Y5 was assigned to Tyr-73 (80). The photo-CIDNP spectra of RNase A and RNase S are essentially identical except for an emission at the position of the upfield Y3 doublet; thus, the tyrosine residue corresponding to Y3 is accessible in RNase S but not in RNase A, and has been assigned to Tyr-25 (80). The fourth normally titrating resonance Y2 is assigned to Tyr-92 (80). 316a. Campbell, I . D . , Dobson, C. M., and Williams, R. J. P. (1975). Proc. Roy. S O C . London A345, 23.

12. PANCREATIC RIBONUCLEASES

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The two sharp doublets of Y3 in the spectrum of RNase A, seen at low pH are exhibited by RNase S at both neutral and low pH, and indicate rotation about the Cp-Ccy bond of Tyr-25. At neutral pH these Y3 signals are less sharp in RNase A, indicating this movement is more hindered at neutral pH in RNase A. Y3 exhibits a pH-dependent chemical shift with a midpoint at pH 6.2; the other tyrosine resonances are unaffected in this pH range. Lenstra et al. (80) were unable to locate the resonances assigned by Markley (316) to Tyr-25. 3. Conformational Trcinsition Involving H i s 4 The different titration behavior of His-48 in the proton NMR spectra of RNase A in the presence and absence of acetate (294, 315-317), and its different behavior in the proton NMR spectra of RNase A and RNase S, (77, 238) led to the suggestion that His-48 is involved in a local conformational transition of the protein (237, 238, 294, 315-317). The C-2 proton resonance of His-48 of RNase A, observed in acetate buffer, exhibits an inflection in its titration curve near pH 4.2 (77-79, 238); this inflection is not observed with RNase S-protein (239). Moreover, the active-site histidine residues of RNase A (77, 79, 238, 317, 318) and RNase S (79) also exhibit an inflection near pH 4.5 in the titration curves of their C-2 proton resonances. Markley and Finkenstadt (318)have proposed a model based on mutual interaction between His-12 and His-119 to describe the pH titration curves of the respective C-2 proton resonances. They suggest that the low pH inflections of the titration curves arise from a conformational transition with a midpoint at pH 3.7. Cohen and Shindo (79) proposed that the inflections at pH -4.2 in the NMR titration curves of histidine residues 12, 119, and 48 of RNase A are derived from a common event, namely a conformational change involving the side chains of Asp-14 and His-48. In the crystal structure of RNase S ( I )and RNase A (119), the p-COOH of Asp-14 is hydrogen-bonded to the phenolic hydroxyl of Tyr-25. The proton NMR data of Lenstra et 01. (80) for the Y3 doublet assigned to Tyr-25 indicate that in solution at acidic pH, Tyr-25 is not involved in a hydrogen bond. They suggest that the behavior of the Tyr-25 (Y3) resonances may be explained by the breaking of the Tyr-25-Asp-14 hydrogen bond as His-48 is protonated, with formation of a His-48-Asp-14 hydrogen bond. Riiterjans and Witzel (77), and later Santoro et a/. ( 7 4 , based 317. Schechter, A. N . , Sachs, D. H . , Heller, S. R., Shrager, R. I . , and Cohen, J. S. (1972). JMB 71, 39. 318. Markley, J. L., and Finkenstadt, W. R. (1975). Biochemistry 14, 3562.

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upon the titration behavior of resonances assigned to Tyr-25 and the /3-COOS of Asp-14 in the proton-decoupled I3C NMR spectra of RNase A, have also proposed a hydrogen bond between Asp-14 and His-48 at acidic pH. They propose that as His-48 is deprotonated, a hydrogen bond forms between the Tyr-25 phenolic group and the Asp- 14 p-COOH or with one of the ring nitrogens of His-48. I3C NMR studies by Cohen and his associates (203,203a),working with the RNase S' complex in which an S-peptide of residues 1 through 15 was synthesized with 13C-enrichedamino acids at His-12 (C'), Met-13 ((3, and Asp-14 (Cy), obtained a pK, of 2.4 for the p-COOH of Asp-14 in the complex by curve fitting $0 the data points; a p Ka of 3.8 was obtained in the free peptide. The titrabion curve of Asp-14 exhibits an inflection with a pK, of 6.1, thought to originate from His-48. On the other hand, the PKa of 2.4 of Asp-14 in this RNase S' complex is too low to account for the inflection seen at pH 4.2 in the proton NMR titration curve of His-48 and those of His-12 and His-119 of RNase A and RNase S. The C' resonance of His-12 of this RNase S' complex exhibited a single pH transition with a pK, of 5.7; an inflection at pH 4.2 was not observed. Santoro et a/. (74) identified several titrating resonances in the carbonyl and carboxyl region of the I3C NMR spectra of RNase A. They tentatively assigned two of these, one to the a-carboxyl of Val-124 (pKa 3.5) and the other to the /3-carboxyl of Asp-14 (pKa 4.33), which demonstrated an inflection and peak splitting with a midpoint in the pH range 6.5 to 7.0, thought to be a result of interaction with His-48. In view of the results of Niu et ol. (203, 203a), this assignment to Asp-14 may require reexamination, but should not be ruled out. 4. His-I 19-Asp121 Interaction

An interaction between these residues was proposed by Sacharovsky et

d.(71). Such an interaction could perhaps explain the results obtained by Santoro et ul. (74) previously described. However, they consider this unlikely since they observed a broadening and splitting of the carboxyl resonance around the imidazole p K,, which they suggest is from His-48. Also, based on curve fitting to their data on the low pH inflection for the C-2 proton resonance of His-119, Cohen and Shindo (79) consider this His-1 19-Asp-121 interaction unlikely. Deprotonation of the imidazolium form of a histidine residue can yield either the Nc2-H(N-3-H)tautomer or the N"-H (N-1-H) tautomer of the imidazole form of the residue. From the titration behavior of the resonances of the nonprotonated aromatic carbons assigned to the four histidine residues in the I3C NMR spectrum of RNase A in acetate buffer, it was found by Walters and Allerhand (75)that only His- 119 exists predom-

12. PANCREATIC RIBONUCLEASES

375

inantly as the N"-H tautomer. The imidazole forms of His-12 and His-48 are predominantly in the N6'-H tautomeric form; His- I05 appears to be between 50 to 90% as the N"-H tautomer. The NC2-Htautomer is predominant for 1-histidine and a number of L-histidyl peptides (319-321). Stabilization of the N6'-H tautomeric forms in the protein is thought to result from hydrogen bonds between residues His-48 and Asp-14, His-I2 and Thr-45, and between NS'-H of His-105 and the a-carboxyl group of Val-124 ( 7 3 . Thus, evidence for a hydrogen bond between the p-COOS of Asp-121 and NC2of His-119 was not found by this technique. 5 . Lysine Amino Groirps

Direct observation of the lysine e N H 2 groups of proteins by NMR spectroscopy is not possible due to the rapid rates of exchange of the amine protons. Lysine residues may be studied, after reductive methylation with formaldehyde and borohydride, by the proton resonances of the resultant N-methyl groups (18, 322). Titration of the N-methyl proton resonances of fully reductively methylated RNase A permitted assignments for the derivatives of the a-NHz group of Lys-1 and the e N H 2 group of Lys-41, and enabled pK, values, corrected for the effects of methylation (323), of 6.6 and 8.8 to be determined, respectively, for their amino groups (18). The pK, values obtained for the other nine lysine e N H 2 groups were between 10.2 and 10.8 (18). The pK, of 8.8 obtained for the e N H 2groups of Lys-41 agrees closely with previously determined values (41, 324, 325). Lys-41 is the last lysine residue to react upon guanidination with l-guanyl-3,5-dimethylpyrazole( I 7); reaction of Lys-41 is accompanied by greater than 95% loss of enzymatic activity (19). The proton NMR spectra of RNase guanidinated on 9 or 10 lysine residues and of RNase A reductively methylated are essentially identical with that of the native enzyme and undergo similar changes on thermal unfolding (18, 19). Following reductive methylation of the e N H Zgroup of Lys-41 of RNase guanidinated on 9 lysine NH2groups, titration of the N-methyl proton resonances indicated a pK, of 8.8 for the e N H 2 group of this residue. The major loss of activity upon modification of Lys-41 does not appear to result from any 319. Reynolds, W. F., Peat, I . R., Freedman, M . H . , and Lyerla, J. R., Jr. (1973). JACS 95, 328. 320. Wasylishen, R. E., and Tomlinson, G. (1977). Cnn. J . Biorhcm. 55, 579. 321. Blomberg, F., Maurer, W., Riiterjans, H. (1977). JACS 99, 8149. 322. Bradbury, J. H . , and Brown, L. R . (1973). EJB 40, 565. 323. Perrin, D. D. (1964). Airst. J . Cl7m. 17, 484. 324. Murdock, A. L . , Grist, K. L., and Hirs, C . H . W. (1966). A B B . 114, 375. 325. Carty, R . P., and Hirs, C . H. W. (1968). JBC 243, 5244.

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PETER BLACKBURN AND STANFORD MOORE

major conformational change in the protein arising from the chemical modifications. Moreover, similar chemical shifts and pK, values for the active-site histidine residues were observed for the fully methylated and native RNase (18). Upon binding phosphate, the pK, of EN-methyl-Lys-41 was increased by 0.3 pH units. Similar pK, values for the respective lysine residues of RNase were obtained by Jentoft et al. (20, 34) in I3C NMR studies after reductive methylation of the enzyme with [‘3C]formaldehyde and cyanoborohydride. Binding of 3’-CMP by the fully methylated RNase caused a small but significant shift in the resonance assigned to eN.N‘-dimethyl-Lys-41 but was without effect on the other N-methyllysine resonances. The spin lattice relaxation time and nuclear Overhauser enhancement effect for theN, N’-dimethyl-Lys-41resonance indicate that the e N H 2 group of this residue is in a relatively restricted environment (37). The pH titration curve of this resonance has two inflections, with pK1 at 9.0 and pK2 at 5.7. Based upon a comparison of the effects of various substrate analogues on pK2 and the pK, values of the active-site histidine residues it was suggested that pK2 arises by a perturbation from His- 12, possibly via a conformational transition of the protein. Markley (316) postulated two slow conformational transitions of the enzyme, one involving His-48, as described previously, and a second with a midpoint at pH 5.6, which might be due to titration of His-12. Also, Riiterjans and Witzel (77) postulated that an electrostatic interaction between the e N H 2 group of Lys-41 and the active-site histidine residues could explain the strong dependence of their pK, values on ionic strength. 6 . Inntemctions wirli Substrate Analogs a. Mononucleotide Phosphates. The effects of nucleotide monophosphates on the histidine C-2 proton resonances have been described by a number of authors (42, 72, 77, 294, 300, 306, 307). The C-2 proton resonances of the active-site histidine residues 12 and 119 shift downfield on binding the pyrimidine mononucleotides 2’- and 3’-CMP; the major shift is that of H3 of His-12. The shift seen on binding of 2’-CMP or 3’-CMP by the C-2 proton resonance of His-12 is beyond the position of a fully protonated histidine, resembles that seen on formation of an imidazoliumphosphate complex (300, 326), and suggests direct contact between the imidazolium group of His-12 and the phosphate of 2’-CMP or 3’-CMP. When 5’-CMP or phosphate binds to RNase, this extensive shift is not observed. The effects on the C-2 proton resonance of His-119 are similar for all three mononucleotides. 326. Cohen, J . S. (1968). BBRC 33, 476.

12. PANCREATIC RIBONUCLEASES

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The C-2 proton resonance of His-48 is shifted slightly downfield by the binding of nucleotide monophosphates, but not by phosphate (300) or pyrophosphate (3061,whereas that of His-105 is not shifted at all. Binding of pyrmidine nucleotide monophosphates to RNase A also produces an upfield shift of an aromatic resonance (294, 300, 306). Four resonances designated F1 through F4, which correspond to the three phenylalanine residues of RNase A, are shown in the high resolution proton NMR spectrum reported by Lenstra et al. (80) and presented in Fig. 3. All four of these resonances are affected upon binding of the active-site inhibitors 2'-UMP or 2'-CMP (Fig. 5). Resonance F3 is most strongly affected and is tentatively assigned to Phe-120 (80,300)since this residue is most directly involved in binding active-site inhibitors (21, 73). A similar shift in the I3C NMR resonance assigned to Phe-120 was also reported by Santoro et al. (74).A resonance, designated I (Fig. 5 ) , which originates from the enzyme-nucleotide complex, titrates with a small shift between pH 4 and 5 and has been assigned to Phe-120 (80,294,300).There appears to be no shift in resonances due to Tyr-25 [contrary to the suggestion by Markley (316) and Antonov et af. (72)1, since the Y3 doublet of Tyr-25 was not affected (80.).The resonances assigned to His-105, and tyrosine residues 25, 73, 76, 92, and 115 do not shift on the binding of either 2'-UMP or 2'-CMP (80). By contrast, the purine mononucleotides 2'- and 3'-AMP have no effect on the C-2 proton resonances of either His-48 or His-105 (306). No downfield shift of the C-2 proton resonance of His-12 is seen on binding of 5'-AMP. Moreover, 2'-, 3', and 5'-AMP are all without effect on the Phe-120 resonance (306, 307). 6. Histidine p K , Changes. The effects of various active-site inhibitors on the pK, values of the histidine residues of RNase A and some of its derivatives are summarized in Table VIII. The various mononucleotide phosphates are not equivalent in their effects, especially with regard to the pK, of His-12. Meadows et al. (300) suggest that the magnitude of the effect on the pK, of His-12 reflects the order of the binding constants, 2'-CMP > 3'-CMP > 5'-CMP. The effect on the pK, of His-12 cannot depend exclusively on the geometry of the phosphate esters, since 2'-UMP and 3'-UMP have much less influence on the pK, of His-12 (305, 306) than do 2'-CMP and 3'-CMP, and do not cause the major downfield shift of its C-2 proton resonances seen with these cytidine mononucleotides (42, 306). Also 2'-UMP and 2'-CMP do not equally affect the resonances F1, F2, and F3 (80). Only small effects on the pK, of either active-site histidine are seen with 2'- and 3'-AMP (306). By contrast, 5'-AMP significantly raises the pK, values of both active-site histidine residues (306,307).

378

PETER BLACKBURN AND STANFORD MOORE 3 mM RNae A, pH 7 1

I

7

65

C-l'H

C-5H

6

6 (ppm) FIG. 5 . Aromatic regions of the proton NMR spectra of RNase A in the presence of active site inhibitors. The dashed lines indicate the positions of the F resonances at pH 7.1 without inhibitor (see Fig. 3). The C resonances are from the inhibitor. From Lenstra e l d. (80).

379

12. PANCREATIC RIBONUCLEASES TABLE VIII T H EEFFECTO F N U C L E O T I D B EI N D I NOGN T H E pK, RIRONUCLEASE A

OF

H I S T I D I NRESIDUES E OF

Histidine residue p K , Nucleotide

His-I2

His-119

His-48

His-105

Native enzyme + 2'-CMP + 3'-CMP + 5'-CMP + Z'deoxy-3'CMP + 2'-UMP + 3'-UMP + 2'-AMP + 3'-AMP + 5'-AMP + Phosphate + UpcA + 2'-FdUpA + 2'-FdUp(Me) + 2'-FdUp Des-( 12I - 124) + 3'-CMP S-protein + Phosphate

5.8

6.2 8.0 7.9-8.0 8.0 8.0 7.8 7.8-8.0

6.3-6.6

6.7 6.7 6.7 6.7 6.7 6.7 6.7 6.7 6.7 6.7 6.7

6.4 6.4 6.3 6.6

6.7 6.7 6.7 6.6 6.7 6.7 6.9

8.0 7.4-7.5 7.0 6.5 6.3 6.2-6.4 6.3 6.0 6.3 6.6 5.8

6.6 6.5 7.3 7.1 7.5

-

6.0 7.6 6.9 6.1 6.7 6.7 7.8 6.8 7.4 6.98 7.4

5.6 6.0

Ref

c. Efects on Sirbstrate Resonances. Two new resonances appearing at high field (Fig. 5 ) are identified as the proton resonances of C-1' of the ribose and C-5 of the pyrimidine base (300, 316, 327). The proton resonances C-5-H and C-6-H of the cytidine bases of 2'-, 3'-, and 5'-CMP, all shift downfield similarly on binding to the enzyme. The proton resonances C-1'-H of the ribose rings are also downfield shifted similarly €or 3'- and 5'-CMP, but slightly differently for 2'-CMP (300). Thus all three mononucleotide bases bind similarly to the enzyme. The shift to lower field of the C-6 and C-5 proton resonances is attributed to anisotropy caused by the base-stacking with Phe-120 (77,300,306);a similar shift in the C-6 proton resonance of 2'- and 3'-UMP is also observed (306). The C-2 and C-8 proton resonances of the adenine base of 2'-, 3'-, and 5'-AMP are all shifted slightly upfield on binding to the enzyme and may arise in part from a stacking interaction with His-1 19 (306, 307). 327. Gorenstein, D. G., and Wyrwicz, A. (1974). Biorlwmisrr~13, 3828.

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PETER BLACKBURN A N D STANFORD MOORE

d. Interactions with the Phosphate Group. While 31Pchemical shifts are a function of the geometry of the phosphate ester (328, 329), they are relatively insensitive to the nature of the group bonded to the phosphate oxygen (328). Gorenstein et a/. (328. 330) have reported the pH dependence of the 31Presonance of pyrimidine nucleotides, 2'(3')- and 5'-CMP, and 3'-UMP, both free in solution and when bound to RNase A. Two ionizations are observed for the complex, found also by Haar et ul. (331), with pK, of 4.0-5.5 and pK2 of 5.9-6.7. The pK1 is associated with ionization of the monoanionic inhibitor, and pK2 with ionization of the protonated His-12, which hydrogen bonds to the phosphate. Gorenstein et a/. (328) calculated the microscopic pK, values that describe the pHdependent equilibria between the monoanionic and dianionic forms of the phosphonucleotides with the enzyme protonated and unprotonated at His-12. For the unprotonated form of His-12, the microscopic pK, for the ionization of the monoanionic phosphonucleotide is nearly the same as the ionization constant for the phosphonucleotide free in solution, which suggests little interaction between the phosphate and the other protonated groups at the active site, His-119, and Lys-41. The pK, values of the 2'and 3'- (but not the 5'-) monoanionic phosphonucleotides bound to the protonated His-12 enzyme are perturbed by 1.5 to 2.0 pH units, a result consistent with the stabilization afforded by hydrogen bonding between the 2'- and 3'-phosphates and His-12 (328). Binding of 3'-CMP to RNase A decreases the spin-lattice relaxation time, T I , of the C-2 proton resonances of His-I2 by 25%, and that of H i s 4 by lo%, but is without effect on those of His-119 or His-105 (332), and further demonstrates the close association between His-12 and this mononucleotide phosphate. The decreased relaxation times, chemical shift changes, and titration behavior of the C-2 proton resonances of His- 12 and His-48 suggest local conformational changes upon binding of active-site inhibitors. e . Ititercrctiuns witlz Dinucleotide Substrate Analogs. The mononucleotide phosphates, around neutral pH, bind in the dianionic state, whereas the phosphodiester substrate is monoanionic and would be expected to have less effect on the pK, values of the active-site histidine residues as a result of charge interactions. Griffin et a / . (305)studied the binding of UpcA, an analog of UpA in which a methylene group re328. 329. 330. 331. 332.

Gorenstein, D. G., Wyrwicz, A. M., and Bode, J. (1976). JACS 98, 2308. Blackburn, G. M . , Cohen, J. S., and Weatherall, I . (1971). Tetrahedron 27, 2903. Gorenstein, D. G., and Wyrwicz, A. (1973). BBRC 54, 976. Haar, W., Thompson, J. C., Maurer, W., and Riiterjans, H. (1973). EJB 40, 259. Benz, F. W., Roberts, G . C. K., Feeny, J., and Ison, R . R. (1972). BBA 278, 233.

12. PANCREATIC RIBONUCLEASES

38 1

places the 5’-oxygen of the adenosine nucleoside (334, and is not cleaved by the enzyme. For the binding of this inhibitor, Griffin et al. (305) found that the pK, values of His-12 and His-119 were not significantly altered. Moreover, they found, by NMR studies (305, 329), little effect on the ionization or conformation of the phosphonate group, which these authors suggest indicates little direct interaction between the phosphonate group and the active-site histidine residues in the RNase A-UpcA complex (305). Antonov rt a/. (72) studied the binding of a nonhydrolyzable analog of UpA to RNase A, 2’-deoxy-2’-fluorouridyl-3‘-p-5’-adenosine (2’-FdUpA). The binding of 2‘-FdUpA, 2’-FdUp methyl ester and 2’-FdUp were studied by proton NMR and 31PNMR. The position of the purine and pyrimidine bases in complex with RNase A were identified from the chemical shifts of the C-6 and C-5 proton resonances of the pyrimidine base, the C-1’ proton resonance of the uridylribose, the C-2 and C-8 proton resonances of the purine base, and the C-1‘ proton resonance of the adenosylribose. The results indicated that the positions of the pyrimidine bases of 2’-FdUpA and 2’-FdUp methyl ester and 2’-FdUp (72) were identical to those previously observed for pyrimidine mononucleotides (300, 306). Binding of the purine base of 2’-FdUpA (72) was somewhat different from that observed for 5’-AMP (306). The chemical shifts of the proton resonances of 2’-FdUpA in complex with RNase A and when free in solution were compared. Measurements were made of the nuclear Overhauser effect for the C-I’ proton resonances of the ribose moieties and those of the C-6 (pyrimidine) and C-8 (purine) proton resonances to determine the glycoside torsion angles in both fragments of the dinucleoside monophosphate when bound to RNase A. The results indicated that the dinucleotide monophospate is bound at the active site in an extended conformation with both nucleotides in the anti conformation. The effects of these fluorine nucleotide analogs on the pK, values of the histidine residues of RNase A are shown in Table VIII. As expected, binding of the monoanionic dinucleotide phosphate and the methyl ester have less of an influence on the pK, values of the active-site histidine residues. A comparison of the effect of 3‘-UMP on the pK, of His-12 with that of 2’-FdUp suggests that the fluorine atom at the 2‘ position of the ribose influences the pK, of His-12, increasing it by about 1 pH unit, and demonstrates the close proximity expected between the 2’-OH of the ribose in UpA and His-12. If the effect of the 2’-fluorine is taken into account, binding of the 333. Jones, G . H . , Albrecht, H. P., Damodoran, N . P., and Moffatt, J . G. (1970). JACS 92, 5510.

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PETER BLACKBURN AND STANFORD MOORE

substrate UpA would not be predicted to affect the pKa of His-12 (72), which agrees well with the results for UpcA reported by Griffin et al. (305). C. OPTICAL PROPERTIES 1. UV Absorption Spectra

Spectrophotometric titrations to study the PKa values of tyrosine residues in RNase are facilitated by the fact that the bovine enzyme and all of the other mammalian pancreatic RNases studied to date (see Section V) contain no tryptophan residues. The absorption near 280 nm is almost entirely due to tyrosine residues. UV spectra have been used to study the degree of exposure of tyrosine side chains in RNase A [reviewed in Ref. (I); also see Ref. (334) for the application of derivative spectra] and the effects of derivatization of available phenolic groups by iodination or nitration (see Section III,A,7). NMR spectroscopy has provided independent data on the pK, values of tyrosine residues (Section IV,B).

2. Circular Dichroism The characteristics of the CD spectra of RNase A in the far- and nearUV regions have been reviewed by Timasheff (335) and by Richards and Wyckoff (I). The near-UV CD spectrum obtained at 25” is characterized by a positive band with a maximum near 240 nm, and a negative band near 275 nm. Strickland and his colleagues (336-338) have shown that the resolution between 250 and 320 nm can be improved by conducting the experiments at 77” K in 1 : 1 water: glycerol solutions (Fig. 6). The nearUV CD spectrum of RNase A shows changes in inflection between 276 and 283 nm that arise from exposed tyrosine residues. The shoulder at 289 nm observed with RNase A (and missing with RNase S, where it may be shifted to about 286 nm) is attributed to a tyrosine residue (probably Tyr-25) that becomes more accessible in RNase S. The shoulders at 268, 261, and 255 nm are attributed to phenylalanine residues. Studies with derivatives of tyrosine have suggested that the major source of tyrosine circular dichroism bands in proteins arises from dipole-dipole coupling between the near-UV transition of tyrosyl side 334. 335. 336. 337. 338.

Brandts, J. F., and Kaplan, L. J. (1973). Biochemistry 12, 2011. Timasheff, S . (1970). “The Enzymes,” 3rd ed., Vol. 2, p. 371. and .,Billups, C. (1970). JACS 92, 2119. Horwitz, J., Strickland, E. €I Honvitz, J., and Strickland, E. H. (1971). JBC 246, 3749. Strickland, E. H. (1974). CRC Crit. Rev. Biochern. 2, 113.

12. PANCREATIC RIBONUCLEASES

383

260 270 280 290 300 Wavelength Inm)

FIG.6. Comparison of the CD spectra of RNase A and RNase S at 77°K. From Horwitz and Strickland (337).

chains with strong far-UV transitions of other nearby moieties. Strong coupling interactions are expected primarily with aromatic amino acids, peptide bonds, and other groups having T orbitals (339-341). Strickland (342) has examined the near-UV CD bands of RNase S on the basis of atomic coordinates for the crystalline enzyme ( 1 ) and theoretical predictions of dipole-dipole interactions within the RNase molecule in order to calculate the rotatory strengths of the individual residues in the near-UV. The interactions between Tyr-73 and Tyr-115 are predicted to provide a major contribution to the negative bands at 276 and 283 nm. Interactions between tyrosine and phenylalanine transitions are considered to contribute little to the optical rotatory properties of the molecule. Despite the numerous peptide groups surrounding some tyrosine side chains, the total rotatory strengths from tyrosyl coupling with peptide bonds are limited by cancellation of contributions having opposite signs. Most of the difference between the calculated and experimental CD spectra comes from the -S-Sbonds of cystine, which contribute to the broad valley (Fig. 6) upon which the finer structure is superimposed. At low temperature (77" K), the bands between 250 and 300 nm in the CD spectra of RNase A (336) and RNase S (337) exhibit similar intensities to those observed at ambient temperature; this result suggests that the 339. 340. 341. 342.

Hooker, T. M . , Jr., and Schellrnan, J. A. (1970). Biopolymers 9, 1319. Chen, A. K . , and Woody, R. W. (1971). JACS 93, 29. Hsu, M.-C., and Woody, R. W. (1971). JACS 93, 3515. Strickland, E. H. (1972). Biochemistry 11, 3465.

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PETER BLACKBURN AND STANFORD MOORE

contributing tyrosine residues have relatively restricted motions (342). Upon cooling solutions of N-acetyl-0-methyl-L-tyrosine ethyl ester from 297" K to 140" K, the negative rotational strength exhibited by this model compound was intensified 10-fold (343). However, Pflumm and Beychok (344) showed that the positive elipticity observed at 240 nm with RNase is temperature-sensitive; a change from 25" to 5" causes an increase in intensity of two- to threefold. Goux and Hooker (345)have extended the calculations of mean residue rotatory strengths for side chain transitions to the positive elipticity at 240 nm. The calculations are based upon the three-dimensional structure ( 1 ) and are fitted to CD data obtained at 27". Their predictions are in general agreement with those of Strickland (342). The contributions from disulfides and the interaction between Tyr-73 and Tyr-115 are considered to be significant for both the positive and the negative bands in the spectrum. Several applications of CD measurements in the course of researches covered in this chapter are included in Sections 111,A; IV,D; V; VI; and VII. 3. Fluorescence Seagle and Cowgill (156, 158) have extended the studies by Cowgill [cf. Ref. (Z)] on the use of fluorescence to study the tyrosine residues in RNase. In the native enzyme, the fluorescence exhibited by RNase A arises from the normally titrating surface tyrosine residues of the molecule. The low quantum yield results from quenching by disulfide bonds and through hydrogen bonding that involves tyrosine residues 25,92, and 97. Seagle and Cowgill (156) reported on the fluorescence characteristics of RNase A modified with tetraintromethane (147)and reduced with bisulfite to produce 3-amino-Tyr residues (148). For one tyrosine residue that was unusually susceptible to such modification, the fluorescence emission maximum was near 395 nm, a high wavelength for 3-amino-Tyrin peptides and on the surface of fibrous proteins; generally A,, is at 350 to 370 nm (158) (see Section III,A,7). Churchich (346) has reported on the fluorescence properties of 4-pyridoxic-5'-phosphate,a competitive inhibitor of the enzyme, with a K1 of M , that is bound firmly to the active site as demonstrated by nanosecond emission anisotropy measurements. Churchich and Wampler (347)reported on the luminescent properties of RNase A at 77" K. The low 343. 344. 345. 346. 347.

Strickland, E. H., Wdchek, M., Horwitz, J., and Billups, C. (1972). JEC 247, 572. Mumm, M. N., and Beychok, S. (1969). JBC 244, 3973. Goux, W. J . , and Hooker, T. M., Jr. (1980). JACS 102, 7080. Churchich, J. E. (1976). Modern Fluorescence Spectroscopy 2, 217. Churchich, J . E., and Wampler, J. (1971). BBA 243, 304.

12. PANCREATIC RIBONUCLEASES

385

phosphorescence yield of RNase A is related to the quenching effect exerted by disulfides on the triplet-excited state of tyrosyl residues. Grandi er al. (348), in studies on the dimeric RNase of bovine seminal plasma, used fluorescence-quenching experiments to observe that the tyrosine residues of the seminal dimer are less exposed to solvent than those of the monomeric species. Iodide and cesium ions and acrylamide [cf., Refs. 349, 3501 have minimal quenching effect with the dimer.

D. THEFOLDING PATHWAY The folding of ribonuclease to its native conformation proceeds in a directed pathway to a final structure of overall minimum free energy (351); the nature of the process has been the subject of several reviews (123,352,353). Interactions within the polypeptide chain must dictate the pathway and the final conformation. Short-range interactions between amino acid residues dominate the determining factors that contribute to the native conformation of a globular protein such as RNase A (354-358). The precise orientation of an amino acid residue in the h a 1 conformation of the protein is further refined via medium-range (359) and long-range interactions with other regions of the polypeptide chain (360). The unfolded protein molecule has been described by Anfinsen and Scheraga (361) as “a fluctuating ensemble of conformations.” By a random mechanism, the folding of a protein molecule would take an inordinately long time (362,363) and can be ruled out (353). The formation of a nucleus, which would then direct subsequent protein folding, has been postulated as the rate-determining step. Both hydrogen-bonded sec348. Grandi,,G., D’Alessio, G., and Fontana, A. (1979). Biochemistry 18, 3413. 349. Eftink, M . R., and Ghiron, C. A. (1976). Biochemistry 15, 672. 350. Eftink, M. R., and Ghiron, C. A. (1977). Biochemistry 16, 5546. 351. Anfinsen, C. B . (1973). Science 181, 223. 352. Baldwin, R. L., and Creighton, T.E . (1980). In “Protein Folding” (R. Jaenicke, ed.), p. 217. Elsevier, Amsterdam. 353. Baldwin, R. L. (1980). In “Protein Folding” (R. Jaenicke, ed.), p. 369. Elsevier, Amsterdam. 354. Kotelchuck, D., and Scheraga, H. A. (1968). PNAS 61, 1163. 355. Kotelchuck, D., and Scheraga, H. A. (1%9). PNAS 62, 14. 356. Finkelstein, A. V., and Ptitsyn, 0. B. (1971). J M E 62, 613. 357. Scheraga, H. A. (1973). Pure Appl. Chem. 36, 1 . 358. Scheraga, H. A. (1978). Pure Appl. Chem. 50, 315. 359. Ponnuswamy, P. K., Warme, P. K . , and Scheraga, H. A. (1973). PNAS 70, 830. 360. Burgess, A. W., and Scheraga, H. A. (1975). PNAS 72, 1221. 361. Anfinsen, C. B., and Scheraga, H. A. (1975). Advan. Protein Chem. 29, 205. 362. Levinthal, C. (1968). J . Chem. Phys. 65, 44. 363. Karplus, M., and Weaver, D. L. (1976). Nature (London) 260, 404.

386

PETER BLACKBURN AND STANFORD MOORE

ondary structures (364, 365) and a cluster of hydrophobic residues (366, 367) have been proposed to form such a nucleus. An alternative view is that folding proceeds via discrete intermediates that determine the pathway and the rate of folding. These alternatives have been discussed by Baldwin (353). 1. Equilibrium Studies

a. Stability of the Native Conformation. The unfolding transition of RNase A was observed originally by spectrophotometric measurements of tyrosine absorption (368-370). Since then, a number of techniques have been applied to the study of this problem. Calorimetric measurements give direct thermodynamic data on the stability of the native conformation, and have demonstrated that the folding transition for RNase A is a highly cooperative process (371). The native protein has been shown to exhibit substantial heat uptake prior to entrance into the cooperative transition zone (371, 372). A pretransition zone preceding unfolding by guanidinium chloride has also been demonstrated (373). These results suggest a loosening of the native structure prior to the cooperative unfolding of the molecule. Laser Raman spectroscopy, which measures the vibrational frequencies of a given class of groups, has been used to study the thermal unfolding of RNase A (374). The results provided evidence for stepwise unfolding of the protein between 32 and 70". NMR spectroscopy has been used to follow the folding transition of RNase A (80, 237, 307, 312). The NMR spectrum of the random coil conformation of a protein represents the sum of the superposition of the spectra of the constitutent amino acids (375,376).The area of a particular resonance is a direct measure of the quantity of the species responsible for that resonance. Thermal and acid unfolding of RNase A produces shifts in 364. Ptitsyn, 0. B., Finkelstein, A. V., and Falk, P.(1979). FEBS Lett. 101, 1. 365. Lim, V. I. (1978). FEES Lett. 89, 10. 366. Wuthrich, K., and Wagner, G . (1978). Trends Biochem. Sci. 3, 227. 367. Matheson, R. R., Jr., and Scheraga, H. A. (1978). Macromolecules 11, 819. 368. Harrington, W. F., and Schellman, J. A. (1956). CR Truv. Lab. Curlsberg, Ser. Chim. 30, 21. 369. Hermans, J . , and Scheraga, H. A. (1961). JACS 83, 3283. 370. Scott, R. A., and Scheraga, H. A. (1%3). JACS 85, 3866. 371. Privalov, P. L., and Khechinashvili, N. N. (1974). J M B 86, 665. 372. Matheson, R. R., Jr., Dugas, H . , and Scheraga, H. A. (1977). BBRC 74, 869. 373. Miller, J. F., and Bolen, D. W. (1978). BBRC 81, 610. 374. Chen, M. C., and Lord, R. C. (1976). Biochemistry 15, 1889. 375. Cohen, J. S . , and Jardetzky, 0. (1968). PNAS 60, 92. 376. McDonald, C. C., and Phillips, W. D. (1969). JACS 91, 1513.

12. PANCREATIC RIBONUCLEASES

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the C-2 proton resonances of His-12, His-119, and His-105 that can be accounted for by the changes in the pK, values and protonation of these residues. However, the C-2 proton resonance of His-48 is an exception; the shift in its C-2 proton resonance indicates that a conformational change occurs in its locale in the pretransition zone. Urea and guanidinium chloride both produce large chemical shifts in the C-2 proton resonances of the histidine residues of RNase A (313). No effect is observed on the resonance of His-105. A small downfield shift of His-119, and a small upfield shift of His-48, are observed throughout changes in concentrations of guanidinium chloride that are insufficient to effect the major unfolding transition of the molecule (up to 1.3 M guanidinium chloride). A large downfield shift for the C-2 proton resonance of His-12 is observed throughout a wide concentration range of the denaturants. Except for that of His-12, the shifts are completed below 1.3 M guanidinium chloride. The areas of the histidine C-2 proton resonances, which give a direct measure of the relative populations of those molecules with an individual histidine in a folded (not necessarily native) state, decrease with increased concentrations of denaturant above 2.0 M guanidinium chloride. The sum of the areas of the individual folded resonances and the resonance that corresponds to the unfolded state remains constant throughout the unfolding transition. The resonance of each histidine residue exhibits a different dependency on denaturant concentration, suggesting that partially folded species must exist during the unfoldirig transition. Chemical trapping experiments have been performed to follow stages in the unfolding of RNase A. One procedure has been to use proteases, which generally hydrolyze the peptide bonds of unfolded proteins much more rapidly than those of the native molecule. RNase A is a particularly good protein for this approach, since its native structure is very resistant to most proteases; as the protein unfolds with increasing temperature, specific peptide bonds are cleaved (377-381). The order in which particular peptide bonds hydrolyze has been taken to indicate the order in which corresponding sections of the molecule unfold. Since the time spans of unfolding steps are commonly of the order of milliseconds (3731, chemical reactions used to trap intermediate stages of unfolding should be fast; photochemical labeling has been used to this 377. Rupley, J . A., and Scheraga, H. A. (1963). Biochemistry 2, 421. 378. Ooi, T., Rupley, J. A., and Scheraga, H. A. (1963). Biochemistry 2, 432. 379. Ooi, T., and Scheraga, H. A. (1964). Biochemistry 3, 648. 380. Klee, W. A. (1967). Biochemistry 6, 3736. 381. Burgess, A. W., Weinstein, L. I., Gabel, D., and Scheraga, H. A. (1975). Biochemistry 14, 197.

388

PETER BLACKBURN AND STANFORD MOORE

end. Information on the thermal unfolding of RNase has been gained using an aryl nitrene generated by flash photolysis (184-186, 188) (see Section III,A,&b); amino acid analysis indicates which types of residues undergo modification. 6 . Contributions of Terminal Segments to the Conformational Stability. The S-protein is folded significantly like RNase A as evidenced by the chemical shifts and titration behavior of the C-2 proton resonances of His-48 and His-105, which are similar to those of RNase A (239). However, the C-2 proton resonance corresponding to histidine residues that are fully exposed to solvent is present to varying degrees in the proton NMR spectra of S-protein at all pH values; there is a pH-dependent equilibrium between folded and unfolded forms of the S-protein. The C-2 proton resonance of His-1 19 is affected by removal of S-peptide, yet this resonance is also shifted significantly by phosphate, suggesting some residual phosphate binding capacity of S-protein (239). Furthermore, it has been shown that S-protein undergoes cooperative thermal unfolding (197, 382), although the melting temperature and thermodynamic parameters (383, 384) indicate that the protein is more labile and exhibits a broader thermal transition than does RNase A. The importance of the six carboxyl terminal residues of RNase to the proper folding of the chain is discussed in Section 111,B,2. c. Properties of the Unfolded Molecule. RNase A, reduced and under nondenaturing conditions, is largely unfolded with all six tyrosine residues exposed to solvent; however, far-UV circular dichroism measurements differ from those typical of a random coil and indicate the presence of some ordered structure (127, 384a). Moreover, fully reduced RNase A retains 0.04% of the activity of native RNase A toward cyclic 2',3'-CMP, with the same substrate concentration-dependence and characteristics of inhibition by 2'-CMP as those of native RNase A (385).This activity is lost after treatment with the sulfhydryl reactant N-ethylmaleimide, indicating that the species responsible for this activity are at most only partially reoxidized (385). NMR studies have shown that, above pH 5.0, thermally unfolded 382. Shenvood, L. M., and Potts, J. T., Jr., (1965). JBC 240, 3799. 383. Tsong, T. Y., H e m , R. P., Wrathall, D. P., and Sturtevant, J. M. (1970). Biochemistry 9, 2666. 384. Rocchi, R., Borin, B., Marchion, F., Moroder, L., Peggion, E., Scoffone, E., Crescenzi, V., and Quadrifoglio, F. (1972). Biochemistry 11, 50. 384a. Takahashi, S., Kontani, T., Yoneda, M., Ooi, T. (1977). J . Biochern. (Tokyo) 82, 1127. 385. Garel, J.-R. (1978).IMB 118, 331.

389

12. PANCREATIC RIBONUCLEASES

RNase, although similar to a random coil (80, 96,31 f -313, 386, 387), still retains some residual structure which may be further unfolded by guanidinium chloride or urea (80,313), or by thermal unfolding below pH 2.0 (3ff , 312). Two of the resonances that shift upon addition of denaturant arise from incompletely exposed His-48 and His-105 (96), and others arise from tyrosine and phenylalanine residues (80). d. Irnrnunochernical Estimation of Protein Conformation. Using an immunochemical approach, Chavez and Scheraga (388) have determined the equilibrium constant, Kconffor the equilibrium between unfolded and folded forms of RNase A and some of its derivatives. Thus, for the equilibrium

where P, and P, are the protein in its random and native forms. The association with anti-native RNase antibodies is defined by Ab

+ P,

Z

AbP,, where K,,,

=

[AbP,I/[Abl[P,I

Thus,

and may be calculated from measurable parameters. At 4" and pH 8.3, for native RNase A the value of Kconf is very large. For reduced RNase A the value of Kconfis 0.06; thus, the antigenic determinants of fully reduced RNase A have about 6% (Kconf/l + &,) of the native conformation of RNase A at equilibrium. This represents a high degree of structure in the fully reduced molecule, since Kconffor a single unstructured determinant is estimated to be to low6(388). Removal of residues 121-124 reduces Kconffor each of the antigenic regions [average value = 0.29, T,,, = 44.5" (252)l; when Phe-120 and His-119 are also removed, Kconffor each region is decreased markedly [average value = 6.8 x lo+, T,,, = 32.5" (252)]. Deletion of the S-peptide to give the Sprotein also reduces Kconf[average value = 0.07, T, = 37.5" (38311. The data clearly indicate the importance of residues in the COOH- and NH2-terminal regions for stability of the native conformation of RNase A. 386. Howarth, 0. W. (1979). BBA 576, 163. 387. Sadler, P. J . , Benz, F. W., and Roberts, G. C. K. (1974). BBA 359, 13. 388. Chavez, L. G . , Jr., and Scheraga, H. A. (1980). Biochemistry 19, 996; ibid., 1005.

390

PETER BLACKBURN AND STANFORD MOORE

2, Kinetic Studies a . Refolding of RNase with Intact Disuljides. RNase, with its native disulfides intact, that has been unfolded by heat or guanidinium chloride, refolds rapidly according to biphasic kinetics when followed by tyrosine absorption or fluorescence emission, and by difference spectra upon 2'-CMP binding. The process has been described by Baldwin and associates (154, 389-392) as one in which Us

k, kz * UF * N k-, k-,

where Us (80%) and UF (20%) represent slow- and fast-folding forms of the unfolded protein, and N represents the native molecule. b. Role of Proline Isomerization. The molecular basis for the difference between Us and UF, first suggested by Brandts et af. (393, appears to result from the cis-trans isomerization of at least two X-Propeptide bonds. RNase A has two cis isomers, at Pro-93 and Pro-114, and two trans isomers, at Pro-42 and Pro-117 (I, 292). Isomerization at the bond at Pro-42 has been considered to be one possibility [see Ref. (399)l. Isomerization at Pro-114 has been observed spectrophotometricaly in RNase nitrated at tyrosine residues 73, 76, and 115, as measured by the effect of isomerization on the ionization of the adjacent 3-nitro-Tyr-115(394,395). The conversion of UF to Us under unfolding conditions and cis-trans proline isomerization are both acid-catalyzed at similar rates and have similar activation enthalpies of the order of 20 kcaUmol(392, 393). The distribution of Us and UF at equilibrium is temperature-dependent (396), and corresponds to the enthalpic difference between cis and trans proline isomers (397) of - 1.4 kcaYmol (396). To account for the fast-folding species UF, Brandts et al. (393) suggested that the four proline residues in this species adopted their native peptide configurations. However, measured under refolding conditions, the activation enthalpy for the Us UF reaction is less than 5 kcdmol (398), as opposed to the 18 to 20 kcaUmol for cis-trans isomerization at

*

389. Garel, J.-R., and Baldwin, R. L. (1973). PNAS 70, 3347. 390. Garel, J.-R., and Baldwin, R. L. (1975). JMB 94, 611. 391. Garel, J.-R., Nall, B. T., and Baldwin, R. L. (1976). PNAS 73, 1853. 392. Hagerman, P. J., and Baldwin, R. L. (1976). Biochemistry 15, 1462. 393. Brandts, J. F., Halvorson, H. R., and Brennan, M. (1975). Biochemistry 14, 4953. 394. Garel, J.-R. (1980). PNAS 77, 795. 395. Garel, J.-R., and SiEert, 0. (1979). BBRC 89, 591. 396. Henkens, R. W., Gerber, A. D., Cooper,M. R . , and Herzog, W. R., Jr. (1980). JBC 255, 7075. 397. Cheng, H. N., and Bovey, F. A. (1977). Biopolymers 16, 1465. 398. Nall, B. T., Garel, J.-R., and Baldwin, R. L. (1978). J M B 118, 317.

12. PANCREATIC RIBONUCLEASES

391

proline (394, 399). Moreover, the rates of refolding do not correlate with the temperature-dependence of the distribution of Us and UF during prefolding conditions (3%); thus isomerization at proline in the unfolded molecule cannot be rate limiting in the folding of Us to N. The rate of isomerization at proline is significantly accelerated during folding, 30-fold at 100 (400).During the refolding of Us to N, the existence of intermediates in the folding pathway prevent the otherwise rapid exchange of amide protons, probably as a result of their engagement in hydrogen-bonded structures (244). Guanidinium chloride (2 M) destabilizes these intermediates and allows the amide protons to exchange. Guanidinium chloride does not affect the rate of conversion of UF to Us (394, 401) and it has no effect on cis-trans isomerization in Lalanyl-L-proline (398);however, it does decrease the rate of isomerization at proline during refolding. Apparently isomerization at proline is accelerated by formation of an intermediate during refolding (400). c . Role of S-Peptide During Refolding. Measurements of protection of amide protons against exchange have indicated that the S-peptide is necessary for the stabilization of early folding intermediates (244). RNase S-protein with its native disulfides intact exists in two unfolded forms, as does RNase A, with a similar distribution of the species, Us (80%) U, (20%) (245). At low temperatures, dissociation of RNase S by pH <2 (226) yields S-protein that remains partially folded (70% at 100) and rapidly recombines with S-peptide (246). RNase A refolds 60 times more rapidly than S-protein; in the presence of S-peptide, RNase S refolds much more rapidly than S-protein. The refolding kinetics of RNase S are concentration-dependent , which suggests that the S-peptide combines with and stabilizes a folding intermediate of the S-protein (245). The refolding kinetics of thermally unfolded RNase A, studied by 360 MH, proton NMR resonances of the C-2 protons of histidine residues, suggest that formation of the a-helical structure in residues 3 to 12 constitutes an early event during refolding of the molecule (243). d . Refolding of RNase with Reduced Disuljides. The refolding of reduced RNase is a complex and slow process. Recovery of the native enzyme by air oxidation requires hours (402, 403). In the presence of a redox mixture of reduced and oxidized glutathione (404&06), reoxidation 399. 400. 401. 402. 403.

Schmid, F. X . , and Baldwin, R. L. (1978). PNAS 75, 4764. Cook, K. H., Schmid, F. X., and Baldwin, R. L. (1979). PNAS 76, 6157. Schmid, F. X., and Baldwin, R. L. (1979). JMB 133, 285. Anfinsen, C. B . , Haber, E., Sela, M., and White, F. H . , Jr. (1961). PNAS 47, 1309. Epstein, C. J . , Goldberger, R . F., Young, D. M., and Anfinsen, C. B. (1962). ABB

Suppl. 1, 223.

404. Saxena, V. P., and Wetlaufer, D. B. (1970). Biochemistry 9, 5015. 405. Ahmed, A . K., Schaffer, S. W., and Wetlaufer, D. B. (1975). JBC 250, 8477.

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PETER BLACKBURN AND STANFORD MOORE

of protein sulfhydryl groups to intramolecular disulfides occurs within minutes near pH 8.0, but with initially incorrect pairing of the protein disulfides (407,408). In a much slower reaction (tllz 90 min by recovery of activity), these bonds then rearrange to the native pairings via glutathione-catalyzed mixed-disulfide exchange reactions (407, 409). In 6 M guanidinium chloride, although the rate of thiol-disulfide exchange is unaltered (410), species with only one or two randomly paired disulfides accumulate along with a significant amount of insoluble material, presumably interchain disulfide cross-linked products (129). e. Trapping of SulJhydryl Intermediates. After trapping sulfhydryl groups by S-alkylation with iodoacetate, during glutathione-facilitated refolding of reduced RNase A, and separation of the S-carboxymethylated products by ion exchange chromatography, Creighton (129,411) identified intermediates with 1,2, 3, and 4 non-native disulfides as well as the native molecule, As the number of disulfide bonds formed increased, the number of non-native disulfide pairings decreased from those statistically possible. Creighton (129) found that cysteine residues 65 and 72 were uniformly more engaged in disulfides throughout refolding than were the other six cysteine residues. Similarly, during Cun-catalyzed air oxidation of reduced RNase, Takahashi and Ooi (412) have found that the native disulfide between cysteine residues 65 and 72 was generated most rapidly. Creighton (413) has also isolated and identified a refolding intermediate (IIIn) with three correctly paired disulfides; the last disulfide between residues Cys-40 and Cys-95 had not formed. This last disulfide is near two proline residues at positions 42 (trans) and 93 (cis); the slower reformation of this disulfide may be related to the isomerization of one or both of these proline residues. The conformation of intermediate IIIn is substantially similar to that of native RNase A; IIIn demonstrates enzymatic activity, and spectral, hydrodynamic, and immunochemical properties similar to those of the native molecule, but its conformation is less stable to denaturation by urea. With the exception of IIIn (asjudged by tyrosine absorption and fluorescence emission, electrophoretic analysis of unfolding in urea gradients (414), and immunochemical cross-reactivity), other intermediates resem-

-

406. 407. 408. 409. 410. 411. 412. 413. 414.

SchaBer, S. W., Ahmed, A. K., and Wetlaufer, D. B. (1975). JBC 250, 8483. Hantgan, R. R., Hammes, G. G . , and Scheraga, H. A. (1974).Biochemistry 13,3421. Creighton, T. E. (1977). J M B 113, 329. Schaffer. S. (1975). Int. J . Peptide Protein Res. 7, 179. Creighton, T. E. (1977). JMB 113, 313. Creighton, T. E. (1978). Progr. Biophys. Mol. B i d . 33, 231. Takahashi, S., and Ooi, T. (1976). Bull. Inst. Chem. Res. Kyoto Univ. 54, 141. Creighton, T. E. (1980). FEES Lett. 118, 283. Creighton, T. E. (1979). J M B 129, 235.

12. PANCREATIC RIBONUCLEASES

393

bled the unfolded species. However, conformational analyses of intermediates trapped by S-carboxymethylation, or with other charged or bulky reagents, may be misleading as a result of charge repulsion and steric effects on the conformations of the intermediates (388,425). Intermediates with two or three correctly paired disulfides, which might be expected to possess native-like structure, would be affected most because the charged groups would be located at precisely the regions predicted to interact with one another. The reshuffling of disulfide bonds in partially regenerated RNase can be arrested without disturbing the disulfide bonds or the conformation of the protein by lowering the pH to 4.0. By this approach, Konishi and Scheraga (416) have studied the temperature-dependence of the initial velocity of cyclic 2',3'-CMP hydrolysis as a probe of the thermodynamic properties of the active site during refolding. Temperature-dependence of tyrosine absorption at 287 nm and changes in optical rotation at 436 nm were also studied as indicators of the polypeptide chain conformation during glutathione-facilitated refolding of reduced RNase A. The thermodynamic parameters AHo (T,) and ASo (T,) and the T, (melting transition temperature) of folding intermediates with enzymatic activity over the range from 0.6 to 100% were identical to those of native RNase. The thermodynamic parameters determined by activity measurements and spectral measurements were essentially identical. Thus, the intermediates formed during refolding of reduced RNase are inactive, and by these techniques their conformations are apparently disordered (416). Proton NMR spectra indicate that the reappearance of the C-2 proton resonances of histidine residues 48, 105, and 119, and the corresponding decrease of the C-2 proton resonance of histidine residues in disordered conformations also parallels the recovery of enzymatic activity during refolding of RNase A (417), again suggesting that the dominant conformations of the intermediates formed during refolding are disordered. f. Immunochemical Approach to RNase Refolding. An alternative approach to seek out intermediates in the folding of reduced RNase has been to use immunochemical methods, based upon the ability of antibodies (toward the native protein) to recognize specifically only native conformation in their antigenic determinants (418). A significant advantage of this approach is its greater sensitivity over physicochemical techniques, permitting low concentrations of folding intermediates to be detected. A 415. Goto, Y.,and Hamaguchi, K. (1979). J . Biochem. (Tokyo) 86, 1433. 416. Konishi, Y., and Scheraga, H. A. (1980). Biochemistry 19, 1308. 417. Konishi, Y., and Scheraga, H. A. (1980).Biochemistry 19, 1316. 418. Sachs, D. H . , Schechter, A. N., Eastlake, A., and Anfinsen, C. B. (1972).P N A S 69, 3790.

394

PETER BLACKBURN AND STANFORD MOORE

disadvantage is that only events on a relatively long time scale can be observed. RNase A has at least four antigenic sites, possibly more, only three of which may be occupied at any one time as a result of competition between two of the sites (279). Subfractions of anti-native RNase A were purified b> affinity chromatography on RNase A-CNBr fragments comprised of residues 1-13, 31-79, and 80-124 that had been coupled to Sepharose (419). The ability of refolded RNase A to compete 100% with lZ5I-labeled native RNase A for binding to anti-native RNase A antibodies and their specific subfractions, and the absence of appreciable crossreactivity with reduced S-carboxymethylated RNase A, made it possible to follow the kinetics of refolding of reduced RNase A (419). Antigenic activity begins to return immediately during air oxidation of reduced RNase A, and returns to 100% after 240 min. Antigenic activity appears first in residues 80-124, followed by residues 1-13, and, finally, residues 31-79. Enzymatic activity reappears after 30 min and correlates with the folding of residues 3 1-79, which therefore probably represents a late event during folding. The thermal stability of these antigenic regions follows the order 1-13 < 31-79 < 80-124 (419). Matheson and Scheraga (367) have proposed that a segment of a polypeptide chain that can fold into a pocket that maximizes hydrophobic interactions forms the primary nucleation site for folding of a protein. They suggest that residues 106 to 118 form the primary nucleation site for RNase A(367). Upon air oxidation of reduced des-( 121-124)-RNase A, randomly paired disulfides are formed (142). RNase S-protein, on the other hand, possesses a limited capacity (20 to 30%) to fold to the native conformation by air oxidation of the reduced S-protein (420,421).Chavez and Scheraga (3881, by the irnmunochemical approach, followed the kinetics of refolding of reduced RNase A, S-protein, and des-( 121- 124)-RNase A, during glutathione-catalyzed thiol-disulfide exchange. It was found that the antigenic site in residues 87- 104 of all three proteins folded before the others. They suggest that nucleation at residues 106- 118 induces early folding of residues 87-104. The fact that S-protein readily folds eliminates residues 1-20 as the primary nucleation site. The partial folding of des(121-124)-RNase A indicates that residues 121-124 are not in the nucleation site; otherwise segment 87-104 would not have folded. Residues 121-124 are required to stabilize later stages of the folding process (388). 419. Chavez, L. G., Jr., and Scheraga, H. A. (1977). Biochemistry 16, 1849. 420. Haber, E., and Anfinsen, C. B. (l%l). JBC 236, 422. 421. Kato, I., and Anfinsen, C. B. (1969). JBC 244, 1004.

12. PANCREATIC RIBONUCLEASES

395

Burgess and Scheraga (422) suggested that segments of RNase that unfold only at high temperatures should provide nucleation sites for the folding of the molecule. Six overlapping stages in the thermal unfolding/ folding process were identified (422), modified by Matheson and Scheraga (186), and later modified by Chavez and Scheraga (388). Some of the observations that have helped to elucidate the thermal unfolding of RNase A, are summarized in Table IX (including Refs. 423-426). The process is first localized to the outermost residues and continues in a series of overlapping stages (186,388,422). In stage I (15-3Y), the side chain of Tyr-92 unfolds, and other localized changes involving Met-13 and Ala-19 and/or Ala-20 occur; in stage I1 (30-4Y), residues 13-25 unfold, and changes occur in the exposure of residues 1-12 to solvent; in stage 111 (40-50"), residues 27-34 and 75-80 unfold; in stage IV (50-60°), residues 51-60 and 121-124 unfold; in stage V (55-65"), residues 1-12, 35-50, and 62-74 unfold; and in stage VI (60-70"), residues 81-102 and 104-120 unfold (186, 388). The outer shell of amino acids, mostly polar residues, unfolds in a broad transition starting at 35-40" and is complete by 70"; the hydrophobic core unfolds in a sharper transition between 60 and 70". The overall transition temperature at pH 5.0 is about 56 to 60". The unfolded form of RNase at 78" still has some residual structure (80, 186, 312, 386, 427).

The folding pathway of reduced RNase is determined primarily by short-range interactions in the protein chain. The formation of disulfide bonds is not considered by Scheraga (123) to influence the folding pathway, but is thought to stabilize the final folded form (361) by reducing the entropy of the unfolded molecule [cf. Refs. 428, 4291. The folding pathway, as proposed by Chavez and Scheraga (388), is as follows: Nucleation by the hydrophobic core of residues 106-1 18, which may include cis-trans proline isomerization, and includes folding of the antigenic determinant in residues 87- 104. Folding continues as residues 63-75 reduce the exposure of the primary nucleation site to solvent. The ,&bend at residues 66-68 brings the chain around as residues 40-48 fold into place, along with residues 63-75 and probably residues 1-12, possibly with stabilizing interactions with the COOH-terminal residues 121422. 423. 424. 425. 426. 427. 428. 429.

Burgess, A. W., and Scheraga, H. A. (1975). J . Theoret. Biol. 53, 403. Li, L.-K., Riehm, J. P., and Scheraga, H. A. (1966). Biochemistry 5, 2043. Bigelow, C. C . , and Sonenberg, M. (1962). Biochemistry 1, 197. Bigelow, C. C. (l%l). JBC 236, 1706. Zaborsky, 0. R., and Mfiman, G. E. (1973). BBA 271, 274. Roberts, G . C. K., and Benz, E W. (1973). Ann. N . Y. Acad. Sci. 221, 130. Lapanje, S ., and Rupley, J. A. (1973). Biochemistry 12, 2370. Johnson, R. E., Adams, P., and Rupley, J. A. (1978). Biochemistry 17, 1479.

396

PETER BLACKBURN AND STANFORD MOORE TABLE IX EVENTS IN

THE

THERMAL UNFOLDING OF RIBONUCLEASE A

Events Stage I 15-35" Tyr-92 side chain unfolds and normalizes Localized changes involve Met-13, Ala-19/20 Stage I1 30-45" Tyr-25-Cys-26 bond becomes accessible Normalization of Tyr-25 Conformational change involves His-48 and Tyr-25 with Asp-14 Disruption of Met-13, Val-47 hydrophobic contact His-12 environment begins to change Stage 111 40-50' Lys-31-Ser-32, Arg-33-Asn-34bonds become accessible Met-79-Ser-80, Tyr-76-Ser-77 bonds become accessible NHpterminal and COOH-terminal sections remain intact Stage IV 50-60" Phe-120-Asp-121accessible at low pH as Val-124 is protonated Ser-123-Val-124 bond becomes accessible a-Helix of residues 51-60 unwinds Stage V 55-65' a-Helix of residues 3-12 unwinds Lys-1 exposed Lys-37-ASP-38, Arg-39-CYS-40, Lys-66-Asn-67 Tyr-73-Gln-74 Stage VI 60-70" Tyr-97 still not normalized L ys-9 1-Or-92

Ile-106-Val-118 nucleation site retained

Evidence

Ref.

Spectrophotornetric titration Photochemical labeling

(370. 423, 425)

a-Chymotrypsin Spectrophotometric titration NMR

(377, 379, 380) (370,423, 425) (312, 426, 427)

Photochemical labeling

(186)

NMR

(312)

Trypsin

(378,.380)

a-Chymotrypsin

(377)

Resistance to exopeptidases

(380,381)

Pepsin

(251

Immobilized carboxypeptidase A Optical rotation

(381 )

NMR Spin labeling; aminopeptidase Stable to trypsin

(375) (372, 380)

(186)

(422

(378)

Stable to a-chymotrypsin

(378)

Spectrophotometric titration Stable to trypsin Theory; antibodies

(425 1

(378) (367, 388)

12. PANCREATIC RIBONUCLEASES

397

124. These events bury the hydrophobic core of the molecule; the remaining residues then pack together in order to decrease the overall free energy of the protein molecule. 3 . Folding in Vivo

Bovine pancreatic RNase is synthesized in vivo with a 25-amino acid extension of its amino terminus (275), the signal peptide (273). This presecretory form of the protein is reported to fold via thiol-disulfide exchange to an enzymically active species. However,this is not always the case; folding of the presecretory proinsulin molecule is inhibited by its signal peptide (430). The folding of presecretory RNase A to a native-like structure is in keeping with observations that the S-peptide segment of the protein is less important for stability and folding in v i m . The observation that the last four residues of the molecule, residues 121-124, are important for maintaining the overall conformational stability of the protein and the inability of reduced des-(121-124)-RNase A to refold, indicates that, at least for RNase A, folding of the nascent protein chain during biosynthesis (431 433) cannot direct the final native conformation. A microsomal enzyme has been described (434, 435) that will catalyze the rearrangement of the disulfide bonds of a protein and facilitate refolding in vitro (436-438). The role of this enzyme in vivo is not clear. The kinetics of refolding for a small protein like RNase A via glutathionecatalyzed mixed-disulfide exchange may be adequate to account for its folding in vivo. Some of the aspects of protein folding in vivo that are pertinent to RNase A have been discussed by Baldwin and Creighton (352).

V.

Species Variations

In the course of studies on pancreatic RNases from a wide variety of mammalian species, Beintema and his colleagues measured the concen430. 797 1. 431. 432. 433. 434. 435. 436. 437. 438.

Lomedico, P. T., Chan, J. S., Steiner, D. F., and Saunders, G . F. (1977). JBC 252, Chantrenne, H.(l%l). “Biosynthesis of Proteins,” p. 122. Pergamon, Oxford. Phillips, D. C. (1967). P N A S 57, 484. DeCoen, J. L. (1970). J M B 49, 405. Goldberger, R. F., Epstein, C. J., and Anfinsen, C. B. (1963). JEC 238, 628. Venetianer, P., and Straub, F. B. (1963). EEA 67, 166. Givol, D., Goldberger, R. F., and Anfinsen, C. B . (1%4). JBC 239, PC3114. Venetianer, P., and Straub, F. B. (1964). BBA 89, 189. Fuchs, S., De Lorenzo, F., and Anfinsen, C. B. (1967). JEC 242, 398.

398

PETER BLACKBURN AND STANFORD MOORE

tration of the enzyme in about one hundred species of mammals (4394 4 0 ~ )The . data extend those of Barnard (441, 442), and show that the concentration in pg/g of tissue ranges from about 0.5 in dog and 5 in man to 1000 in cow and 2000 in eland. The generally high concentrations found in ruminants led Barnard to conclude that the enzyme’s special role in that instance is in the digestion of bacterial RNA. I N AMINO ACIDSEQUENCE A. VARIATIONS

Beintema and associates have conducted sequence determinations on the purified RNases from a wide variety of species. Taken together with the earlier sequence data, the results summarized in Table X (see Refs. 4 4 3 4 6 5 ) demonstrate the variable and conserved regions of the sequences for thirty-eight homologous RNases. The bovine seminal enzyme (see 439. Zendzian, E. N., and Bamard, E. A. (1967). ABB 122, 699. 440. Gaastra, W. (1975). Ph.D. Thesis, Univ. of Groningen, The Netherlands. 440a. Beintema, J. J., Scheffer, A. J., van Dijk, H., Welling, G. W., and Zwiers, H. (1973). Nature New Biol. 241, 76. 441. Barnard, E. A. (1969). Nature (London) 221, 340. 442. Bamard, E. A. (1969). Annu. Rev. Eiochem. 38,677. 443. Beintema, J. J., and Gruber, M. (1973). BBA 310, 161. 444. Beintema, J. J., and Gruber, M. (1967). BEA 147, 612. 445. Lenstra, J. A., and Beintema, J. J. (1979). EJB 98, 399. 446. Jekel, P. A , , Sips, H. J., Lenstra, J. A., and Beintema, J. J. (1979). Biochirnie 61, 827. 447. van Dijk, H., Sloots, B., van den Berg, A., Gaastra, W., and Beintema, J. J. (1976). t i i t . J . Peptide Protein Res. 8, 305. 448. van den Berg, A., and Beintema, J. J. (1975). Nature (London) 253, 207. 449. van den Berg, A., van den Hende-Timmer, L., Hofsteenge, J., Gaastra, W., and Beintema, J. J. (1977). EJB 75, 91. 450. van den Berg, A., van den Hende-Timmer, L., and Beintema, J. J. (1976).BBA 453, 400. 451. Emmens, M., Welling, G. W., and Beintema, J. J. (1976). EJ 157, 317. 452. Jackson, R. L., and Hirs, C. H. W. (1970). JBC 245, 637. 453. Phelan, J. J., and Hirs, C. H. W. (1970). JEC 245, 654. 454. Welling, G. W., Groen, G., and Beintema, J. J. (1975). BJ 147, 505. 455. Welling, G. W., Mulder, H., and Beintema, J. J. (1976). Biochem. Gener. 14, 309. 456. Beintema, J. J. (1980). BEA 621, 89. 457. Suzuki, H., Greco, L., Parente, A., Farina, B., La Montagna, R., and Leone, E. (1976).In “Atlas of Protein Sequence and Structure” (M. 0. Dayhoff, ed.), Vol. 5, Suppl. 2, p. 93. National Biomedical Research Foundation, Washington, D. C. 458. Russchen, F., De Vrieze, G., Gaastra, W., and Beintema, J. J. (1976).BBA 427,719. 459. Groen, G., Welling, G. W., and Beintema, J. J. (1975). FEBS Lett. 60, 300. 459a. Kuper, H., and Beintema, J. J. (1976). BEA 446, 337. 460. Welling, G. W., Scheffer, A. J., and Beintema, J. J. (1974). FEBS Lett. 41, 58. 461. Kobayashi, R., and Hirs, C. H. W. (1973). JBC 248, 7833. 461a. Beintema, J. J., Gaastra, W., and Munniksma, J. (1979). J. Mol. Evol. 13, 305.

12. PANCREATIC RIBONUCLEASES

399

Section VI) is included in the comparison. Residues essential for maintaining the correct secondary and tertiary structure of the protein (including the four -S-Sbridges) and residues with essential roles for binding of substrate and for catalytic function should be expected to remain constant throughout evolution; only nonessential residues should occupy variable domains. Among the pancreatic RNases studied, a number have been found to exhibit sequence heterogeneity (466). A gene duplication has occurred in the guinea pig where two forms of the enzyme RNase A and B are found. They differ from each other at 3 1 positions in the sequence, and one of the forms, RNase B, exhibits heterogeneity at position 64 where the major species has Leu and a minor species has Pro (448, 449). The enzyme in bovine seminal plasma is considered to arise through gene duplication (466); the seminal enzyme is a dimer of identical subunits cross-linked by two adjacent disulfide bonds at positions 31 and 32 (Section VI). The sequence of the subunit is homologous with that of pancreatic RNase A (457).

Allelic polymorphism has been demonstrated in the dromedary pancreatic RNase, where Gln or Lys is found in the ratio 3 : 1 at position 103; the pancreatic RNase of the bactrian species has the same sequence with Gln at this position (454, 455). In roe deer, either Ile or Ala is found at position 64 in the ratio 7 : 3 (464,465). Sequence heterogeneities have also been found in the pancreatic RNase from chinchilla, where either Gly or Asp is found at position 32 (448), in porcupine RNase, where either Gly or Arg is found at position 98 (466), and in hippopotamus RNase where either Gln or Lys is found at position 37 (10). Mutations that have affected the amino terminus are seen in the pancreatic RNase from rat (443, 444), which has three extra residues, and those from kangaroo (9) and wallaby, both of which have a single amino acid deletion at the amino terminus. These events probably represent mutations that have affected cleavage of :he signal peptide from the polypeptide chain during biosynthesis (see Section 111,D). Mutations of the termination codon that have led to chain extensions of the carboxyl-terminus have been found in guinea pig RNase B (448, 449) and in the pancreatic RNase from horse (139), coypu (448), casiragua, 462. Gaastra, W., Groen, G ., Welling, G. W., and Beintema, J. J. (1974). FEBS L e f t . 41, 227. 463. Leijenaar-van den Berg, G., and Beintema, J. J. (1975). FEBS Lett. 56, 101. 464. Zwiers, H., Scheffer, A. J., and Beintema, J. J. (1973). EJB 36, 569. 465. Oosterhuis, S., Welling, G. W., Gaastra, W., and Beintema, J. J. (1977). BBA 490, 523. 466. Beintema, J. J . (1980). Proc. 28th Colloq. Profides B i d . Fluids, p. 139. Pergamon, Oxford.

TABLE X AMINOACIDSEQUENCES Order

Infraorder Superfamily or suborder or family

OF MAMMALIAN PANCREATIC RIBONUCLEASES'

Species (Ref.) 1

10

20

1. ox fBoo DIu11u61. bison IBiJon bi6onl ( 1 5 1 K E T A A A K F E R Q H M D S S T S A A 2. water buffalo swamp type IBubaeu bubaEbI (4561 K E T A A A K F Q R Q H U D S S T S S A 3. river type K E T A A A K F Q R Q H M D S S T S S A 4. eland ITauno-tnagub omizl 14561 K E T A A A K F E R Q H U D S S T S S A 5. nilgai lBo4ePaphud t h a g o c a n ~ u 6 1 (4561 K E T A A A K F E R Q H M D S S T S S A Hippo6. gnu ICclnnwltaetcla tnuninubl (4591 K E S A A A K F E R Q H M D S S T S S A thaqinae 7. topi [ W b C t M kotnigiunl I 4 m l K E S A A A K F E R Q H U D S S T S S A AMbpinae 8. impala [Aepycmob mr'an)wdI K E S A A A K F E R Q H M D S S T S S A 9. Thomson's gazelle (CflzclCn thoMorul (4561 K E S A A A K F Z R Z H M B S S T S S A K E S A A A K F E R Q H U D S S T S S A 10. goat [ C a w hihCUA1, sheep [OUid a h i e ~ l T 6 0 . 4 6 1 1 Caphinae AnZiPocap%idae 11. pronghorn IAtdLPocapta anphicanal (461aI - K E T A A A K F E R Q H I D S N P S S V Gihad6idae 12. giraffe IGina66a c a n c ( q n t d a l i s 1 i 4 T K E S A A A K F E R Q H I D S S T S S V CPnuidae Odoicoiee- 13. reindeer [Rangi@t &zhnnrlw) 14631K E S A A A K F E R Q H U D P S P S S A inae 14. roe deer [ C a p e o h c a ~ . v 1 e o t u l ~ 6 4 , 4 6 5 I K E S A A A K F E R Q H U D P S P S S A 15. moose IACceS dceal 14631 K E S A A A K F E R Q H U D P S A S S I Cehvinae 16. red deer ICehvw &p7LTl 1464,4651 K E S A A A K F E R Q H U D P S T S S A K E S A A A K F E R Q H U D P S M S S A 17. fallow deer (Oana d m l 1 4 6 ' j r Tylopoda Camdidae 18. dromedary [Caneew d f i o ~ d Z & 1 4 ] 1454) S E T A A E K F E R Q H M D S Y S S S S 19. bactrian camel ( C a m d m Iiacthinnud-[4551 S E T A A E K F E R Q H M D S Y S S S S Ancodonta 20. hippopotamus [H4ppOtJOh-u3 mphibiub P I K E T A A E K F Q R Q H M D T S S S L S SUitlL7 21. pig (Sub 4 c h u 6 a l 17,452,5531 K E S P A K K F Q R Q H M D P D S S S S R E s P A u K F Q R Q H M D s G N s P G CetaCU 22. lesser rorqual I&a?aenop& acdoaostnatn) P t A i hdo d a c t y t a 23. horse I E q u cahLepu6I 11391 K E S P A U K F E R Q H M D S C S T S S Rod& Myomqha Mcuidae 24. rat (Rattub n o 4 u e g i c u I m 3 . 4 4 4 1 G E S R E S S A D K F K R Q H M D T E G P S K R E S A A Q K F Q R Q H U D P D G S S I 25. m u s e (ku m u n c u p ~ ~(~4] 4 5 r Chicetidae 26. hamster I#eooc%icetlo nrulafub) (4461 K E S A A H K F E R Q H U D S T V A T S K E T S A Q K F E R Q H U D S T C S S S 27. muskrat IUndatna z i h u t l u c a ) HybthiHybtnicoidea 28. porcupine I H I ( 4 f h i X c h i 4 t a & z l K E S S k M K F E R Q H M D S S C S P S 29. guinea pig 1Cavi.a p o t c e U u 4 I (446,449) A A E S S A M K F E R Q H V D S G C S S S cononpha Cauoidea 30. guinea pig ( C a v i a pozcePPuaI ( I J Z , ~ B] A E S S A M K F Q R Q H M D P E G S P S Ckinckieeoidea 31. chinchilla (CkinckiPPa baovicalZ€iibT(446,450) K E S S A M K F Q R Q H M O S S C S P S K E S S A K K F Q R Q H I D S S C S P S Dotodontoidea 32. casiragua IP.toeckimip guaihnelb S E S S A K K F E R Q 11 M D S R C S P S 33. coypu IMyoroMtua co!it~u51 (446,4501 K E T A A M K F Q R Q H U D S C S S L S 34. two-toed 510th [ C ~ O ~ O P ~h I ~U m & Z l I I01 EdenK E S S A A K F Q R Q H M D S D S S(X X 35. three-toed 510th iB4adypu5 g'Li6eu1 - E T P A E K F Q R Q H M D T E H S T A bllmupiaLia 36. red kangaroo [ M a c t o t w i u d u l I 9 1 - E T A A E K F Q R Z(H U B T)E H S T(A 37. wallaby [lhcrlopw zul(oqGbeu6J - / 9 l b i R i b o n d u e om 4eminaP pPaMma:l 38. ox (804 tarnu61 (4571 K E S A A A K F E R Q H M O S G D S P S Wodac&~fa

Pecona

Bouidne

Bovinae

-

ml

[s]

cr

im

30 40 Y) 60 70 1 . S S S N Y C N Q M M K S R N L T K D R C K P V N T F V H E S L A D V Q A V C S Q K N V A C K N C Q T N C Y Q S Y

2 3 4 5

. . . .

S S S S

S S S S

S S S S

N N N N

Y Y Y Y

C C C C

N N N N

Q Q Q Q

M n M M

M M M M

K K K K

S S S S

R R R R

S N D S

M M M M

T T T T

S S K Q

D D D N

R R R R

C C C C

K K K K

P P P P

V V V V

N N N N

T T T T

F F F F

V V V V

H H H H

E E E E

S S S S

L L L L

A A A A

D D D D

V V V V

Q Q Q Q

A A A A

V V V V

C C C C

S S S S

q Q Q Q

K E K K

N N N N

V V V V

A A A A

C C C C

K K K K

N N N N

G G G G

Q Q Q Q

T T T T

N N N N

C C C C

Y Y Y Y

Q Q Q Q

S S S S

Y Y Y Y

6 . S S S N Y C N Q M M K S R N L T Q D R C K P V N T F V H E P L A D V Q A V C S Q K N V A C K N G Q T N C Y Q S Y 7 . S S S N Y C N Q M M K S R N L T Q D R C K P V N T F V H E S L A D V Q A V C S Q K N V A C K N G Q T N C Y ~ S Y

6 . S S S N Y C N Q n M K S R N L T Q S R C K P V N T F V H E S L A D V Q A V C S Q K N V A C K N G Q T N C Y Q S Y 9 . S S S N Y C N Q M M K S R N L T Q D R C K P V N T F V H E S L A D V Q A V C S Q K N V Q C K N G Q T N C Y Q S Y 1 O . S S S N Y C N Q M M K S R N L T Q D R C K P V N T F V H E S L A D V Q A V C S Q K N V A C K N G Q T N C Y Q S Y 1 1 1 2 1 3 l ~ 1 5

. . . . .

S S S S S

S S S S S

S S S S S

N N N N N

Y C N Q M M Y S B N L T Q C R C K P V N T F V H E S L A D V ~ A V C S Q K N V A C K N G Q T N C Y Q S Y Y C N Q M M T S R N L T Q D R C K P V N T F V H E S L A D V Q A V C S Q K N V A C K N C Q T N C Y Q S Y Y C N Q M M Q S R D L T Q D R C K P V N T F V H E S L A D V Q A V C F Q K N V A C K N G Q S N C Y Q S N Y C N Q M M Q S R N L T Q D R C K P V N T F V H E S L A D V Q A V C F Q K N V I C K N G Q S N C Y Q S N Y C N Q M M Q S R N L T Q D R C K P V N T F V H E S L A D V Q A V C F Q K N V A C K N G Q S N C Y Q S N

1 6 . S S S N Y C N Q M M Q S R K M T Q D R C K P V N T F V H E S L A D V Q A V C F Q K N V A G K N G Q S N C Y Q s N 1 7 . S S S N Y C N Q M M Q S R K M T Q D R C K P V N T F V H E S L A D V Q A V C F Q K N V A C K N G Q S N C Y O S N 1 6 . S N S N Y C N Q M M K R R E M T N G - C K P V N T F I H E S L E D V Q A V C S Q K S V T C K N G Q T N C H Q S S

1 9 . S N S N Y C N Q M M K R R E M T N G - C K P V N T F I H E S L E D V Q A V C S Q K S V T C K N G Q T N C H Q S S Z 2 2 2

O . N D S N Y C N Q M M V R R N M T Q D R C K P V N T F V H ~ S E A D V K A V C S Q K N V T C K N G Q T N C Y Q ( S N 1 . N S S N Y C N L M M S R R N M T Q G R C K P V N T F V H ~ S L A D V Q A V C S Q I N V N C K N G Q T N C Y Q S N 2 . N N P N Y C N Q M M M R R K M T Q G R C K P V N T F V H E S L E D V K A V C S Q K N V L C K N G R T N C Y E S N 3 . N P T N Y C N Q M M K R R N M T Q G - C K P V N T F V H E P L A D V Q A I C L Q K N I T C K N C Q S N C Y Q S S

2 4 . S S P T Y C N Q M M K R Q C M T K G S C K P V N T F V H E P L E D V Q A I C S Q G Q V T C K N G R D N C H K S S 2 2 2 2 2 3 3 3 3

5 C 7 8 9 0 1 2 3

. N S P T Y C N Q M M K R R D M T N C S C K P V N T F V H E P L A D V Q A V C S Q E N V T C K N R K S N C Y K S S S S P T Y C N Q M M K R R N M T Q G Y C K P V N T F V H E S L A D V H A V C S Q E N V A C K N G K S N C Y K S H . S S ( P T Y ) C N Q M M K R R E M T Q G Y C K P V N T F V H E P L A D V Q A V C S ~ E N V T C K N G N S N C Y K S R . S N S N Y C N E M M R R R N M T Q D R C K P V N T F V H E P L A D V R A V C F Q K N V A C K N G Q T N C Y Q S N , S N A N Y C N E M M K K R E M T K D R C K P V N T F V H E P L A E V Q A V C S Q R N V S C K N G Q T N C Y Q S Y . N S S N Y C N V M M I R R N M T Q G R C K P V N T F V H E S L A D V Q A V C F Q K N V L C K N G Q T N C Y Q S Y . T N A N Y C N E M M K G R N M T Q G Y C K P V N T F V H ~ P L A D V Q A V C F Q K N V P C K N G Q S N C Y Q S N . T N P N Y C N A M M K S R N M T Q ~ R C K P V N T F V H E P L A D V Q A V C F Q K N V P C K N G Q S N C Y ~ S T . T N P N Y C N E M M K S R N M T Q C R C K P V N T F V H E P L A D V Q A V C F Q K N V L C K N G Q T N C Y Q S N

3 4 , S S S D Y C N K H M K V R N M T Q E S C K P V N T F V H E S L Q D V Q A V C F Q E N V T C K N G Q Q N C H Q S R 3 S . X X X X X X X ) K M M K S R N M T Q E S C K A V N T F V H ~ P L T D V ~ A V C L Q E N V T C K B G Q Q B C H X X X

3 6 . S S S N Y C N L M M K A R D M T S C R C K P L N T F I H E P K S V V D A V C H Q E N V T C K N G R T N C Y K S N 37. S S S B Y C B L M M)K A R E M T S D R C K P V N T F I H E P K S V V B A V(C 2 Z ) E B ( V T C)K N G Q T N ( C Y)K S ( N 3 6 . S S S N Y C N L M M C C R K M T Q C K C K P V N T F V H E S L A D V K A V C S Q K K V T C K N C Q T N C Y Q S K

(continued)

TABLE X (Coritinuetl) 80 90 M 110 1a0 1 . S T M S I T D C R E T G S S K Y P N C A Y K T T Q A N K H l I V A C E G N P Y V P V H F D A S V

2 . S T M S I T D C R E T G S S K Y P N C A Y K T T Q A N K H l I V A C E G N P Y V P V H F D A S V 3 . S T U S I T D C R E T C S S K Y P N C A Y K T T Q A N K H l l V A C E C N P Y V P V ~ Y D A S V

~ 5 6 7

. S . S T . S T . S T

T M M M

~ S I S I T D S I T D S I T D

T D C C R E T C R E T C R E T

R E T G S S K G S S K G S S K

G Y Y Y

S S P N C P N C P N C

K A A A

Y P N ~ A Y K T T Q A ~ K H I I ~ A ~ E ~ N P Y ~ P ~ H F D A ~ ~ Y T T T Q A K K H I I V A C E G N P Y V P V H F D A S V Y K A T Q A K K H I I V A C E G N P Y V P V H F D A S V Y K T T Q A K K H I I V A C E G N P Y V P V H F D A S V

6 . S T M S I T D C R E T G S S K Y P N C A Y K T T Q A K K H I I V A C E G N P Y V P V H F D A S V

9 . S T M S I T D C R E T C S S K Y P N C A Y K T T Q A Q K H I I V A C E G N P Y V P V H F D A S V 1 1 1 1 ~ 1 1 1 1 1 2 2 2

D 1 2 3

. S T M S I T D C R E T C S S K Y P N C A Y K T T Q A E K H I I V A C E G N P Y V P V H F D A S V . S T M S I T D C R E T C S S K Y P N C A Y K T T Q A K K H I I V A C E C N P Y V P V H Y D A S V . S A M S I T D C R E T C N S K Y P N C A Y Q T T Q A E K H I I V A C E G N P Y V P V H Y D A S V . S A M H I T D C R E T G S S K Y P N C V Y K T T Q A E K H I I V A C E G N P Y V P V H F D A S V . S A M H I T D C R E S G N S K Y P N C V Y K T T Q A E K H I I V A C E C N P Y V P V H F D A S V S . S A M H I T D C R E S C N S D Y P N C V Y K T T Q A E K H I I V A C E G N P Y V P V H F D A S V 6 . S A M H I T D C R E S G N S K Y P N C V Y K A T Q A E K H I I V A C E G N P Y V P V H F D A S V 7 . S A M H I T D C R E S C N S K Y P N C V Y K A T Q A E K H l I V A C E C N P Y V P V H F D A S V 6 . T S M H I T D C R E T G S S K Y P N C A Y K A S N L K K H I I I A C E C N e Y V P V H F D A S V 9 . T S M H I T D C R E T G S S K Y P N C A Y K A S N L Q K H I I I A C E C N P Y V P V H F D A S V D . S T ) M H I T D C R E T C S S K Y P N C A Y K T S Q L Q K H I I V A C E C D P Y V P V H Y D A S V l . S T M H I T D C R Q T C S S K Y P N C A Y K A S Q E Q K H I I V A C E G N P P V P V H F D A SV 2 . S T M H I T D C R Q T G S S K Y P N C A Y K T S Q K E K H I I V A C E G N P Y V P V H F D N S V

2 3 . S S M H I T D C R L T S G S K Y P N C A Y Q T S Q K E R H l I V A C E G N P Y V P V H F D A S V Q T 2 2 2 2

~ 5 . 6 . 7 .

. S S S

S T L R I T D C R L K G S S K Y P N C T Y N T T " P Y V P A L H I T D C H L K G N S K Y P N C D Y K T T Q Y Q K H I I V A C E C N P A L H I T D C R L K G N A K Y P N C D Y Q T S O L Q K Q I I V A C ~ G N A L H I T D C R L K G N S K Y P N C D Y Q ( T S Q L ) Q K Q V I V A C E G S P

V H F D A S V Y V P V H F D A T V P F V P V H F D A S V F V P V H F D A S V

2 8 . S L M H I T D C R V T G S S K Y P D C S Y R M S Q L E R S I V V A C E C S P Y V P V H F D A S V G P S T 2 9 . S S M H I T E C R L T S G S K F P N C S Y R T S Q A Q K S I l V A C E G K P Y V P V H F D N S V 3 D . S R M R I T D C R V T S S S K F P N C S Y R M S Q A Q K S I I V A C E G D P Y V P V H F D A S V E P S T 3 3 3 ~ 3 3

1 2 3 . 5 6

. S N M H I T D C R L T S N S K Y P N C S Y R T S R E N K G I I V A C ~ : G N P Y V P V H F D A S V . S N M H I T D C R L T S N S K F P D C L Y R T S Q E E K S I I V A C E G N P Y V P V H F D A S V A A S A . S N M H I T D C R V T S N S D Y P N C S Y R T S Q E E K S I V V A C E G N P Y V P V H F D A S V A A S A S N M H I T D C R Q T S C S K Y P N C L Y Q T S N M N R H I I I A C E G N P Y V P V H F D A S V E D S T . X X M H I T D C R Q ( T S G S T Y P N C L Y ) K T T N K X X X X X X X X X X X X X V P V H F D A T V . S R L S I T N C R Q T C A S K Y P N C Q Y E T S N L N K Q l l V A C E C Q - Y V P V H F D A Y V 37. S)R L(S 1 T N C)R Q(T G A S B Y P B C 7. Y 2 T S B ) L Q K Q ( I I V A C ) E G Q Y(V P V H F ) D A Y V 3 6 . S T M R I T D C R E T G S S K Y P N C A Y K T T Q V E K H I I V A C C G K P S V P V H F D A S V

-

Classified according to Beintema and Lenstra (468), with sequence data from the cited references. Many residues in peptides have been positioned for the RNases of bovidae and pronghorn by homology with the ox enzyme; a similar procedure has been used with the RNases of deer species (with reference to red deer) and bactrian camel (with reference to dromedary). J . J. Beintema et al., unpublished. Single letter code: A = Ala, B = Asx, C = Cys, D = Asp, E = Glu, F = Phe, G = Gly, H = His, I = Ile, K = Lys, L = Leu, M = Met, N = Asn, P = Pro, Q = Gln, R = A r g , S = Ser, T = Thr, V = Val, W = Trp. X = ?, Y = Tyr, Z = Glx, - = Gap, ( ) = By analogy, from composition, / = )(, back-to-back parentheses.

'

403

12. PANCREATIC RIBONUCLEASES True ruminants Came 1s

ALA (GCX) G U L‘

pig

LYS (AApu)

Horse

MET (AUG)

Lesser rorqual

MET (AUG)

Rat Muskrat

ASP (GAPY) GLU ‘ GLN (CApu)/

Guinea pig

MET (AUG)

Chinchilla

MET (AUG)-

COYPU

LYS (AApu)

Kangaroo

GLU (GApu)

(GAG) \ G L U ( a G )

(GAG)-LYS

(AAG)

GLU (GAG)

F I G .7. Evolutionary history of mutations at position 6 in RNase. Of the ancestral codons, only those that require minimal base changes are given. From Welling et cil. (472).

porcupine (Table X), and two-toed sloth (10). Examples of deletions have not been found at the carboxyl terminus. Havinga and Beintema (10) pointed out that carboxyl terminal additions do not occur frequently, and when they are present the sequences of the extensions are not random. The molecular evolution of mammalian pancreatic RNases has been discussed in detail by Beintema and his colleagues (467, 468). Pancreatic RNase is a rapidly evolving protein by comparison with cytochrome c and several other proteins (469471). The evolution rate of bovid RNases is 2-3 times slower than the average rate observed among the other mammalian RNases (456).On the other hand, rat RNase has evolved at an unusually high rate, 2.5 to 4.3 times as high as the enzyme in related rodent species, and 23 times as high as the average rate in the bovid RNases (445). An example of the evolutionary changes that affect a particular residue is shown in Fig. 7. The residue at position 6 is rather variable, although not all amino acids are permitted at this position. Probably one reason for this restricted variability is the importance of the a-helix in this region of the 467. Beintema, J . J . , Gaastra, W., Lenstra, J. A , , Welling, G . W., and Fitch, W. M. (1977). J . Mol. E t d . 10, 49. 468. Beintema, J. J., and Lenstra, J. A. (1982). / / I “Macromolecular Sequences in Systematics and Evolutionary Biology” (M. Goodman, e d . ) , in press. Plenum, New York. 469. Dickerson, R. E. (1971). J . Mol. Evol. 1, 26. 470. McLaughlin, P. J . , and Dayhoff, M. 0 . (1973). J . Mol. Evol. 2, 99. 471. Smith, E. L. (1970). “The Enzymes,” Vol. 1 , p. 267.

404

PETER BLACKBURN AND STANFORD MOORE

molecule (472). In general, the various amino acid substitutions can be accomodated into the three-dimensional model of RNase S (1, 16) without altering the folding of the backbone. However, mutations that result in the deletion of an amino acid have occurred at residue 39 in horse (472), and dromedary and bactrian camel RNases (454,455). In order to accommodate the deletion of residue 39, the loop that comprises residues 36-39 must be shortened in such a way that the hydrogen bond between the phenolic hydroxyl of Tyr-92 and the carbonyl oxygen of residue 37 is broken, causing Tyr-92 to turn away into the solvent (139). With the exception of Tyr-92, which is conservatively substituted by phenylalanine in guinea pig RNases A and B and in the RNase from casiragua, the surface tyrosine residues have been variously substituted nonconservatively; the two buried tyrosine residues at positions 25 and 97 are invariant. Proline substitution into a polypeptide chain restricts the number of possible chain conformations available at that site because the dihedral angle imposed by proline must be around -70". Among the various pancreatic RNases, proline substitutions occur at ten different sites in addition to the four positions found in bovine RNase A. Oosterhuis et al. (465) have listed the dihedral angle at these ten sites in the X-ray structure of RNase S (16) and find that proline can be accommodated at all of these sites without main chain distortion. Of the four proline residues in bovine RNase, the two at positions 93 and 114 are in the cis configuration; no changes at these positions are found, with the exception of the kangaroo RNase in which residue 114 is deleted (9). No substitutions have been found for the two proline residues at positions 42 and 117, with the exception of Ala at position 42 in the three-toed sloth RNase (10). Further evidence that amino acid substitutions, comprising the variable regions of the pancreatic RNases, have not appreciably affected the three-dimensional structure of the molecule comes from studies on S-peptide and S-protein interactions. Although, the first 24 residues of the sequence contain some of the most variable positions of the RNase molecule, Glu-2, Ala-5, Phe-8, Arg-10, Gln-11, His-12, and Asp-I4 are invariant. Hybrids composed of the S-peptides derived from cow, dromedary, and kangaroo RNases, and a synthetic 17-residue S-peptide corresponding to that sequence of rat RNase, with the S-proteins derived from cow and dromedary camel had very similar properties (190). The dissociation constants for the S-peptides, K , values for cyclic 2',3'-CMP, and maximal velocities showed no significant differences. Immunological 472. Welling, G. W., Leijenaar-vanden Berg, G., van Dijk, B., van den Berg, A., Groen, G., Gaastra, W., Emmens, M., and Beintema, J. J. (1975). Bio Systems 6, 239.

12. PANCREATIC RIBONUCLEASES

405

cross-reactivities with antibodies toward bovine RNase S failed to reveal any difference between the various hybrids. By NMR, the conformation of a hybrid RNase S' composed of bovine S-protein and the synthetic S-peptide corresponding to residues 3-20 of the rat enzyme, was essentially identical to that of bovine RNase S' (473). However, not all pancreatic RNases are cleaved by subtilisin (474,475). The region of the RNase molecule recognized by the protease, residues 16 to 22, is a highly variable part of the sequence; minor changes in the threeLdimensiona1structure of this region due to slightly different conformational preferences of the different amino acids might account for the resistance of some RNases to subtilisin cleavage (475). In particular, the presence of proline residues in this loop is associated with resistance to subtilisin (465). Despite the many amino acid substitutions in the S-peptide segments of the pancreatic RNases, residues that have been shown to be important from binding studies with synthetic S-peptides (see Section II1,B) are invariant, with the exception of the conservative substitutions of Met-13 by Ile in giraffe (462), pronghorn, and casiragua (Table X), and by Val in guinea pig RNase B (448). Most of the variable residues of the pancreatic RNases are hydrophilic and are located in less structured, looped regions on the surface of the molecule. Of the hydrophilic residues that remain constant, most can be ascribed roles. His-12, His-119, and Lys-41 are important for the catalytic process; Lys-7, Lys-66, and Arg-85 are close to the active site and their positively charged side chains may be important for enzymic function. However, Arg-85 has been substituted by histidine in mouse RNase (445). Substitutions of other residues thought to have important roles include Arg-39, which may be responsible for the lower pK, of the e N H 2group of Lys-41 (see Section III,A,3), and has been substituted by tyrosine in chinchilla (448), muskrat (447), and hamster RNase (446),by serine in rat (443,444)and mouse RNase (35),and by lysine in bovine seminal plasma RNase (457). Ser-123 has been substituted by threonine in mouse (445) and three-toed sloth RNase (lo), and by tyrosine in kangaroo (9) and wallaby RNase (Table X); Ser-123 is engaged in a hydrogen bond with uridine-containing substrates (260) (see Section 111,B,2). Other residues on the surface of the molecule that are invariant, or relatively so, include those in the adjacent sequences that involve residues 42-45 and 90-95, both of which contain nonpolar residues and appear to 473. Beintema, J. J . , and Lenstra, J . A . (1980). I n t . J . Pepfide Protein Res. 15, 455. 474. Klee, W. A , , and Streaty, R. A. (1970). JBC 245, 1227. 475. Welling, G . W., Groen, G . , Gabel, D., Gaastra, W., and Beintema, J. J. (1974). FEBS L e f t . 40, 134.

406

PETER BLACKBURN AND STANFORD MOORE

form part of the binding domain for the naturally occurring RNase inhibitor (45) (see Section VII). Residues 65-72 form part of the second basebinding site (21, 73). These relatively nonpolar domains are also among the more thermostable regions of the surface of bovine RNase A (see Table IX, Section IV,D). The eight half-cystine residues are conserved among all of the mammalian pancreatic species examined in detail to date. Weickman et al. ( / I ) have reported on the human pancreatic RNase; amino acid analyses for S- carboxymethylcysteine of the reduced, carboxymethylated protein have indicated 6 half-cystine residues per molecule. This result merits further study. Residues involved in hydrophobic contacts, and residues that shield such contacts, are in general invariant. They appear to be the most important features of the amino acid sequence for the formation of secondary and tertiary structure. Lenstra et a/. (476) applied the predictive methods of Burgess et (11. (477),Chou and Fasman (478,479),and Lim (480, 481 ) to the sequences of some 24 different mammalian pancreatic RNases. The predictions by the method of Lim, based on the relative positions of hydrophobic residues in a-helices and P-sheet structures, gave the best agreement with the X-ray structure of the bovine RNase. All of the residues that according to Lim’s theory are essential for the formation of secondary structure are invariant or conservatively substituted in the ribonucleases tested (476). Moreover, residues 106- 118, proposed by Matheson and Scheraga (367) (see Table IX, Section IV,D) as the nucleation site for the folding of bovine RNase A, form one of the more conserved regions of the sequence. Proton NMR studies have been performed on a number of the mammalian pancreatic RNases (42, 482, 4 8 4 , and pK, values have been determined for their histidine residues (Table XI). His-48 has a higher pK,, in rat (42) and pig (482) RNase than in bovine RNase A. It was suggested for the rat enzyme that this arises because of the influence of the negatively charged glutamate residue at position 16 of the rat enzyme (42); the pig RNase has aspartate at position 16 ( 7 , 452, 453). The high pK, values of 476. Lenstra, J. A . , Hofsteenge, J . , and Beintema, J. J. (1977). J M B 109, 185. 477. Burgess, A . W., Ponnuswamy, P. K . , and Scheraga, H. A. (1974). I s r . J . Biochem. 12, 239. 478. Chou, P. Y., and Fasman, G. D. (1974). Biochemistry 13, 211. 479. Chou, P. Y., and Fasman, G . D. (1974). Biochemistry 13, 222. 480. Lim, V. I. (1974). J M B 88, 857. 481. Lim, V. I. (1974). J M B 88, 873. 482. Wang, F.-F. C., and Hirs, C. H. W. (1979). JBC 254, 1090. 483. Leijenaar-van den Berg, G., Migchelsen, C., and Beintema, J . J. (1974). FEBS L e f t . 48, 218.

407

12. PANCREATlC RIBONUCLEASES TABLE XI pK, VALUES OF H I S T I D I NRESIDUES E IN

THE

PANCREATIC RNASES

p K , values Position

Reindeer"

Ratb

Chinchilla"

Coypu"

12 119 48 73 80 105

6.1 6.5 6.3 >7.0 6.5

6'2 6.6 7.6 6.1 6.3

6.0-6.1"

6.3d

4.9 7.2

5.8 8.0 -

-

Pig' 6.4 6.3 >7.5 >7.0 6.6

From van den Berg and Migchelsen (48.1). From Migchelsen and Beintema (42). ' From Wang and Hirs (482). Unassigned.

His-80 in coypu, chinchilla (42),reindeer (483),and pig RNase (482) arise most probably as the result of a salt bridge with Glu-49 (450). From NMR data, the active-site conformations of the bovine and rat RNAses resemble one another more closely in the presence of pyrimidine mononucleotides. This result suggests evolutionary constraints on the preservation of the active-site structure in the enzyme-substrate complex rather than in the substrate free state (42). Myer et a/. (484) arrived at a similar conclusion, based on the near-UV circular dichroism difference spectra of bovine RNase A and turtle RNase (442,485)in the presence of 2'- and 3'-CMP. B.

VARIATIONS IN

CARBOHYDRATE MOIETIES

Post-translational modifications that attach carbohydrate side chains to pancreatic RNases occur in many species. Beintema rt a/. (486) have tabulated (Table XII) (see refs. 487-490) the points of attachment and the approximate compositions of the carbohydrate side chains found with the enzymes from different species. Four carbohydrate attachment sites have 484. Myer, Y. P., Barnard, E. A., and Pal, P. K. (1979). JBC 254, 137. 485. Barnard, E. A . , Cohen, M. S., Gold, M. H., and Kim, J. (1972). Nrrrrrre (Londun) 246, 395. 486. Beintema, J. J., Gaastra, W., Scheffer, A. J., Welling, G. W. (1976). EJB 63, 441. 487. Plummer, T. H., Jr. (1968).JBC 243, 5961. 488. Becker, R. R., Halbrook, J. L., and Hirs, C. H. W. (1973). JBC 248, 7826. 489. Kabasawa, I . , and Hirs, C. H. W. (1972). JBC 247, 1610. 490. Tsuruo, T., Yamashita, S., Terao, T., and Ukita, T. (1970). BBA 200, 544.

TABLE XI1 APPROXIMATE COMPOSITION

OF T H E

CARBOHYDRATE MOIETIESOF GLYCOSIDATED RIBONUCLEASES" ~~

Ratio of monosaccharide residuesb

Species Cow pancreas

Sheep Pig

Giraffe

Okapi

Component B C D B C

Attachment site 34 34 34 34 34 21 34 76 all 34a 34b 34c 34a 34b 34c

G~UCOSamine 2 4 4 2 5 7- 10 2 8-11

11

4 7 3 2 5

4

Mannose 6 4 3 6 6 3 6 3

7 5 6 5 4 4 19?

Galactose

2 2 2 2-4

Fucose

1 1 -

2 1

3-4 2 3 3

2 3

1

1

2 2 -

1 1 1

1 1

Sialic acid

21

4

Refs.

Moose Horse

Lesser rorqual B COYPU Chinchilla

I I1

Guinea pig

Roe deer Hippopotamus Two-toed sloth Hamster River-type water buffalo Porcupine Casiragua

B

B

34 21 34 62 76 34 34 34 21a 21b 34 34 34 34 34 34 34 34

3

5 6 5 3 2 4 3 2 3 2 3

4 2 Not determined 5

4-6 10- IS 6 7 3 9 4 9

+

5

+

" Adapted from Beintema et a / . (486);galactosamine not present. not found; + , number of residues not determined. Approximate values, J. J. Beintema et d..unpublished.

(463 )

5

4

* - Residue

1

5

+

4 3

1

(450)

(449)

2 -

4 6

1

1

I1

Trace

410

PETER BLACKBURN AND STANFORD MOORE

been found at positions 2 1,34,62, and 76. All occur at asparagine residues in an Asn-X-Thr/Ser sequence, coupled via N- acetylglucosamine in an N-glycosidic linkage (491, 492). The four carbohydrate attachment sites occur at exposed regions of the RNase molecule, removed from the active site, in variant parts of the sequence. The enzymes in even closely related species differ by the presence or absence of carbohydrate (486). Moreover, not all potential glycosylation sites are coupled to carbohydrate and there are notable differences in the complexity of the carbohydrate chains. The carbohydrate side chain of bovine RNase B (66, 493) has been sequenced by Liang ei al. (494) and is Mand (Mancvl

+ 2)0--3

Manal

I

!Mancvl

I

f

!Man01

+

4GlcNAcpl .+ 4GlcNAc

f

Mancvl

The side chain is variously elaborated through the addition of one to three mannose residues joined by an a(1 + 2) linkage. Most of the pancreatic RNases studied have experienced acid conditions at some point during their isolation, with the exceptions of some of the preparations of the proteins from cow (66,493), pig (489,495,496), and sheep (488). Carbohydrate side chains, especially at sialic acid residues, undergo gradual degradation when exposed to acid (496). Sialic acid residues have been found in gl ycopeptides isolated from various pancreatic RNases that have been exposed to acidic conditions; thus their carbohydrate side chains cannot have undergone extensive acid degradation. The function of the carbohydrate chains attached to some of the pancreatic RNases is unknown. Beintema et nl. (486) have suggested that since species with cecal digestion (like pig, horse, guinea pig, chinchilla, and coypu) produce RNases with large carbohydrate chains attached to one or several sites on the surface of the molecule; the carbohydrate perhaps protects the RNase molecule from absorption by the gut. This 491. Eylar, E. H. (1965). J . Tlieorrt. 861. 10, 89. 492. Neuberger, A., and Marshall, R. D. (1%9). “Symposium on Foods-Carbohydrates and Their Roles” (H. W. Schultze, R. F. Chain, and R. W. Wrotstad, eds.), p. 115. Avi, Westport, Connecticut. 493. Plummer, T. H., Jr., and Hirs, C. H. W. (1963). JBC 238, 1396. 494. Liang, C.-J., Yamashita, K., Kobata, A. (1980). J. Biochem. (Tokyo) 88, 51. 495. Reinhold, V. N., Dunne, F. T., Wriston, J. C., Schwartz, M., Sarda, L., and Hirs, C. H. W. (1968). JEC 243,6482. 496. Jackson, R. L., and Hirs, C. H. W. (1970). JBC 245, 624.

12. PANCREATIC RIBONUCLEASES

41 1

woutd then facilitate its transport to the large intestine where it should hydrolyze the RNA from the cecal microflora, analogous to the postulated function of pancreatic RNases in ruminants (441). The effect of the carbohydrate side chains on the properties of the porcine pancreatic RNase molecule has been studied by Wang and Hirs (497). They compared a number of physicochemical properties of the native molecule and of molecules in which the carbohydrate was substantially reduced after digestion with a mixture of exoglycosidases. The size of the carbohydrate side chains had no influence on the rate at which the fully reduced, denatured protein reassumed the native folded structure on reoxidation, nor on the overall conformational stability of the molecule. The results of spectrophotometric titrations at high and low pH indicated that the carbohydrate chains increased the conformational stability of local surface regions associated with tyrosine residues. Circular dichroism measurements and UV absorption-difference spectra suggested that the carbohydrate chains affected the local tertiary structure around at least one tyrosine residue. Based upon the results of Puett (498), who found by circular dichroism measurements that the carbohydrate side chain at Asn-34 of bovine RNase B has no effect on the tertiary structure of the molecule and no influence on the nearby residue Tyr-92, Wang and Hirs (497) tentatively identified Tyr-25 of the porcine RNase as the residue most likely affected, possibly through interactions with the carbohydrate attached to nearby residue Asn-21. The conclusion from the results with pig pancreatic RNase was that no special mechanisms are necessary for correct polypeptide chain folding when glycoproteins are synthesized on membrane-associated ribosomes; this may not be true for all glycoproteins.

VI.

Bovine Seminal Plasma RNase

The ribonuclease of bovine seminal plasma was first described by D’Alessio and Leone (499); the enzyme represents more than 2% of the total protein of the fluid (500).Two components were identified, BS-1 and BS-2, and the major component, BS-1, was purified and shown to have a MW of 29,000 and an isoionic point at pH 10.3 (500, 501). Similar proper497. 498. 499. 500. 153. 501.

Wang, F.-F. C., and Hirs, C. H. W. (1977). JBC 252, 8358. Puett, D. (1973). JBC 248, 3566. D’Alessio, G . , and Leone, E. (1963). BJ 89, 7P. D’Alessio. G . , Floridi, A , , De Prisco, R . , Pignero, A , , and Leone, E. (1972). EJB 26, Floridi, A., and De Prisco, R . (1973). I d . J . Biochern. 22, 1.

412

PETER BLACKBURN AND STANFORD MOORE

ties were reported by Hosakawa and Irie (502)for ribonucleases isolated from bovine seminal vesicles. The early studies demonstrated that while the seminal plasma enzyme had some properties in common with the bovine pancreatic enzyme, the differences in molecular size and charge were prominant (503-506). RNase BS- 1 is a pyrimidine-preferring endoribonuclease that yields 3’-phosphonucleotides via cyclic 2’ ,3‘intermediates (507), as is bovine RNase A (508), but differs from the pancreatic enzyme in its ability to hydrolyze double-stranded RNA (509). Seminal RNase BS-1 has been found to be comprised of two identical subunits (510) with an amino acid sequence (457, 511-513) that is homologous with that of the mammalian pancreatic RNases (see Table XI. There are 23 substitutions in nonessential positions of the chain; all key residues (such as His-12, His-119, and Lys-41) are present, and the four intrachain disulfides (514) are at positions identical with those found in the pancreatic RNases. The two subunits are covalently cross-linked by two readily reduced, adjacent disulfides at positions 31 and 32 of the polypeptide chain (515-517). Selective reduction of the interchain disulfides by dithiothreitol occurs rapidly with 30% dissociation of RNase BS-1 into monomers, and 70% into noncovalently associated dimers (517); complete dissociation to monomers requires mildly denaturing conditions. The refolding of fully reduced, denatured RNase BS-1 is facilitated optimally at pH 8.0 and 30” with a mixture of 3.0 mM reduced glutathione and 0.6 mM oxidized 502. Hosakawa, S. , and hie, M. (1971). J . Biochem. (Tokyo) 69, 683. 503. Forlani, L., Chiancone, E., Vecchini, P., Floridi, A., D’Alessio, G., and Leone, E. (1967). BBA 104, 170. 504. Floridi, A., and DAlessio, G. (1%7). Boll. Soc. Itul. Biol. Sper. 43, 32, 505. Floridi, A. (1968). BBRC 32, 179. 506. D’Alessio, G., Demma, G., Farina, B., Leone, E., and Parente, A. (1970).Boll. Soc. Ital. B i d . Sper. 46, 96. 507. Floridi, A., D’Alessio, G., and Leone, E. (1972). EJB 26, 162. 508. Volkin, E., and Cohn, W. E. (1953). JBC 205, 767. 509. Libonati, M., and Floridi, A. (1969). EJB 8, 81. 510. D’Alessio, G., Parente, A., Guida, C., and Leone, E. (1972). FEBS Lett. 27, 285. 511. D’Alessio, G., Parente, A., Farina, B., La Montagna, R., De Prisco, R., Demma, G. B., and Leone, E. (1972). BBRC 47, 293. 512. Leone, E., Suzuki, H., Greco, L., Parente, A , , Farina, B., and La Montagna, R. (1972). Proc. 8th FEBS. Meet. Amsterdrim, Abstr., p. 359. 513. Suzuki, H., and Greco, L . (1972). Boll. Soc. Itnl. B i d . Sper. 48, 1124. 514. Di Donato, A., and D’Alessio, G. (1979). BBA 579, 303. 515. Malorni, M. C., Di Donato, A., and D’Alessio, G. (1972). Boll. Soc. Ifnl. B i d . Sper. 48, 606. 516. Di Donato, A., and D’Alessio, G. (1973). BBRC 55, 919. 517. D’Alessio, G., Malorni, M. C., and Parente, A. (1975). Biochemistry 14, 1116.

12. PANCREATIC RIBONUCLEASES

413

glutathione (518), in much the same way as for bovine RNase A (405). The major product is the monomeric species with two moles of glutathione per mole of subunit as mixed disulfides at positions 31 and 32 (518).Selective reduction of the interchain disulfides with reduced glutathione yieids bis-S-glutathione-RNase-BS- 1 monomers (5f9 ) that are quite stable and show no tendency to re-form disulfide cross-linked dimers. Selective reduction of the interchain disulfides of RNase BS-1 followed by bis-Salkylation with iodoacetate (to give MCM-BS- 1) ( 5 / 7 ) , iodoacetamide (MCAM-BS-I), and ethylenimine (MAE-BS-l), yields catalytically active subunits (520). Smith and Schaffer (521) showed that these derivatives, after full reduction, will refold to the active species. Bovine RNase A (522-524) and seminal RNase BS-1 (525) aggregate when lyophilized from 30 to 50% acetic acid. The alkylated monomeric species, MCM-, MCAM-, and MAE-BS-1 showed less tendency to aggregate ( 5 , 8, and 11%, respectively) than did RNase A (24%) when similarly lyophilized (520).A structure has been proposed (522-524) for the aggregated dimer of RNase A in which the NH2-terminal segment (the S-peptide region) of one molecule adsorbs on the main chain (the S-protein region) of another, and vice versa. Low resolution X-ray diffraction studies on crystalline RNase BS-1 (526) provide data that indicate that a similar structure is possible for the seminal dimer, but the resolution is not sufficient to eliminate alternative orientations. The two monomeric subunits are disposed in the anti-parallel configuration, with half-Cys-3 I of one linked to residue 32 of the other, and vice versa. The dissociated monomeric subunits of RNase BS-1 adopt a different conformation from that which they have in disulfide cross-linked dimers. The circular dichroism spectra of bis-S-glutathione-BS-1 (518, 519) and MCM-BS- 1 monomers (348) closely resemble that of native bovine pancreatic RNase A, but the monomeric subunits appear to contain more a-helical content and less @-structure than the covalently cross-linked dimer (348). Near-UV circular dichroism measurements, fluorescence and fluorescence-quenching studies, and UV-absorption difference spectra all 518, Smith, G. K . , D'Alessio, G . , and Schaffer, S. W. (1978). Biochemistry 17, 2633. 519. Smith, G. K . , and Schaffer, S. W. (1979). ABB 196, 102. 520. Parente, A . , Albanesi, D . , Garzillo, A . M., and D'Alessio, G . (1977). I r a / . J . Biorhem. 26, 451. 521. Smith, G. K . , and Schaffer, S. W. (1980). ABB 203, 282. 522. Crestfield, A. M., Stein, W. H., and Moore, S . (1962). ABB Suppl. 1, 217. 523. Fruchter, R. G., and Crestfield, A. M. (1965). JBC 240, 3868. 524. Fruchter, R . G., and Crestfield, A . M . (1965). JBC 240, 3875. 525. Libonati, M. (1969). Itol. J . Eiochem. 18, 407. 526. Capasso, S . , Giordano, F., Mattia, C. A , , Mazzarella, L., and Zagari, A. (1979). Go::. Chim. Itci(. 109, 55.

414

PETER BLACKBURN AND STANFORD MOORE

suggest an increased exposure of tyrosine residues upon reduction of the interchain disulfides and dissociation to monomers (348). Of the four tyrosine residues, at positions 25, 73, 92, and 97, only one, presumably Tyr-73, titrates spectrophotometrically with a normal pK, (527). Other evidence of conformational differences between monomeric and dimeric forms of RNase BS-1 subunits is suggested from the differences in the extinction coefficients, E!Fm, of native RNase BS-1, 4.65 (500); MCMBS-1, 4.82 (517); MCAM-BS-1, 5.37; and MAE-BS-1, 5.49 (520). Also, the monomeric species MCM-BS- 1 exhibits susceptibility to digestion with trypsin, whereas RNase BS-1 does not (528), and MCM-BS-1 is less stable to heat, acid, and urea denaturation than pancreatic RNase A and dimeric RNase BS-1 (348). Immunologically, RNase BS-1 dimer demonstrates less reactivity with anti-RNase A serum than does RNase A, while the monomeric species is very similar to RNase A in its interaction with this antiserum (529). Bovine pancreatic RNase A, at physiological ionic strength and pH, has essentially no activity toward double-stranded RNA (530) or poly(A); activity toward the latter is observed at high concentrations of both enzyme and poly(A) (53f). On the other hand,-RNase BS-1 is active under physiological conditions toward double-stranded RNA, and will hydrolyze the polypyrimidine strand of a poly(A). poly(U) complex (509) and the RNA strand of a DNA-RNA hybrid (532). Similarly, aggregates of bovine pancreatic RNase A and its chemically cross-linked dimers (55, 533) act on double-stranded RNA (509,525), the RNA strand of a DNARNA hybrid (534), and measurably on poly(A) (530). Libonati and Floridi (509) considered it unlikely that the two active sites in the dimeric enzymes were important in the activity toward doublestranded substrates and suggested that the determining factor was probably the positive charge density of the enzyme. This hypothesis received support from studies with monomeric derivatives of RNase BS-1 (520, 535) and with species variants of the enzyme that exhibited different 527. Irie, M., and Suito, F. (1975). J . Eiochetn. (Tokyo) 77, 1075. 528. Parente, A., Branno, M., Malorni, M. C.. Welling, G. W., Libonati, M., and D’Alessio, G. (1976). EEA 445, 377. 529. Floridi, A., and Fini, C. (1972). f m l . J . Biochem. 21, 72. 530. Libonati, M. (1971). BEA 228, 440. 531. Beers, R. F., Jr. (1960). JEC 235, 2393. 532. Taniguchi, T., and Libonati, M. (1974). BERC 58, 280. 533. Bartholeyns, J . , and Moore, S. (1974). Science 186, 444. 534. Libonati, M., Sorrentino, S. , Galli, R., La Montagna, R., and Di Donato, A . (1975). BBA 407, 292. 535. Libonati, M . , Malorni, C., Parente, A., and D’Alessio, G. (1975). EBA 402, 83.

12. PANCREATIC RIBONUCLEASES

415

basicities (536);both the net positive charge and the positions of the basic residues are considered to contribute to the effectiveness of the enzyme toward double-stranded substrates. In that connection, Wang and Moore (59) have shown that the highly basic derivative of RNase A cross-linked to octaspermine with dimethyl suberimidate is 115 and 380 times as active as RNase A toward poly(A) 'poly(U) and the hybrid poly(rU) 'poly(dA), respectively, the increased activity being primarily a result of a 100-fold decrease in the K m for these substrates (537). The resistance of double-stranded RNA and poly(A) to hydrolysis by bovine RNase A is only observed at physiological ionic strength. At onetenth the physiological ionic strength, the activity of bovine RNase A is equal to or greater than the activities of RNase BS-1, the more basic whale pancreatic RNase, and chemically cross-linked dimers of bovine RNase A (538, 539). Although the mechanisms are not fully established (540), the effects of ionic strength on the relative activities of the various RNases toward single- and double-stranded substrates reflect changes in the K , of the substrates for each enzyme. For example, the K ifor poly(A) inhibition of bovine RNase A activity toward yeast ribosomal RNA decreases 10-fold with respect to the substrate K , as the ionic strength is decreased from 0.1 M to 0.005 M in Tris-HC1 buffer at pH 7.5 (P. Blackburn and R. Jacoby, unpublished results), making it a relatively stronger competitive inhibitor, whereas at physiological ionic strength the Ki for poly(A) and the K , for RNA are almost identical. The K, of RNase BS-I for yeast RNA increases significantly as the ionic strength is increased above 0.2 M salt (541). Bovine RNase A dimers and RNase BS-1 destabilize the structure of double-stranded DNA (542) much more efficiently than does bovine RNase A (543). When the known interactions of nucleotides with bovine pancreatic RNase are considered with respect to the X-ray structure of the molecule ( I , I 6 , 2 / , 7 3 ) it is clear that at least local strand separation of a double-stranded substrate must occur to accommodate the polynucleotide into the enzyme active site. 536. Libonati, M . , Furia, A , , and Beintema, J . J . (1976). EJB 69, 445. 537. Wang, D. (1979). BBA 568, 488. 538. Palmieri, M., and Libonati, M. (1977). BBA 474, 456. 539. Libonati, M., and Palmieri, M . (1978). BBA 518, 277. 540. Sorrentino, S . , Carsana, A., Furia, A , , Doskofil, J., and Libonati, M. (1980). BBA 609, 40. 541. Floridi, A . , and Fini, C. (1973). ftu1. J . Biocliern. 22, 7 . 542. Libonati, M . , and Beintema, J . J. (1977). Biochem. Soc. TrcrnA. 5, 470. 543. Felsenfeld, G., Sandeen, G., and von Hippel, P. H . (1963). P N A S 50, 644.

416 VII.

A.

PETER BLACKBURN AND STANFORD MOORE

Cytoplasmic RNase Inhibitor

PURIFICATION A N D CHEMICAL PROPERTIES

Nonpancreatic tissues contain small amounts of RNases that in many respects resemble the pancreatic enzyme. These RNases have not been thoroughly characterized, primarily as a result of the minute quantities available for study. Levy and Karpetsky (544) and Maor and Mardiney (545) have reviewed studies on these RNases from human tissue, serum, plasma, and urine in relation to neoplastic and nonneoplastic diseases. Investigations of tissue RNases of the pancreatic type require consideration of the presence of an endogenous RNase inhibitor. Normally, more than 95% of the available RNase activity measured near neutral pH in the postmitochondrial supernatant fraction of mammalian tissues is in a latent form. The RNase, bound by an inhibitor, forms an inactive complex maintained by a 6- to 8-fold molar excess of free inhibitor over the enzyme. The presence in mammalian tissues of this inhibitor of neutral RNase activity was first described in 1952 by Pirotte and Desreux (546). Since that time, most if not all mammalian tissues have been found to contain small amounts (e.g., 1 part in 10,000 on a protein basis) of this RNase inhibitor in the cytoplasm. Many of the early studies on the inhibitor were performed by Roth and his colleagues (547-550) and have been reviewed by Roth (551). The inhibitor is not restricted to mammalian species and has been found in marsupial (552,553),amphibian (553n), and avian livers (552, 554, 55.5) where it exhibits species specificity toward its respective neutral RNase. 544. Levy, C. C., and Karpetsky, T. P. (1981).//1"Enzymes as Drugs'' ( J . S. Holcenberg, ed.), p. 103. Wiley, New York. 545. Maor, D . , and Mardiney, M. R., Jr. (1979). CRC Crit. Rev. Clin. Lab. Sci. 10, 89. 546. Pirotte, M., and Desreux, V. (1952). Bull. Soc. Chim. Belges 61, 167. 547. Roth, J. S. (1956). BBA 21, 34. 548. Roth, J. S. (1958).JBC 231, 1085. 549. Roth, J. S . (1958). JBC 231, 1097. 550. Roth, J. S . (1962). BBA 61, 903. 551. Roth, J. S. (1967). Merhods Cancer Res. 3, 153. 552. Kraft, N., and Shortman, K. (1970). Austr. J . B i d . Sci. 23, 175. 553. Meyer, D. H . , Meyer, W. L . , and Kakulas, B. A. (1976). I n "Recent Advances in Myology" (W. G. Bradley, D. Gardner-Medwin, and J. N. Walton, eds.), p. 277. Excerpta Medica, Amsterdam. 553a. Malicka-Blaskiewicz, C. (1978). Proc. 12th FEES Meet. Dresdeti. Absrr. 0122. 554. Kraus, A. A., and Scholtissek, C. (1974). EJB 48, 345. 555. Dijkstra, J., Touw, J., Halsema, I., Gruber, M., and AB, G. (1978). BBA 521, 363.

12. PANCREATIC RIBONUCLEASES

417

The researches by Roth (547, 548) and Shortman (556, 557) on the RNase inhibitor of rat liver demonstrated that the inhibitor was a heatand acid-labile, sulfhydryl-dependent protein readily inactivated by p-hydroxymercuribenzoate with concomitant activation of the latent RNase. The protein does not inhibit acid lysosomal RNase (557) and is specific for neutral RNases of the pancreatic type. Generally, most purification procedures have used combinations of salt fractionation of the postmicrosomal or postmitochondrial supernatant fractions of isotonic extracts of mammalian tissues, followed by ion exchange chromatography and gel filtration. Using this approach Gribnau et a/. (558) determined that the molecular weight of the rat liver RNase inhibitor was near 50,000; although a 3000-fold purification was achieved, the protein was still impure by SDS gel electrophoresis. This purification was further extended by Gribnau et af. (559) using affinity chromotography on RNase A coupled to carboxymethyl-cellulose. A similar approach was later used by Gagnon and de Lamirande (560). Studies on the properties of the rat liver inhibitor purified by Gribnau el al. (5.58) demonstrated the importance of EDTA and free thiol, especially dithiothreitol, for maintaining the protein in its active form (561). However, the best preparations (559) were contaminated by a potent leucine aminopeptidase that was detrimental to studies on cell-free protein synthesis (562). A number of the properties of the partially purified RNase inhibitor have been described (563-571 ). The inhibitor was observed to be present 556. Shortman, K. (1961). BBA 51, 37. 557. Shortman, K . (1962). BBA 55, 88. 558. Gribnau, A. A. M., Schoenmakers, J. G. G . , and Bloemendal, H. (1969). ABB 130, 48. 559. Gribnau, A. A. M., Schoenmakers, J. G . G., van Kraaikamp, M., and Bloernendal, H. (1970). BBRC 38, 1064. 560. Gagnon, C., and de Lamirande, G. (1973). BBRC 51, 580. 561. Gribnau, A. A. M., Schoenmakers, J . G . G . , van Kraaikamp, M., Hilak, M . , and Bloemendal, H. (1970). BBA 224, 55. 562. Berns, A. J. M., Zweers, A., Gribnau, A. A. M., and Bloemendal, H. (1971). BBA 247, 62. 563. Ortwerth, B. J . , and Byrnes, R . J. (1972). Expt. Eye R e s . 14, 114. 564. Greif, R . L . , and Eich, E. F. (1977). Metabolism 26, 851. 565. Takahashi, Y., Mase, K., and Suzuki, Y. (1967). Experientia 23, 525. 566. Suzuki, Y., and Takahashi, Y. (1970). J . Neiirochem. 17, 1521. 567. Takahashi, Y., Mase, K., and Suzuki, Y. (1970). J . Neirrochem. 47, 1433. 568. Goto, S., and Mizuno, D. (1971). ABB 145, 64. 569. Goto, S . , and Mizuno, D. (1971). ABB 145, 71. 570. Bishay, E. S., and Nicholls, D. M. (1973). ABB 158, 185. 571. Nicholls, D. M., and Markle, H . V. (1974). C h e m . B i d . Inrercictions 8, 225.

418

PETER BLACKBURN AND STANFORD MOORE

in human placenta (572, 573), a tissue with considerable biosynthetic activity. From this readily available human tissue, the homogeneous inhibitor was isolated through the use of ion exchange chromatography and affinity chromatography on RNase A Sepharose (574). A simplified procedure employs only (NH&SOa fractionation and affinity chromatography (575). In common with the inhibitors of pancreatic RNase from other tissues, the placental inhibitor is an acidic protein with an isoionic point at pH 4.6 to 4.8. It has a mean average molecular weight, determined by SDS-polyacrylamide gel electrophoresis and gel filtration, of about 50,000, and forms a I : 1 complex with bovine pancreatic RNase .4. Ortwerth and Byrnes (563) reported a molecular weight of 32,000 for bovine lens RNase inhibitor. Bloemendal et (11. (576) obtained a molecular weight by gel filtration of near 55,000 for a preparation of calf-lens RNase inhibitor; an inactive preparation obtained by electroelution of the protein after polyacrylamide gel electrophoresis at pH 8.9 was found to give two bands on subsequent polyacrylamide electrophoresis in the presence of 6 M urea or sodium dodecyl sulfate. RNase inhibitor activity is generally assayed against bovine pancreatic RNase A by its ability to inhibit the enzymatic activity toward RNA according to the procedures of Roth (547)and Shortman (556). One unit of RNase inhibitor activity is defined as the amount of inhibitor required to inhibit the activity of 5 ng of RNase A by 50% (556). The spectrophotometric assay for bovine pancreatic RNase A toward 2' ,3'-cyclic CMP (252. 577, 578) has been adapted to the assay of purified RNase inhibitor (575). While this assay is not sensitive enough to be used with tissue extracts it provides a convenient method, and is the assay of choice for measuring the activities of purified preparations of the inhibitor. As is the case with the RNase inhibitors from bovine lens (563), rat kidney (570), and rat liver (570, 579) the placental inhibitor is a strong noncompetitive inhibitor of bovine pancreatic RNase A, with a K i of 3 x lO-'OM (574). Like the RNase inhibitor from rat liver (547),the placental RNase inhibitor is rapidly inactivated by agents that react with sulfhydryl groups, especially by p - hydroxymercuribenzoate. The inactivation is not reversed by the presence of excess free thiol, and results in dissociation of 572. Brody, S. (1957). BBA 24, 502. 573. Bardon, A., Pamula, Z , and Hillar, M.(1969). A m Biochim. Polon. 16, 119. 574. Blackburn, P., Wilson, G., and Moore, S. (1977). JBC 252, 5904. 575. Blackburn, P. (1979). JBC 254, 12484. 576. Bloernendal, H., Zweers, A., Koopmans, M., and van den Broek, W. (1977). BBRC 17, 416. 577. Richards, F. M. (1966). CR Troi.. Lob. Corlshrrg Ser. Chim. 29, 315. 578. Crook, E. M., Mathias, A. P., and Rabin, B. R. (1960). BJ 74, 234. 579. Bartholeyns, J . , and Baudhuin, P. (1977). BJ 163, 675.

12. PANCREATIC RIBONUCLEASES

419

TABLE XI11

A M I N OACIDCOMPOSlTlONS

OF

MAMMALIA RNASE N INHIBITORS Residues/Molecule"

Amino acid

Human placentab

Bovine brain'

Asx Thr Ser Glx pro Gly Ala Val Met I le Leu TY Phe His LYS Arg t cys + cys Trp Total residues

47 16 45 64 17 36 34 24 2 12

43 20 40 65 18 53 38 22 2 9 88 5 3 5 15 20 30 -5 48 1

85

4 6 6 17 23 30 5 473

' I Calculated to the closest integer fit based on a molecular weight of 51,000. From Blackburn et rrl. (574). ' From Burton ef d.(581).

active enzyme from the RNase-inhibitor complex. In the absence of free thiol, the free RNase inhibitor is rapidly inactivated; the depletion of free thiol from extracts of human placenta can lead to inactivation of the inhibitor during extraction (574. 575). Electrophoretic studies on the inhibitors from rat, ovine, and bovine tissues indicated that their RNase inhibitors were similar (580). The RNase inhibitor has been purified to apparent homogeneity from bovine brain (581) and mammalian liver (582)by procedures based upon those of Blackburn ef NI. (574, 575). For comparison, the amino acid composition of the inhibitors from placenta and brain are shown in Table XIII. Of the 580. van den Broek, W. G. M., Koopmans, M. A. G., and Bloemendal, H . (1974). W o l . B i d . RtJp. 1, 295. 581. Burton, L. E . , Blackburn, P., and Moore, S. (1980). 1u/.J . P ~ p / i d r/ndPro/ein t~ Res. 16, 359. 582. Burton, L . E., and Fucci, N . P. (1982). f i i / . J . Prpritlc Proreill Res.. in press.

420

PETER BLACKBURN AND STANFORD MOORE TABLE XIV

INTERACTION OF MODIFIED RNASEA

Reagent or derivative Reduced and carboxamido methylated BJtanedione C-clohexanedione Iodoacetate Iodoacetate Methylacetimidate Methyl-p -hydroxybenzimidate Cyanate Cyanate Bromoacetate Des-( 121- 124)-RNase Des-(l19-124)-RNase RNase S-protein RNase S-peptide RNase S

WITH

RNASEINHIBITOR

Type and number of residues modified

4 Cystine 8 Arginine 4.0 Arginine 3.34 Histidine (residue-12) Histidine (residue-1 19) Lysine 9.1 Lysine 10.0 Lysine 3 Lysine 6.6 Lysine (residue-41)

FROM

HUMAN PLACENTA"

Enzymatic activity (% of RNase A)

Strength of interaction with inhibitor (l/R&

0

5.1 3.2 7 <1 0.9 0 0 0 0 1.2 0 0.4 0 99

1.0 0.45 1.3 3.6 0.27 0.25 0.1 <0.1 0.1 1.o 1.o 1.o 10.1 1.or

" From

Blackburn and Jailkhani (47) and Blackburn and Gavilanes ( 4 5 ) . of the inhibition of RNase A by the inhibitor. ' Determined from percentage inhibition of RNase S by inhibitor.

* Rm = Molar ratio of derivative to RNase A that gives 50% reversal

30 half-cystine plus cysteine residues per molecule of placental RNase inhibitor, at least 8 are present as free sulfhydryl (574). Studies on the functional groups involved in the enzyme-inhibitor interaction to date have been performed primarily with bovine pancreatic RNase A used as a model. Knowledge of the sequence of the enzyme (15) and the details of its three-dimensional structure (1, 16), along with the wealth of data concerning the importance of specific functional groups of the enzyme, have permitted conclusions to be drawn as to which regions of this enzyme are involved in binding to the inhibitor. Specific proteolytic and chemical modifications of RNase A were made. Competition-binding experiments were performed using the 2' ,3'-cyclic CMP assay described by Blackburn (575) to examine the effect of these modifications on the ability of RNase to interact with the inhibitor (45,47). The results, some of which are shown in Table XIV, were consistent with the noncompetitive mode of inhibition. The active-site residues of

12. PANCREATIC RIBONUCLEASES

42 1

the enzyme, His-12 and His-119, and the auxiliary residues Phe-120, Asp-121, and Ser-123 are not essential for the interaction with the inhibitor. Also, it was demonstrated that the binding site for the inhibitor resides within the S-protein part of the molecule; residues 1 through 20 are not essential for the interaction. The results suggest that one or more lysine residues might be involved. When bound to the enzyme, the inhibitor protected the enzyme from inactivation by reagents that react at Lys-41 of RNase A, indicating that this residue was essential for the interaction (47). Subsequently, specific carboxymethylation of Lys-41 of RNase A with bromoacetate (81) was shown to reduce significantly the strength of the interaction between the enzyme and its inhibitor (45). Modification of the four arginine residues of the enzyme had little effect on the interaction ( 4 7 ) . Unexpectedly, carboxymethylation of the active-site residue His- 119 of RNase A (583)resulted in a 3.5-fold increase in the strength of the interaction between the enzyme and the placental inhibitor (45, 47). To assess whether the enzyme modifications that altered the interaction with the inhibitor were specific, circular dichroism measurements were performed on a number of the derivatives listed in Table XIV (45). The results, coupled to the CD data that had been reported previously by others for des-(121-124)-RNase A (142, 143), des-(119- 124)-RNase A (263), and RNase S-protein (584), suggested that one or more tyrosine residues of the enzyme were important for its interaction with the inhibitor. Based on studies of the binding of the inhibitor with different pancreatic RNases of known sequence (26/, 443, 445, 446, 454), and with the bovine seminal plasma RNase BS-1 (457)and its monomeric component, it was concluded that Tyr-92 was important for the interaction. It is hypothesized that Tyr-92 is rendered more accessible for interaction with the inhibitor as a result of carboxymethylation of His- 119 of the enzyme (45). To more clearly define the regions of the RNase molecule that are in contact with the inhibitor, the available lysine residues of the enzyme inhibitor complex have been amidinated with methyl acetimidate (585589) under conditions that preserve the complex functionally intact (47). 583. 584. 585. 586. 587. 595. 588. C89.

Crestfield, A. M . , Stein, W. H . , and Moore, S. (1963). JBC 238, 2413; ibid.. 2421. Pflumrn, M. N . , and Beychok, S . (1969). JBC 244, 3973. Lambert, J . M . , and Perham, R. N . (1977). BJ 161, 49. Hunter, M . J . , and Ludwig, M. L. (1962). JACS 84, 3491. Ludwig, M. L . , and Hunter, M . J. (1967). “Methods in Enzymology,” Vol. 1 1 , p. Browne, D. T., and Kent, S. B. H. (1975). BBRC 67, 126. Browne, D. T., and Kent, S. B. H . (1975). BBRC 67, 133.

422

PETER BLACKBURN AND STANFORD MOORE TABLE XV THEPROTECTIVE EFFECTO F RNASEINHIBITORO N THE A M I D I N A T IOF ON LYSINE RESIDUESI N RNASEA

Lysine residue Protection afforded (56)”

1 <5

7

31

37

100

100

71

41 100

61 100

66 <5

91 100

98

<5

104 <5

Calculated from relative recovery of individual tryptic peptides and their overlap peptides, the sum of which equals 100%. From Blackburn and Gavilanes (44a). If

The resistance of eacetimidyllysine residues to hydrolysis by trypsin (590) has allowed (44n), after peptide mapping, the identification of the lysine residues of the enzyme that are fully protected by the inhibitor from amidination (Table XV). Lysine residues in the enzyme that are protected by the substrate analogue polyadenylic acid from amidination are identified in Section III,A, Table I. The results indicate that the contact regions for substrate and inhibitor are not identical, as to be expected for a noncompetitive inhibitor, but there is some overlap. From the data presented in Table XV, and those reported earlier (45. 4 7 ) , studied in reference to the three-dimensional structure of the enzyme (Fig. 2), the known contact points with the inhibitor can be placed into three groups, A, B, and C, with respect to their locations in the molecule. Group A includes (a) Lys-7, (b) Lys-41, Pro-42, and Val-43, and (c) Lys91, Tyr-92, and Pro-93. Group B includes Lys-31 and Lys-37. Group C is represented by Lys-61 and adjacent residues. Groups A and B are adjacently located, whereas C lies distal to both A and B. The current hypothesis to describe the interaction involves an extensive contact between the inhibitor and RNase A that spans from A through B and on around through C. Other contact areas must lie between A , B, and C; these cannot involve the regions of the RNase A molecule where the e N H 2 groups of lysine residues 1, 66, 98, and 104 are located, since these residues are not protected from amidination. The intervening contacts between B and C possibly involve the groove formed by residues Ser-77 to Thr-82, which are in a P-structure with residues Thr-100 to His-105. The interaction between the enzyme and the inhibitor involves both polar and nonpolar residues. A key ionic interaction involves the positively charged eNH2-group of Lys-41, probably through an interaction with a negatively charged group of the inhibitor. This interaction of Lys-41 may account for the inactivation of the enzyme, since the activity of the enzyme is sensi590. Hunter, M. J., and Ludwig, M. L. (1972). “Methods in Enzymology,” Vol. 25, p. 585.

12. PANCREATIC RIBONUCLEASES

423

tive to modification of the e N H , group of this residue (19, 23, 81, 591, 592). B.

STUDIES O N IN V I T R O PROTEIN SYNTHESIS

Kraft and Shortman (552, 593) first noted that the ratio of the inhibitor to neutral RNase activity tends to increase in tissues characterized by increased rates of RNA synthesis and accumulation (e.g., 594-598); conversely, tissues in which protein synthesis decreases and catabolic activity increases usually demonstrate lower levels of the inhibitor and elevated neutral RNase activity (e .g., 596-600). A detailed discussion of the extensive literature on this subject is beyond the scope of this review. Evidence has been obtained in iGtvo (601, 602) that the inhibitor serves to preserve fully functional messenger RNA in the course of protein biosynthesis. Inclusion of the inhibitor purified from the human placenta into such systems significantly increases the incorporation of radioactive amino acids into the larger molecular weight translation products (Fig. 8). Also, it has been shown by de Martynoff crl. (603) that the synthesis of complementary DNA by reverse transcriptase is significantly improved in the presence of a preparation of placental RNase inhibitor. This finding has wide significance for genetic engineering experiments as an aid in the synthesis of full-size reverse transcripts for insertion into plasmid DNA and also for mRNA sequence determination studies. Further evidence for the role of the RNase inhibitor comes from studies on the isolation of polysomes, the stability of which is greater in tissues characterized by high levels of inhibitor activity, as seen in regenerating rat liver (604, 605) and the liver of estrogenized roosters (555). Moreover, the endogenous RNase inhibitor in rat liver high-speed supernatants (558, 591. Hirs, C. H. W. (1962). Brookhaven Syrnp. Biol. 15, 154. 592. Carty, R . P., and Hirs, C. H. W. (1968). JBC 243, 5244; ibid.. 5254. 593. Kraft, N . , and Shortman, K. (1970). BBA 217, 164. 594. Liu, D. K., Williams, G. H., and Fritz, P. J. (1975). BJ 148, 67. 595. Kyner, D., Christman, J. K., and Acs, G. (1979). EJB 99, 395. 596. Greif, R. L., and Eich, E. F. (1972). BBA 286, 350. 597. Murthy, P. V. N., and McKenzie, J. M. (1974). Endocrinology 94, 74. 598. Brewer, E. N., Foster, L. B., and Sells, B. H. (1969). JBC 244, 1389. 599. Liu, D. K., and Matrisian, P. E. (1977). BJ 164, 371. 600. Karplus, M.,and Weaver, D. L . (1976). Nrrt/rrr (London) 260, 404. 601. Scheele, G., and Blackburn, P. (1979). PNAS 76, 4898. 602. Robbi, M., and Lazarow, P. B. (1978). PNAS 75, 4344. 603. de Martynoff, G., Pays, E., and Vassart, G. (1980). BBRC 93, 645. 604. Moriyama, T., Umeda, T., Nakashirna, S., Oura, H., and Tsukada, K. (1969). J. Biochrm. (Tokyo) 66, 151. 6 0 5 . Bont, W. S., Rezelman, G., Meisner, I., and Bloemendal, H. (1967). ABB 119, 36.

;:I 424

PETER BLACKBURN AND STANFORD MOORE

(B)

12

0

O O ’

/

-1

0

C .-

a

o0 0

0

C .-

E 0

‘0

30

60 90 Time (minl

120

FIG.8. (A) Effect of human placental RNase inhibitor on the in virro translation of dog pancrease mRNA in the wheat germ system. Radioactivity incorporated into protein was measured with (+I) and without (-1) inhibitor. (B) Fluorographic analysis of SDSpolyacrylamide gel patterns obtained with equal volumes of the translation mixtures prepared as described for A. The M, values correspond to those of the presecretory proteins preamylase (55K), preprocarboxypeptidases (46K), and the serine preproteases (27K). From Scheele and Blackburn (60/).

606-609) and the purified human placental RNase inhibitor (601) have been used to protect polysomes from degradation during their extraction. VIII.

Catalytic Properties

A. ASSAYS In the decade since a review of procedures for RNase assay was published in this treatise ( I ) ,there have been improvements in the sensitivity and modes of measurement of the transphosphorylation step and the hy606. 607. 608. 609.

Bont, W. S., Rezelman, G., and Bloemendal, H. (1965). BJ 95, 1%. Blobel, G., and Potter, V. R. (1966). PNAS 55, 1283. Takahashi, Y., Mase, K., and Sugano, H . (1966). BBA 119, 627. Burghouts, J. Th. M., Stols, A. L. H., and Bloemendal, H. (1970). BJ 119, 749.

12. PANCREATIC RIBONUCLEASES

425

drolysis step. The most commonly used procedures continue to be modifications of the spectrophotometric assay toward RNA as introduced by Kunitz (610), the measurement of acid-soluble nucleotides released from polynucleotides by Anfinsen et al. (61I), and the spectrophotometric assay of the course of hydrolysis of 2’,3’-cyclic CMP by Crooker al. (578) [cf., see Richards Ref. (189)l. The sensitivity of the precipitation assay has been increased by the use of a variety of radioactively labeled RNAs. With [3H]tRNAor [32P]tRNA, less than 1 part of RNase per 10 million in preparations of DNase can be measured (12). Mendelsohn and Young (612) determined the acid-soluble radioactivity from the action of RNase on leucyl-tRNA charged with [14C]leucine.For a direct assay of the transphosphorylation step, White et al. (30) used Up[’T]C as substrate; after thin-layer chromatography, the radioactivity was counted in the areas containing [“CIC and Up[ 14C]C. When the assay of Anfinsenet al. (611) was modified (613) to precipitate acid-soluble nucleotides from very dilute solution (0.02% RNA instead of the usual 0.25% RNA), anomalous results due to incomplete precipitation were obtained (55), particularly at acid pH in the presence of low concentrations of phosphate (0.005 M ) (614). Anfinsen et al. (611) diluted the sample after (not before) the precipitation step, and this is preferable. A spectrophotometric assay has been based upon measurement of color solubilized by RNase action on an insoluble RNA-acridine orange complex (615). Fluorometric assays have been described that measure the drop in fluorescence when RNase acts on the complex of RNA and ethidium bromide (616, 617). The drop in radioactivity of an insolubilized substrate, [1251]RNA-agarose,has been used to detect RNase down to a concentration of lov8pdml(618). The measurement of the hydrolysis step, as described by Crook et af. (578), requires substrate concentrations close to the K , value for RNase A and measures the spectrophotometric change at 286 nm, which is on the 610. Kunitz, M. (1946). JBC 164, 563. 611. Anfinsen, C. B., Redfield, R. R . , Choate, W. L., Page, J . , and Carroll (1954). JBC 207, 201. 612. Mendelsohn, S . L., and Young, D. A . (1978). BBA 519, 461. 613. Bartholeyns, J., Peeters-Joris, G . , Reychler, H . , and Baudhuin, P. (1975). EJB 57, 205. 614. Bartholeyns, J . , Wang, D., Blackburn, P., Wilson, G., Moore, S . , and Stein, W. H. (1977). I n t . J . Peptide Protein Res. 10, 172. 615. Chaplinski, T., and Webster, D. A. (1973). Anal. Biochem. 54, 395. 616. LePecq, J. B . , and Paoletti, C. (1966). Anal. Biocllern. 17, 100. 617. Kamm, R. C., Smith, A . G . , and Lyons, H. (1970). Anal. Biochem. 37, 333. 618. Egly, J. M., and Kempf, J . (1976). FEES Lett. 63, 250.

426

PETER BLACKBURN A N D STANFORD MOORE

descending arm of the absorption spectrum of the substrate. Precautions necessary for reproducible recordings have been discussed by Hugli et al. (619) and Blackburn (575). An assay has been described that measures phosphate liberated by the action of alkaline phosphatase on the products of RNase action on RNA (620).

The detection of nuclease activity in bands obtained electrophoretically in SDS-polyacrylamide gels that contain RNA has been studied by Rosenthal and Lacks (621). The SDS prevents adsorption of the enzyme to nucleic acid; after extensive washing with pH 7.6 buffer to remove SDS (to allow renaturation of RNase and its subsequent action on the substrate) the presence of RNase was evidenced by a dark band under UV light after staining the gel with ethidium bromide. Alternatively, Karpetsky et al. (622) conducted the electrophoresis at acid pH, without SDS, and then incubated the gels at neutral pH to allow the enzyme to act on the incorporated polynucleotide. Spermine was included to prevent binding to the polynucleotide during electrophoresis; staining was with pyronine Y. Both techniques were sensitive to 0.5 ng of RNase A. Sierakowska and Shugar ( 2 ) have reviewed the use of chromogenic substrates, such as uridine-3‘-(a-naphthyl phosphate) and 5’-O-benzyluridine-3’-(a-naphthyl phosphate) for assay of pancreatic-like RNases. A N D ACTIVATORS B. INHIBITORS

Additions to the list of inhibitory nucleotides ( 1 ) include substrate analogs in which the carbohydrate moiety is arabinose; Pollard and Nagyvary (623) have found that Ara-3’-CMP (Ki= 0.1 mM) is bound five times as strongly as the ribose analog under the same conditions (pH 7.0, 25”, with 2’,3’-cyclic CMP as substrate). The anti conformation of arabinonucleosides (624) is considered to be a possible contributing factor to the stronger binding. White ef d.(30) have studied forty oligonucleotides as competitive inhibitors of the hydrolysis of RNA. ApUp ( K , = 0.5 mM) was the strongest of the series. Folk acid is an inhibitor of RNase when cyclic 2’,3‘-CMP is the substrate but not when RNA is the substrate (624 a ) . 619. Hugli, T. E . , Bustin, M., and Moore, S. (1973). Brtiin Res. 58, 191. 620. Stern, R . , and Wilczek, J. (1973). A n d . Biochem. 54, 419. 621. Rosenthal, A. L., and Lacks, S. A. (1977). AnuI. Biocliern. 80, 76. 622. Karpetsky, T. P., Davies, G . E . , Shriver, K . K . , and Levy, C. C. (1980). BJ 189, 277. 623. Pollard, D. R., and Nagyvary, J. (1973). Biochernisrry 12, 1063. 624. Emerson, T. R . , Swan, R. J . , and Ulbricht, T. L. V. (1967). Biochemistry 6, 843. 624a. Sawada, F., Kamesaka, Y.,and Irie, M. (1977). BBA 479, 188.

12. PANCREATIC RIBONUCLEASES

427

The inhibition of RNases is crucial in a variety of experiments that depend upon biologically active RNA. The role of the cytoplasmic inhibitor of RNases of the pancreatic type in the maintenance of functional RNA is reviewed in Section VII. Mendelsohn and Young (612) found a combination of sodium dodecyl sulfate and diethyl pyrocarbonate to be effective in the protection of RNA from degradation during its isolation. Chirgwin et ul. (274)demonstrated the efficiency of homogenization in 4 M guanidinium thiocyanate plus 0.1 M 2mercaptoethanol for this purpose. Oxovanadium ion (VO'+) forms stable complexes with nucleotide monophosphates, which are strong competitive inhibitors of RNase A (625).These complexes have been demonstrated to inhibit RNase activity very efficiently during extraction of polysomes from lymphocytes (626); they cannot be used during translation. Compounds that bind to nucleic acids can either decrease or increase the action of RNase. Chloroquine (627)at 0.13 mM can double the rate of hydrolysis of tRNA by RNase A in 0.02 M Tris-HC1 buffer at pH 7.3 and 37"; there was no effect with 2',3'-cyclic CMP as substrate. Spermine at 0.13 mM has been found to reduce the hydrolysis of tRNA by RNase A to about 50% of the control value under the above conditions (627). At lower concentrations (0.02 mM), spermine increased the activity of RNase twoto fourfold toward cyclic substrates and poly(C), but not toward poly(U) (628). The apparent K , of the substrate was not influenced; the effect was on the velocity of the reaction. In 0.01 M phosphate buffer at neutral pH, spermine has been observed to stimulate the cleavage of RNA by a human plasma RNase (629);the effect, being in part electrostatic in nature, can be expected to be less at physiological salt concentration. Wang (537) studied the kinetics of the action of RNase A cross-linked to polyspermine, a combination that shows increased ability to hydrolyze double-stranded substrates (59) at p H 7.5 in the presence of 0.125 M NaCI. The coupling of a single chain of octaspermine to the enzyme strengthens the binding to poly(A). poly(U) ( K , decreases from 270 to 2.7 pM in total U) and increases the V,,,, for hydrolysis of the susceptible poly(U) strand from 2.5 to 16.2 AAZ5*min-lmg-' of enzyme. There is evidence for inhibition by the RNase-resistant poly(A) tracts in the substrate; free poly(A) shows a K 1 of about 8 pM in total A (537). 625. Lienhard, G . E., Secemski, 1. I . . Koehler, I<. A , , and Lindquist, R . N . (1971). CSHSQB 36, 45. 18, 5143. 626. Berger, S. L . , and Birkenmeier, C. S. (1979). Eioclic~mistq~ 627. Holbrook, D. J . , Jr., Whichard, L . P., and Washington, M. E. (1975). EJB 60, 317. 628. Kumagi, H . , Igarashi, K . , Tsuji, I., Mori, C., and Hirose, S . (1980). Cliem. Phcirm. Blrll. (Jcipcrn) 28, 1189. 629. Schmukler, M., Jewett, P. B . , and Levy, C. C . (1975). JBC 250, 2206.

428

PETER BLACKBURN AND STANFORD MOORE

Jensen and von Hippel (630) examined the kinetics and the thermodynamic parameters that characterize the complex formation between RNase A and DNA; the system provides a model for studies of the destabilization of double-stranded DNA by a protein that binds to singlestranded sequences, which may be transiently exposed by fluctuations below the normal transition temperature for the double-stranded structure. C. KINETICS Rubsamen et t i / . ( 6 3 / ) observed sigmoidal kinetics in the action of RNase A at pH 7.6 on 2',3'-cyclic CMP and on esters of 3'-UMP in the substrate concentration range 1-15 mM; the results were explained by proposing a conformation equilibrium between two enzyme species in terms of the model for RNase A action proposed by Witzel and his colleagues (631).Working in a higher substrate concentration range, the initial rates of hydrolysis were observed by Walker rf t i / . (632) to change markedly with substrate concentration at pH 7.0 in Tris-HC1 buffer or in a pH-stat; there was a dip in the initial rate at 25 to 30 mM substrate, followed by a second rise and a gradual decline. Such data provided an explanation of the variations reported for K , values for 2' ,3'-cyclic CMP, which have ranged from near 1 mM to 7 mM with upper substrate concentrations of 10 to 250 mM. The authors propose an allosteric model for RNase A in which there is a substrate-dependent change in the equilibrium between the two enzyme conformations. They postulate the binding of six substrate molecules in the course of a cooperative substrateinduced transition; the binding is reminiscent of Crestfield and Allen's (633) early observation that the apparent isoionic point of RNase in phosphate buffer varied with phosphate concentration as a result of multiple binding sites for phosphate. The kinetic treatment was extended (634) to include inhibition by product or by a competitive inhibitor such as phosphate, and demonstrated (635) the decrease in the affinity for nucleotides with a decrease in the net positive charge of the protein, which can vary with the method of preparation of the enzyme, with the phosphate or sulfate content, or with the degree of deamidation upon storage of the phosphate-free protein (636). 630. Jensen, D. E . , and von Hippel, P. H. (1976). JBC 251, 7198. 631. Rubsamen, H . , Khandker, R., and Witzel, H. (1974). H o p p e Seyler's Z . Plrysiol. Cham. 355, 687. 632. Walker, E. J . , Ralston, G. B . , and Darvey, I. G. (1975). BJ 147, 425. 633. Crestfield, A. M., and Allen, F. W. (1954). JBC 211, 363. 634. Walker, E. J., Ralston, G . B . , and Darvey, 1. G. (1976). BJ 153, 329. 635. Walker, E. J . , Ralston, G. B., and Darvey, I. G. (1978). BJ 173, 1. 636. Walker, E. J., Ralston, G. B . , and Darvey, 1. G . (1978). BJ 173, 5 .

12. PANCREATIC RIBONUCLEASES

429

The temperature-dependence of the hydrolysis of 2‘ ,3‘-cyclic CMP by RNase A at pH 5 has been studied by Matheson and Scheraga (637); the data indicate that a small conformational change occurs in the enzyme near 32”, well below the temperature of the main thermal transition. Walz (638) measured the binding of deoxyuridine 3’-phosphate to assess the role of the 2’-OH group in the affinity for the enzyme. Kinetic and equilibrium binding studies showed that the deoxy derivative has an apparent Kd of 0.38 mM compared with 0.07 mM for 3‘-UMP and that the bound 2’-hydroxyl group of 3’-UMP interacts with RNase in a specific fashion that influences the interactions of the 3’-phosphate group with the enzyme, as well as an isomerization process associated with formation of the RNase-nucleotide complex. Li and Walz (639) studied the influence of a phosphate group on the 5‘-OH of 2‘,3‘-cyclicUMP; at pH 5.5 the derivative is bound more strongly ( K , is 0.03 mM, which is 23-fold lower than that for 2‘,3‘-cyclic UMP) but the turnover number was fivefold lower. The authors consider it likely that the 5’-phosphate group is subject to attraction by a group such as Lys-41. As noted in Section III,A,I, chemical and physical studies provide data for the presence [see Fig. 20 of Ref. ( f ) ] of three base- and phosphate-binding sites (B and R for base and ribose), PO,B1, R1, p1, R2, B2, pZ, R3, B3. A comparison of the kinetic parameters for the action of pancreatic RNases of known sequence from five mammalian species toward 2‘,3’cyclic CMP and UMP have been determined by Ronda et al. (2.58). The results for all five enzymes are similar, a finding consistent with the preservation of the overall features of the molecule deemed essential for activity; the main difference is in the turnover numbers rather than in the K , values. Avramova et crl. (640) used CD spectra to confirm that the very slow depolymerization of poly(A) at pH 6.5 by RNase A is a result of a low V,,,,,rather than to a change in K , ; the latter is similar to that for poly(U). Cozzone and Jardetsky (641) studied the transphosphorylation step with poly(A) by 3’P NMR. The reaction proceeds slowly at pH 7.9 in 0.1 M Tris-HC1 buffer at temperatures above 35”. The enzymatic process with poly(A) (1) is known to stop at the cyclic phosphate stage; at 160 hours the presence of a small amount of 2’- and 3’-AMP is attributed to nonenzymatic hydrolysis at pH 7.9. Libonati and associates (530, 540) studied the action 637. 638. 639. 640. Biol. 8, 641.

Matheson, R . R . , J r . , and Scheraga, H. A. (1979). Biorhemisrry 18, 2446. Walz, F. G . , Jr. (1971). B k x - h t ~ m b t r 10, . ~ 2156. Li, J. R.-T., and Walz, F. G . , Jr. (1974). A B B 161, 227. Avrarnova, 2. V., Dudkin, S . M . , and Karabashyan, L. V. (1974). Mo/eXir/yirn~r.wr 501. Cozzone, P. J . , and Jardetsky, 0. (1977). FEBS Lett. 73, 77.

430

PETER BLACKBURN AND STANFORD MOORE

RNase- A

O\/O

H,O"

"nu

s o

on

Q

s

(Et O), P 0 C I

2-

FIG.9. The method used to determine the geometry of the ring opening step. From Usher cf t i / . ( 6 4 3 ) .

of RNase A on poly(A) and on double-stranded substrates such as poly(A) . poly(U), with emphasis on the multiple effects of ionic environment on the substrate and/or on the enzyme-substrate interaction.

D. MECHANISM O F CATALYSIS Steps 1. Geometry of the TUY>

The synthesis of the two stereoisomers of uridine 2',3'-cyclic phosphorothioate by Eckstein (642) opened the way to the establishment of the geometry of the two steps in the action of RNase A [Usher et d.(643, fj44)]. The hydrolysis step, the opening of the ring of isomer N (Fig. 9) by the enzyme, was conducted in water enriched in lXO.The ring was then reclosed by a known in-line (SN2)cyclization. If the geometry of the enzymatic ring opening is also in line (as is shown in Fig. 9), then the isomer ( I that is produced should contain no excess IXO;the isotope should be in the h isomer. The results of measurement of the IXOincorporation in the two isomers were consistent with the in-line hypothesis. The transphosphorylation step was then studied (644). Advantage was taken of the reversibility of the transphosphorylation reaction (Fig. 10). The dinucleotide Up(S)C was synthesized from the CI isomer of the cyclic substrate and cytidine by the catalytic action of RNase A. The reformation of the cyclic phosphorothioate from the dinucleotide by base catalysis, which is known to follow an in-line mechanism, gave the origi642. Eckstein, F. (1970). JACS 92, 4718. 643. Usher, D. A., Richardson, D. I . , Jr., and Eckstein, F. (1970). Nutiire (London) 228, 663. 644. Usher, D. A . , Erenrich, E. S . , and Eckstein, F. (1972). P N A S 69, 115.

43 1

12. PANCREATIC RIBONUCLEASES RNase A

c +

UC(S)

d

Up(S)C

Base Up(S)C ---+U^P(S)

+ c

In-Line

F I G .10. The overall method used to test the geometry of the first step of RNase action. From Usher cf t r l . ( 6 4 4 ) .

nalu isomer (determined by NMR). The simplest explanation of these geometric results, and the one consistent with most evidence from organic chemistry, is that both steps in the action of RNase A proceed by in-line mechanisms, each of which results in the inversion of the absolute configuration around the phosphorus. Witzel and associates ( 6 3 / ) ,in their proposal for the mode of action of RNase A, postulate an adjacent mechanism; to accommodate the data of Usher et ti/. (643, 644), they invoke turnstile rotation to permit inversion of the absolute configuration as a consequence of an adjacent mechanism. 2. Confbmiirtioti of tlir Siibstmte

Several lines of investigation have been directed toward the question of the torsional angle for the glycosidic bond in the enzyme-substrate complex. Interpretation of nuclear Overhauser effects have led Karpeisky and Yakovlev (64s) to conclude that the pH-dependent conformation of 3’-CMP holds in the anti orientation up to near pH 7. Earlier NMR data (327) suggested that the syn conformer may prevail at neutral pH. Gorenstein rt (11. (646) applied molecular orbital calculations to the torsional activation of phosphodiester bonds, with reference to RNase A; they conclude that molecules in a gauche, transconformation are activated for cleavage, and consider that the data on the preferential hydrolysis of the intranucleotide linkage 33-34 in Phe-tRNA is evidence that the stereoelectronic consequences of a gauche, trans conformation are taken advantage of in this instance. 3. M e c h t i i s t i c , Models

Richards and Wyckoff ( I ) and Benz and Roberts (647)have discussed in detail the chemical and physical data upon which proposals for the mechanism of action of RNase A can be built. The two main lines of reasoning continue to yield two quite different hypotheses. The initial suggestion, 645. Karpeisky, M . Ya, and Yakolev, G . I . (1977). FEBS Lett. 75, 70. 646. Gorenstein, D. G., Findlay, J. B . , Luxon, B. A , , and Kar, D. (1977).JACS 99, 3473. 647. Benz, F. W., and Roberts, G. C. K. (1973). 1~ “Physico-Chemical Properties of Nucleic Acids” (J. Duchesne, e d . ) , Vol. 3, p . 77. Academic Press, London.

432

PETER BLACKBURN AND STANFORD MOORE

proposed on kinetic grounds by Mathias and Rabin and their colleagues, [see Ref. (I)] has over the years been fitted into the accumulated framework of structural and kinetic data on the enzyme. Deakyne and Allen (648) have reviewed this approach and applied molecular orbital theory calculations to some of the steps. The imidazole ring of His-119 is considered to activate the leaving group and to facilitate the in-line addition of 0-2’ to the phosphodiester group. Hydrogen bonds between the peptidy1 carbonyl of Thr-45 and N-1 of His-12, and N-3 of His-12 and the 2’-OH may increase the nucleophilicity of 0-2’. The backbone N-H from residue 120 can increase the electrophilicity of the phosphorus; Gln11 * HOH * phosphoryl hydrogen bonds are expected to have a similar function. Lys-41 increases the electrophilicity of the phosphorus and stabilizes a trigonal-bipyramidal intermediate. Holmes (649) discussed a square pyramidal model as an alternative to the trigonal intermediate. Rein et a / . (650)combined molecular orbital and perturbation theory to consider the configurational flexibility of the active site; their conclusions on the positioning of the two imidazole rings are consistent with those of earlier reviews (I, 647). In applying molecular orbital theory to the second step, Umeyama et al. (651) postulate that coincident with a change in the shape of the active site, a charge relay system involving Asp- 121 and His- 119 may be involved in the opening of the 2’ ,3’-cyclic substrate. Bellmann and Witzel’s (652) observation that fully purified carboxymethyl-His- 12-RNase A has no activity strengthens the view that in RNase A His-12 should have a well-defined role in whatever mechanism is proposed. The approach of Witzel and his associates (631) is entirely different. Their central premise is that the 2-0x0 group of the pyrimidine activates the 2’-OH group of the sugar in the first step, or a water molecule in the second step. A diimidazole system is proposed in which His-119 is hydrogen-bonded to His- 12. Lys-41 contributes to the electrostatic binding of the phosphate group. The ring of Phe-120 interacts with the pyrimidine ring. [Lin et a / . ’ s (255) observation that a substitution of Leu for Phe at position 120 gives a product with 13% activity indicates that an aromatic ring at this position is helpful but not essential.] Witzel and

-

648. Lkakyne, C. A . , and Allen, L. C. (1979). JACS 101, 3951. 649. Holmes, R. R. (1976). I n ? . J . Peptide Protein Res. 8, 445. 650. Rein, R., Renugopalakrishnan, V., and Barnard, E. A. (1971). In “Proceedings of the First European Biophysics Congress” (E. Broda, A. Locker, and H. Springer-Lederer, eds.), Vol. 6, p. 35. Verlag der Wiener Medizinischen Akademie, Vienna, Austria. 651. Umeyama, H . , Nakagawa, S. , and Fuji, T. (1979). Chem. Pharm. Bull. (Japan) 27, 974. 652. Bellmann, B . , and Witzel, H. (1980). Hoppe-Sryler’s 2. Physiol. Cheni. 361, 218.

12. PANCREATIC RIBONUCLEASES

433

colleagues presented kinetic analyses and NMR data in support of individual structural features of the formulation (631 ).

IX.

Research Applications

Among the uses of RNase as a probe in the study of ribonucleic acids, the enzyme is one of the nucleases widely employed in the course of the sequencing of RNA; references to the current methodology are included in the article on RNase TI (653) in this volume. ThF usefulness of the cytoplasmic inhibitor of RNase to protect RNA in the course of the synthesis of complementary DNA by reverse transcriptase is cited in Section VII; that section also covers the use of the inhibitor to protect mRNA during the course of in vitro translation and during the preparation of rough microsomes and detached polysomes. Clinical applications, in general, are subject to further research; these include the possible value of plasma or serum levels of neutral RNase as indicators of neoplastic disease (544,545).Encouraging clinical trials have been reported on the efficacy of intramuscular administration of bovine RNase A for the treatment of infection by the RNA-containing virus of tick-borne encephalitis (654). The antitumor activities of bovine seminal plasma RNase BS-1 (e.g., 655, 656) and cross-linked dimers of bovine RNase A (e.g., 57, 58, 657) merit further study. Ac K NOW L EDGMENTS The literature survey for this review grew in part from the researches under NIH Grant GM 25323 on the Biochemistry of Nucleases. The library research, organization of the bibliography, and the assembly of the manuscript were completed with the skillful cooperation of Lorraine Ackerman. The authors are especially indebted t o J. J. Beintema and associates for making available unpublished sequence data on species variations and to readers of several sections of the manuscript who provided counsel and papers in press from their laboratories.

653. Takahashi, K . , and Moore, S. (1981). “The Enzymes,” 3rd Ed., this volume, Chap. 13. 654. Glukhov, B. N., Jerusalimsky, A. P., Canter, V. M., and Salganik, R. I. (1976). Arch. Nertrol. 33, 598. 655. MatouSek, J. (1973). Experienria 29, 858. 656. Vescia, S . , Tramontano, D., Augusti-Tocco, G., and D’Alessio, G. (1980). Cuncer Res. 40, 3740. 657. Bartholeyns, J . , and Baudhuin, P. (1976). P N A S 73, 573.

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Ribmuclease T 1 KENJI TAKAHASHI

STANFORD MOORE

I. Introduction . . . . . . . . . . . . 11. Purification and Chemical Properties . A. Purification . . . . . . . . . . . B . Modification of Functional Groups . C . Physical Parameters . . . . . . . 111. Reactions Catalyzed . . . . . . . . A. Specificity of the Catalytic Reaction B. Interaction with Substrate Analogs C. Steady State Kinetics . . . . . . D. Mechanism of Catalysis . . . . . IV. Research Applications . . . . . . . A. Determination of Sequences in RNA B . Additional Applications . . . . . V. Other Guanine-Specific RNases . . ,

I.

. . . . . . .

. ,

. , ,

. ,

,

. . .

.

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

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

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.

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. . . . . .

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

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.

. . .

435 436 436 437 444 447 448 449 458 461 463 463 464 465

Introduction

Ribonuclease T I was discovered by Sato and Egami ( I ) in 1957 as the major ribonuclease in Takadiastase, a commercial enzyme mixture from Aspergillus orryztre. They demonstrated that the acidic protein is a guanyloribonuclease (EC 3.1.27.3) that at neutral pH effects a two-stage endonucleolytic cleavage of RNA to 3‘-phosphomono- and oligonucleotides ending in Gp, with 2’ ,3’-cyclic phosphate intermediates. Uchida and Egami (2 reviewed the knowledge of RNase T I to 1970 in Vol. IV of 1. Sato, K., and Egami, F. (1957). J . Biochrtn. (Tokyo) 44, 753. 2. Uchida, T., and Egami, F. (1971). “The Enzymes,” 3rd ed., Vol. IV, p. 205.

435 THE ENZYMES. VOL. XV Copyright @ 1982 by Academic Press. Inc All rights of reproduction in any form reserved.

ISBN 0-12- 1 2 2 7 6 4

436

KENJI TAKAHASHI AND STANFORD MOORE

this series; Takahashi (3) summarized the chemistry to 1971. Egami el al. ( 4 ) reviewed the specific interactions of both RNase T1 and RNase Uz with their substrates. This chapter covers mainly the information accumulated between 1970 and 1980 on the chemistry of RNase T, and its biochemical applications. II. Purification and Chemical Properties

A.

PURIFICATION

The previously reviewed (2) methods of purification, based primarily upon ammonium sulfate precipitation and chromatography on basic ion exchangers, have been extended by Fields et a/. ( 5 ) to a procedure that utilizes only DEAE-cellulose in several steps to give 350-380 mg of RNase T, (yield 48%) from 500 g of Takadiastase Y powder; Walzst a/. (6) find that phenol extraction is necessary to remove minor contaminants from the enzyme thus prepared. Fulling and Ruterjans (7) isolated the enzyme in 80% yield by chromatography on sulfopropyl-Sephadex at pH 3 and elution of the enzyme as a complex with 2‘-GMP. Affinity chromatography has been applied to the preparation of RNase T1by several investigators. Jervis and Pettit (8) used buffer extraction, gel filtration on Sephadex G-100, and chromatography on a column of 5’(4-aminophenylphosphoryl)guanosine 2’(3’)-monophosphate-glass.Other affinity adsorbents have been used, such as acetylated phosphocellulose containing guanosine 2’ (3’)-phosphate ( 9 ) , a Sepharose derivative carrying 5’-(4-aminophenylphosphoryl)guanosine 2’(3’)-phosphate ( l a ) , NADP-agarose ( 1 1 ), and guanylyl-(2‘-5‘)guanosine coupled to aminohexyl-Sepharose (12). 3. Takahashi, K. (1972). In “Proteins: Structure and Function” (M. Funatsu, K. Hiromi, K. Imahori, T. Murachi, and K. Narita, eds.), Vol. 1 , p. 285. Kodansha, Tokyo; Wiley, New York. 4. Egami, F., Oshima, T., and Uchida, T. (1980).In “Molecular Biology, Biochemistry and Biophysics” (F. Chapeville and A. L. Haenni, eds.), Vol. 32, p. 250. Springer Verlag, Berlin. 5 . Fields, R., Dixon, H. B. F., and Law, G. R. (1971).BJ 121, 591. 6 . Walz, F. G., Jr., Osterman, H. L., and Libertin, C. (1979).ABB 195, 95. 7 . Fulling, R., and Ruterjans, H. (1978).FEBS Left. 88, 279. 8 . Jervis, L., and Pettit, N. M. (1974).J. Chromufogr. 97, 33. 9. Waki, M., Mitsuyasu, N., Terada, S., Matsuura, S., Kato, T., and Izumiya, N. (1974). BBRC 61, 526. 10. Jervis, L. (1974). Phytochernisfry 13, 723. 1 1 . Janski, A. M., and Oleson, A. E. (1976).Anal. Biochem. 71, 471. 12. Ishiwata, K., and Yoshida, H. (1978).J . Biochem. (Tokyo) 83, 783.

437

13. RIBONUCLEASE T ,

Kanaya and Uchida (13) have developed a method for purification of RNase T, using AH-Sepharose 4B coupled to 5’-GMP as an affinity adsorbent. The procedure utilizes batchwise treatment with DEAEcellulose, DEAE-cellulose chromatography, and affinity chromatography. B.

MODIFICATION OF FUNCTIONAL GROUPS

The chemistry of the enzyme can be discussed in reference to the sequence established by Takahashi (14, 15-19) (Fig. 1). The amino acid residues present in the 104 residue chain are Asp-6, Asn-9, Thr-6, Ser-15, Glu-6, Gln-3, Pro-4, Gly-12, Ala-7, t Cys-4, Val-8, Met-0, Ile-2, Leu-3, Tyr-9, Phe-4, Trp-1, Lys-1, His-3, and Arg-1; MW 11,085; PI 2.9 (20). Martin et ul. (21) have obtained crystals suitable for X-ray diffraction studies of the enzyme; Heinemann et a/. (22) have crystallized the RNase T1-2’-GMP complex.

-

1.

Cctrboxyl Groups

The finding that the inactivation of RNase T1 by iodoacetate at pH 5.5 involves specific esterification (23) of Glu-58 led to the conclusion ( 3 )that the yCOOH group of this.residue is essential for activity. The pH-rate profile (23, 2 4 ) for the carboxymethylation indicates the participation of two groups with pK, values of about 4 and 7, which probably correspond to the functional groups of Glu-58 and a histidine residue. The carboxymethylated derivative retains the 1 : 1 binding capacity of the enzyme for 3’-GMP (25);thus Glu-58 is more directly concerned with the catalysis than with the binding of substrate. Tosylglycolate similarly inactivates RNase T1 at pH 5.5 (26) and protection is afforded by substrate analogs. The protective effect decreases in the order 3’-GMP > 5‘-GMP > 3’Kanaya, S ., and Uchida, T. (1981). J . Biochem. ( T o k y o ) 89, 591. Takahashi, K. (1965). JBC 240, 41 17. Takahashi, K. (1971). J . Biochem. ( T o k y o ) 70, 477. Takahashi, K. (1971). J . Biochem. ( T o k y o ) 70, 603. Takahashi, K. (1971). J . Biochem. ( T o k y o ) 70, 617. Takahashi, K. (1971). J . Biochem. ( T o k y o ) 70, 803. Takahashi, K. (1971). J . Biochem. ( T o k y o ) 70, 945. Takahashi, K . (1962). J . Biochem. ( T o k y o ) 51, 95. 36, 95. Martin, P. D . , Tulinsky, A , , and Walz, F. G., Jr. (1980). 22. Heinemann, U . , Wernitz, M., Pahler, A , , Saenger, W., Menke, G., and Riiterjans, H. (1980).EJB 109, 109. 23. Takahashi, K., Stein, W. H . , and Moore, S. (1967). JBC 242, 4682. 24. Takahashi, K . (1970). J . Biochem. ( T o k y o ) 68, 517. 25. Takahashi, K . (1972). J . Biochem. ( T o k y o ) 72, 1469. 26. Oshima, H . , and Takahashi, K. (1976). J . Biochem. ( T o k y o ) 80, 1259. 13. 14. 15. 16. 17. 18. 19. 20. 21.

1

11

12

13

I4

15

16

17

18

19

20

21

23

22

25

24

26

29

28

27

-Ser~Thr~Ala-*Gln-...\la-r;\la-*Gl~-tT~-r -*Gln+Leu+His+Glu

TIhr 32

ucTyTcAsntr\sntT!.reL!.stHistPro.-T!.rt Ser+Asn+Ser+

51

, 98

Asn

97

Gly

96

SLr

55

56

43

44

45

57

58

42

59

40

41

60

61

62

37

38

39

63

64

35

36

65

66

34

67

Pro+T~r+T!.r+Glu-tTrp~Pro+Ile+Leu-tSer +Ser+GIy-tAsp+val-Tyr

r

A

6, - 70 53

92

91

90

89 FIG.

88

1.

87

86

85

84

83

82

81

80

79

a;

77

76

The primary structure of RNase TI. From Takahashi (14).

75

i4

73

72

13. RIBONUCLEASE TI

439

AMP > 3'-CMP > guanosine. The relatively weak effect of guanosine suggests that Glu-58 is located near the site where the phosphate group of the substrate analog interacts. The introduced Cm group is more stable than the Cm group of free y-Cm-glutamic acid, suggesting that the introduced Cm group is stabilized by interaction with other groups in the enzyme. This interaction also seems to stabilize the conformation of RNase T1, since Cm-RNase T I shows a higher transition temperature than native RNase T1,as examined by measuring the decrease in absorbance at 278 nm as a function of temperature (K. Takahashi, unpublished). 2. Atnitio Croirps The nonessentialities of the a-NH2 group of Ala-1 and the e N H 2 group of Lys-41 ( 3 ) have been further demonstrated by nitrotroponylation (27) of the two groups; the derivative retains 70% of the native activity. The two amino groups can be nitroguanidinated by N-methyl-N'-nitro-Nnitrosoguanidine without much loss of activity (K. Takahashi, unpublished). On the other hand, acylation of the two amino groups with several dicarboxylic acid anhydrides, such as succinic, maleic, citraconic, and c6.-aconitic anhydrides, leads to extensive inactivation of the enzyme (28).The inactivation is probably aresult of the carboxy! group introduced at the €-amino group of Lys-41, which is adjacent to His-40. The added carboxyl group may interact with a positively charged group near the active site, or interfere with the binding of RNA to the active site through electrostatic repulsion and/or steric hindrance. The introduced acyl groups, except the succinyl group, can be removed at pH 3.6 with concomitant regeneration of activity. Upon maleylation, the K, value toward RNA is increased about 20-fold while the V,,,, value toward RNA is reduced about 3-fold. With GpC as substrate, Kay and Dixon ( 2 9 ) found that RNase TI in which the two amino groups are maleylated has the same K, as the native enzyme but a threefold lowered kc,, value. The introduced maleyl groups thus interfere more with the interaction of the enzyme with RNA than with GpC. The amino groups can be coupled to various solid supports to prepare active water-insoluble derivatives of RNase TI. The enzyme has been coupled to carboxymethylcellulose ( 3 0 ) and to a cross-!inked polyacrylamide (Enzacry!) (31) by the acid azide method and to cyanogen 27. Tamaoki, H . , Sakiyarna, F., and Narita, K . (1976). J . Eioclic~i?i.( T o k i w ) 79, 579. 28. Takahashi, K . (1977). J . Biochrnz. ( T o k y o ) 81, 641. 29. Kay, J . , and Dixon, H . B . F. (1973). Ahstr-. 9fk 117fcrii.Coirgr-. Bioi,/it'iii., Sfock/io/rn, July 1-7, p. 107. 30. Kuriyama, Y . , and Egami, F. (1966). Seiktrgrrkrr 38, 735. 31. Ito, H . , Hagiwara, M . , Takahashi, K . , and Ichikizaki, I. (19771.J. Bioclwn. ( T o k y o ) 82, 877.

440

KENJI TAKAHASHI AND STANFORD MOORE

bromide-activated Sepharose and Sephadex ( 3 2 ) . These derivatives are generally more stable than the original enzyme, although the activity is reduced. The carboxymethylcellulose-bound enzyme had about 2 and 50-60% of the original activity toward yeast RNA and 2' ,3'-cyclic GMP, respectively. On the other hand, the Enzacryl-bound enzyme had about 45 and 77% of the original activities toward yeast RNA and 2' ,3'-cyclic GMP, respectively; the coupled enzyme is fairly stable in the range of pH 1 to 10 at 37", and is active toward RNA even above pH 9 (at 37") or above 60" (at pH 7 . 3 , conditions under which the native enzyme is inactive. 3. Histidine Residirrs The sensitivities of the enzymic activity and of the histidine residues to methylene-blue catalyzed photooxidation (2 ) have been examined further by Waku and Nakazawa ( 3 3 ) and by Irie ( 3 4 ) . They have shown that two histidine residues are protected from methylene blue-catalyzed photooxidation by the presence of 2'(3')-GMP. Irie (34) has also shown that a group with pK, value of 7.5 is involved in the photooxidative inactivation and that the binding ability toward 2'(3')-GMP is decreased by photooxidation while the gross conformation around tyrosine and tryptophan residues is not affected significantly. Similar results were obtained with rose bengal as a photosensitizer ( 3 ) ; over 70% inactivation takes place when, on the average, one histidine residue per molecule is oxidized, and only 0.2 residue of tryptophan is altered. That this modification involves primarily one of the three histidine residues was demonstrated through study of the histidine-containing peptides obtained by chymotryptic hydrolysis ( 3 ) ;His-92 is the most reactive of the three. Substrate analogs protect the enzyme from photooxidation; the order of protective effect is 2'(3')-GMP > 5'-GMP > 2'(3')-AMP2'(3')-CMP has no effect. Upon photooxidation the enzyme markedly loses the reactivity of Glu-58 toward iodoacetate ( 3 )and the binding ability toward 3'-GMP (34, 3 5 ) . Through study of the inactivation of the enzyme with iodoacetamide, which does not react with Glu-58 ( 2 3 ) , Takahashi (36, 3 7 ) has shown that the loss in activity toward RNA proceeds in parallel with the loss of two histidine residues; the pH-rate profile of this inactivation implicates residues with pK,'s in the 7.5-8 range (36, 3 7 ) . His-92 and His-40 react with 32. 33. 34. 35. 36. 37.

Lee, J. C. (1971). BBA 235, 435. Waku, K., and Nakazawa, Y. (1970). J. Biochern. (Tokyo) 68, 63. Irie, M. (1970). J . Biochern. (ToXvo) 68, 69. Takahashi, K. (1971). J . Bioclwrn. ( T o k y o ) 69, 331. Takahashi, K. (1973). J. Biocliem. (Tokyo) 74, 1279. Takahashi, K. (1976). J . Biochem. (Tokyo) 80, 1267.

13. RIBONUCLEASE TI

44 1

iodoacetamide most rapidly and at similar rates, whereas His-27 is least reactive. Alkylation of His-92 is much slower when the Glu-58carboxymethylated enzyme is treated with iodoacetamide. On the other hand, alkylation of His-40 is slowed down most in the presence of 3’GMP. These results suggest that His-92 and His-40 are involved in the active site and that His-27 is partially buried in the enzyme molecule or interacts strongly with some other residue, thus becoming relatively unreactive. The reactivity of histidine residues is reduced by prior modification of Lys-41 with maleic or cis-aconitic anhydride or 2,4,6trinitrobenzene- I-sulfonate, or of Arg-77 with ninhydrin. Similar alkylation of histidine residues takes place optimally at around pH 8-8.5 by reaction with NO-bromoacetyl-1.-arginine methyl ester, w-bromoacetamidoethylnicotinic acid amide, or 4-(iodoacetamido)salicylic acid, and in most cases the NHz-terminal alanine residue is alkylated concomitantly ( 2 4 ) . As in the case of iodoacetamide reaction, the enzyme is inactivated in parallel with the loss of two histidine residues. The enzyme is not inactivated by such alkylating reagents as bromoethylamine, ethyleneimine, or l-chloro-3-tosylamido-7-amino-2-heptanone (TLCK). 4. Argiriine Modification of the single arginine residue at position 77 by phenylglyoxal results in parallel loss of activity toward RNA and 2‘,3’-cyclic GMP (3). The reactivity of the y-carboxyl group of Glu-58 is lost almost in parallel with the loss of the arginine residue. The binding ability of the enzyme toward 3’-GMP is also lost by this modification. A similar loss of 3’-GMP-binding ability has been reported by Tamaoki ef al. (27) upon selective cleavage of the Arg-77 bond with trypsin. These results indicate that Arg-77 is present at or near the active site of the enzyme. It may be involved in the binding of negatively charged substrates or in building the active site structure of the enzyme. Arg-77 is also modified by ninhydrin with loss of activity, and the two amino groups of the enzyme are modified simultaneously (38). 5.

Trypf o pha n

Studies on the single tryptophan residue at position 59, adjacent to Glu-58, have shown that in the native enzyme the indole ring is relatively inaccessible to chemical modification (2, 3) but that its integrity is essential for folding the chain into the active conformation. This conclusion is supported by experiments on the selective oxidation of Trp-59 by ozone to N’-formylkynurenine and subsequent conversion of the residue to one of 38. Takahashi, K . (1976). J . Biochem. ( T o k y o ) 80, 1173.

442

KENJI TAKAHASHI AND STANFORD MOORE

kynurenine by acid-catalyzed deformylation in the frozen state (39). These two enzyme derivatives are devoid of activity at pH 7.5, but retain measurable activity at pH 4.75. The reactivity of Glu-58 toward iodoacetate and the binding ability of the enzyme toward 3'-GMP are also decreased by these modifications. 6. Tvrosine Residrres Chemical and physical experiments ( 2 , 3 )have shown that most of the 9 tyrosine residues are not exposed on the surface of the enzyme; there is no evidence for direct involvement of tyrosine in the catalytic site. This conclusion is supported by studies with N-acetylimidazole as an acetylating agent (40). Three to four tyrosine residues are acetylated fairly readily at pH 7.5 without extensive loss of activity. p -Diazobenzenesulfonic acid reacts readily with 1 to 2 tyrosine residues. In 8 M urea, all of the tyrosine residues are acetylated with nearly complete loss of activity; the acetyl groups can be removed by incubation with hydroxylamine with concomitant regeneration of full activity.

7. Disuljde Bonds Cleavage of both of the disulfide bonds by reduction reversibly inactivates the enzyme (2 ). The bridge between Cys-2 and Cys- 10 is reduced much more rapidly than that between Cys-6 and Cys-103 (41), and measurable but not complete loss of activity results from reduction of the former. In 2 M NaCl a folding of the fully reduced chain can proceed without formation of -S-Sbonds to yield 25% of the original enzymatic activity (42 1. 8. Enzymutic ModiJiccitions Early results ( 2 ) showed that the threonine residue at the COOH terminus can be removed without loss of activity, that the NHrterminal alanine residue is resistant to leucine aminopeptidase, and that in the presence of 0.2 M phosphate the molecule is relatively resistant to the action of trypsin or chymotrypsin at pH 7 and 37". In the absence of phosphate, the bonds most susceptible to tryptic cleavage at 50" are Lys41-Tyr-42 and Arg-77-Val-78 (43). When the derivative in which the two amino groups are selectively blocked by nitrotroponylation (27) or trinitrophenylation (43) is treated 39. 40. 41. 42. 43.

Tamaoki, H., Sakiyama, F., and Narita, K. (1978). J . Biochem. ( T o k y o ) 83, 771. Kasai, H., Takahashi, K., and Ando, T. (1977). J . Biochem. (Tokyo) 81, 1751. Hayakawa, S . , and Takahashi, K . (1973). J . Biochem. (Tokya) 74, 1075. Oobatake, M., Takahashi, S., and Ooi, T. (1979). J . Biochem. (Tokyo) 86, 65. Takahashi, K . , and Inoue, N. (1977). J . Biochem. ( T o k y o ) 81, 415.

13. RIBONUCLEASE TI

443

with trypsin, the Arg-77-Val-78 bond can be fairly specifically cleaved with concomitant loss of activity. The modified enzyme is devoid of the ability to bind 3’-GMP, and its circular dichroism spectrum suggests that its conformation has been extensively altered (27). These results indicate the importance to the activity of RNase T1of Arg-77 andor the continuity of the peptide chain at this residue. 9. Synthetic Studies Hofmann and his colleagues have aimed for the total synthesis of RNase TI by a solution method based on fragment condensation (44-47). They considered the peptide chain of the enzyme in terms of seven fragments, A (1-1 I), B (12-23), C (24-34), D (35-47), E (48-65), F (66-80), and G (81-104). The individual fragments were synthesized by stepwise procedures. They combined, by the azide method, the appropriate fragments to give AB and CD. These two peptides were then condensed to give fragment ABCD. Fragment E F was similarly prepared. A coupling to yield CDEFG (residues 24-104) has been achieved (48), but the synthesis of the active enzyme in this manner remains to be accomplished. Waki et al. (9) have prepared, by the stepwise solid-phase method, polypeptides with the sequence of RNase T, and of its analog, [59-Tyr]RNase T1, in which Trp-59 is replaced with a tyrosine residue. After gel filtration on Sephadex G-50, reduction with 2-mercaptoethanol and reoxidation in air, the crude peptide mixture was extensively fractionated and an active component was purified by ion exchange chromatography on DEAE-cellulose and acetylated phosphocellulose. The purified peptide corresponding to RNase TI (yield, 0.07% from the crude peptide) had 59 and 44% of the specific activity of the native enzyme toward RNA and 2’ ,3’-cyclic GMP, respectively. The purified peptide corresponding to L59-TyrI-RNase TI (yield, 0.02% from the crude peptide) had 61 and 5796, respectively, of the native specific activities. This result indicates that the residue at position 59 of RNase T1 need not necessarily be a tryptophan residue, but may be replaced by other aromatic amino acid residues. 44. Yanaihara, N., Yanaihara, C . , Dupuis, G . , Beacham, J . , Camble, R., and Hofmann, 91, 2184. 45. Storey, H. T., Beacham, J . , Cernosek, S . F., Finn, F. M., Yanaihara, C., and Hofmann, K. (1972). JACS 94, 6170. 46. Kawasaki, K., Carnble, R . , Dupuis, G . , Romovacek, H . , Storey, H. T., Yanaihara, C., and Hofmann, K. (1973). JACS 95, 6815. 47. Romovacek, H . , Drabarek, S., Kawasaki, K . , Dowd, S. R., Obermeier, R., and Hofmann, K. (1974). Intern. J . Peptide Protein R e s . 6 , 435. 48. Romovacek. H., Dowd, S. R., Kawasaki, K . , Nishi, N . , and Hofmann, K. (1979).

K. (1969).JACS

JACS 101, 6081.

444

KENJI TAKAHASHI AND STANFORD MOORE

C. PHYSICAL PARAMETERS 1. Dissociable Groups

In extension of the studies on the dissociation constants of functional groups in RNase TI (2), the results of pH-induced spectral titrations at around 300 nm led Walz (49) to the identification of a group with a pKa of 4.29 that was attributed to the y-COOH of Glu-58; this pK, value decreases at low ionic strength, which suggests the proximity of cationic side chains. The PKa values of the three histidine residues have been determined by nuclear magnetic resonance (NMR) titration of the signals of the C-2 protons of each histidine residue separately (50,51); the direct assignments (Table I) were made by NMR studies coupled with tritium or deuterium exchange and chemical modifications (7, SO). The pKa values have also been estimated by hydrogen-tritium exchange titration coupled with isolation of the His-containing peptides from proteolytic hydrolysates (52).The relatively high values for these pK,'s could reflect interaction with neighboring negatively charged groups in the three-dimensional structure of the enzyme. Through steady-state kinetic studies of RNase Tl-catalyzed transesterification reactions, Osterman and Walz (53) obtained evidence for two protonated groups having apparent pK, values of 7.5 and 8.1, which they correlate on mechanistic grounds with His-40 and His-92. In their NMR studies, Arata et al. (50) found that in the presence of 3'-GMP the His-40 and His-92 peaks are strongly influenced, but the His-27 titration curve is affected to a much less extent. This is consistent with the conclusion reached by Takahashi (36,37) that His-40 and His-92 are involved in the active site of the enzyme. His-40 seems to interact with a charged group with a pKa of 4.1, which may be Glu-58. Inagaki et al. (51) analyzed the pH dependencies of the C-2 and C-4 proton chemical shifts at 270 MHz in detail, and confirmed the assignments of histidine C-2 proton resonances. Further, they analyzed the pH dependency of the C-2 and C-4 proton chemical shifts of histidine residues of Cm-RNase T1, and of the I3C chemical shifts of the y-carboxymethyl group of IT-enriched Cm-RNase T1 (Table I). An inflection around pH 4 on the C-2 proton titration curve of His-40 disappears upon car49. Walz, F. G., Jr. (1977). Biochemistry 16, 4568. 50. Arata, Y.,Kimura, S., Matsuo, H., and Narita, K. (1979). Biochemistr.y 18, 18. 51. Inagaki, F., Kawano, Y., Shimada, I., Takahashi, K., and Miyazawa, T. (1981). J . Biochem. (Tokyo) 89, 1185.

52. Kimura, S ., Matsuo, H . , and Narita, K. (1979). J . Biochem. (Tokyo) 86, 301. 53. Osterman, H. L., and Walz, F. G . , Jr. (1978). Biochemistry 17, 4124.

445

13. RIBONUCLEASE TI

TABLE I pK,

V A L U E S FOR I N D I V I D U A L H I S T I D I N E

Method

RESIDUES IN RNASET1 A N D CM-RNASE TI

His-27

IH NMR ‘H NMR Tritium exchange

RNase TI RNase TI Cm-RNase TI RNase TI

7.2 7.26 7.07 7.3

?

5

0.01 0.01

His-40

His-92

Ref.

7.9 7.92 5 0.02 7.70 2 0.01 7.7

-8 7.80 2 0.03 7.64 5 0.01 7.6

(50 ) (51) (51) (52 )

boxymethylation of the enzyme, a finding consistent with the hypothesis of proximity to Glu-58. From the pH titration curve of the ‘ T O O H of the carboxymethyl group of Glu-58 in Cm-RNase T1, an unusually low pK, value (0.82) for the carboxymethyl group was found, which is ascribed to the formation of a salt bridge a n d o r a hydrogen bond between the carboxymethyl group and a positively charged group, possibly Arg-77. They also measured the pH dependencies of the fluorescence intensity at 360 nm of RNase TI and Cm-RNase T I to show that the inflections at about pH 4 and 7.5 observed for RNase T I disappear upon carboxymethylation. Kimura et al. (52) measured the hydrogen-tritium exchange rates of the C-2 protons of the individual histidine residues in RNase TI as a function of pH at 37” and 0.2 M ionic strength. Inagaki et 01. (51 ) measured the hydrogen-deuterium exchange rates of the C-2 protons of the individual histidine residues in RNase T1 and Cm-RNase T1 at 37” in 0.2 M NaCl using NMR spectroscopy. Both sets of results, which are summarized in Table 11, show that the hydrogen exchange rates decrease in the order: His-40 > His-92 > His-27. Kimura et a/. (52) deduced from a Bransted plot (log k, versus pK,) for model imidazole compounds and the histidine residues in RNase A and RNase T1 that His-40 is exposed on the molecular surface, while His-27 and His-92 are embedded in the molecule to extents similar to that of His-12 in RNase A. Similar conclusions have been reached by Inagaki et id. (51 ). 2. Spectral Studies In spectrophotometric studies [see Ref. (211 on RNase T1 = 21,200 M-’ cm-’, pH 7) (20), Campbell et al. (54) measured the solvent accessibility of tyrosine and tryptophan residues by solvent perturbation difference spectroscopy, and reported that 2 of the 9 tyrosine residues are at the surface and freely accessible to solvent, and that the single tryptophan residue and 2-3 additional tyrosine residues are close enough to 54. Campbell,

M .

K., Shipp, S . , and Jantzen, E. (1976). BERC 72, 1014.

446

KENJI TAKAHASHI AND STANFORD MOORE TABLE I1

HYDROGEN-TRITIUM A N D HYDROGEN-DEUTERIUM E X C H A N GRATES E OF T H E c-2 PROTONS OF INDIVIDUAL HISTIDINE RESIDUES I N RNASETi A N D CM-RNASE TI'' Tritium exchangeb RNase TI

Residue His-27 His-40 His-92

(X

RNase T1 (pH 8.7)

x. wax

tila

lO-'hr-')

(hrs)

0.31 2.13 0.42

Deuterium exchange'

223 34 166

(X

kJI lo-' hr-') 0.88 4.6

1.17

t 112 (hrs)

Cm-RNase TI (PH8.0) k$ ( x lo-' b - l )

79 15 59

0.2 2.7 0.63

" k + = Pseudo first-order rate constant; t l i 2= Calculated from the corresponding k $ value. At 37" and 0.2 ionic strength, pH near 8. From (52). At 37" in 0.2 M NaCl. From ( 5 [ ) .

the surface to be available for long-range interactions with solvent. Eftink and Ghiron (55) reported that the fluorescence of the supposedly buried Trp-59 residue is collisionally quenched by acrylamide with a rate constant of 3 x lo8 M -' sec-'; they conclude that the result indicates fluidity in the protein matrix. Oobatake ef nl. (56) found the thermal transition temperature of the enzyme at pH 5.0 to be 56.7" (AHtr= 117 kcal/mol) from the measurements of fluorescence intensity at 320 nm as a function of temperature. From the fluorescence measurements, they also showed that the conformation of the enzyme can be stabilized greatly by the presence of salts, such as 1-2 M KF or NaCl (42); disulfide-reduced RNase Tz regains a spectrum similar to that of the native enzyme in 2 M KF. Similar results were obtained through circular dichroism spectra (42, 56). From the curve-fitting analysis of the circular dichroism data, they estimated the secondary structures as follows: a-Helix contents, 11.8% at 32", 11.9% at 54", and 11.6% at 66.5"; P-structure, 51.3% at 32", 35.3% at 54", and 27.8% at 66.5". From these data they conclude that the thermal denaturation of RNase TI involves a decrease in &structure but not in a-helix content. Kasai et al. (40) used optical rotatory dispersion to study the effect of acetylation of the tyrosine residues by N-acetylimidazole. 55. Eftink, M. R., and Ghiron, C. A. (1975). PNAS 72, 3290. 56. Oobatake, M., Takahashi, S., and Ooi, T. (1979). J. Biocliem. ( T o k y o ) 86, 55.

13. RIBONUCLEASE TI

111.

447

Reactions Catalyzed

The initial characterization of RNase TI as a guanyloribonuclease has been reviewed by Uchida and Egami (2). Further evidence on the resistance to RNase T, of nucleosides in which the N-7 of guanosine is substituted is provided by studies on the modified nucleoside Q [7-(4,5-cisdihydroxy-I-cyclopenten-3-ylaminomethyl)-7-deazaguanosine](57). The cleavage of RNA by transphosphorylation can be assayed by spectrophotometric measurement of acid-soluble nucleotides after digestion of yeast RNA at pH 7.5 and 37” ( I , 20, 58). The enzyme is inhibited by organic solvents (59); at 20% by volume, the extents of inhibition were 36, 41, 62, and 90% with dioxane, methanol, ethanol, and 1-propanol, respectively. RNase T, can also be determined with a highly sensitive recording spectrophotometer (60) by measuring small changes in absorbance at 298.5 nm of RNA upon digestion. The second and slower step, the hydrolysis of the 2’,3‘-cyclic ester to the 3’-nucleotide can be determined by measuring alkali consumption, with the use of a pH-stat, upon hydrolysis of 2’,3’-cyclic GMP at pH 7.2 (61, 62), or by measuring the extent of hydrolysis by separating the substrate and 3’-GMP chromatographically on a column of Amberlite IRA-400 (XE-119) (23) or other appropriate anion exchanger (31). The use of a recording spectrophotometer equipped with a flow-through cell or a high-performance liquid chromatography apparatus is convenient for this purpose; RNase T, activity can also be determined by recording the increase in absorbance at 280 nm or its vicinity upon the action of the enzyme on GpC or other GpNs (49, 63, 64). Among metal ions tested, the strong inhibition by Zn2+ and Cu2+ is especially notable (65). No naturally occurring macromolecular inhibitors of RNase TI are known, except for antisera to the enzyme (66). 57. Kasai, H . , Ohashi, Z . , Harada, F., Nishimura, S., Openheimer, N . J., Crain, P. F., Liehr, J . G . , von Minden, D. L., and McCloskey, J. A. (1975). Biochemistry 14, 4198. 58. Takahashi, K . (1961). J . Biocliem. (Tokyo) 49, I . 59. Takahashi, K. (1974). J . Biochem. (Tokyo) 75, 201. 60. Oshima, T., Uenishi, N . , and Imahori, K. (1976). Anal. Biochem. 71, 632. 61. Kuriyama, Y. (1966). J . Biochem. (Tokyo) 59, 596. 62. Yoshida, N . , and Otsuka, H . (1971). BBA 228, 648. 63. h i e , M. (1968). J . Biocliem. ( T o k y o ) 63, 649. 64. Ipata, P. L., Felicioli, R. A., and Zucchelli, G. C. (1969). Ifcrl. J . Biocliem. 18, 114. 65. Egami, F., Takahashi, K., and Uchida, T. (1964).Progr. Nttcleic Arid Res. Mol. Bid. 3, 59. 66. Uchida, T. (1970). J . Biockem. ( T ~ k y o68, ) 255.

448

KENJI TAKAHASHI AND STANFORD MOORE

Ribose binding

site

Secondary

Catalytic site

binding site

FIG.2. A model for the mode of interaction of the substrate with the active site of RNase TI. Bl, B2;catalytic residues.

A.

SPECIFICITY OF THE CATALYTIC

REACTION

A discussion of the enzyme-substrate interaction can be made in reference to a working hypothesis for the factors involved in the binding and the catalysis with a common binding site assumed for the two steps in the catalytic process (Fig. 2). The free 2’-OH is obligatory for the formation of the 2‘,3‘-cyclic product. The similarity of the process to that with RNase A (67) suggests that P-0 bond cleavage rather than C - 0 bond cleavage occurs. Oshima and Imahori (68) have concluded from circular dichroism studies on the binding of substrate analogs that the preferred conformation of protonated guanosine (69) is the syn form and the base has been so diagrammed. The base specificity studies (2) have shown that preferred natural sub67. Hilmoe, R. J . , Heppel, L. A., Springhorn, s. s.,and Koshland, D. E., Jr. (1961). BEA 53, 214. 68. Oshima, T., and Imahori, K. (1972). I n “Proteins: Structure and Function” (M. Funatsu, K. Hiromi, K. Imahori, T. Murachi, and K.Narita, eds.), Vol. 1, p, 333. Kodansha, Tokyo; Wiley, New York. 69. Guschlbauer, W., and Courtois, Y. (1968). FEBS Lett. I , 183.

449

13. RIBONUCLEASE TI

strates for RNase TI have the keto group at position 6, trivalent N at position 7, a proton at N-1, an NH2group at position 2, and p-D-ribose in the carbohydrate moiety. Ivanova et crl. (70)have confirmed the need for the keto group at position 6 by finding that hydrolysis does not occur with the 2’ ,3‘-cyclic phosphate of purine riboside; they also observed that the cyclic derivatives from 8-aminoguanosine and 8-bromoguanosine are cleaved, which shows that the structure at C-8 is not crucial. The cleavage of 8-bromoguanosine 2’ ,3’-cyclic phosphate has also been reported by Yuki and Yoshida (71). As has been noted ( 2 ) , the base specificity of RNase T I is not absolute; at high concentrations for long periods of time, other cleavages are measurable, although double-stranded substrates are remarkably resistant. The bases in the adjacent nucleotides (R and R’ at the secondary binding sites in Fig. 2) can be purine or pyrimidine.

B.

INTERACTION W I T H SUBSTRATE

ANALOGS

Insight into the binding process has been gained through study of the interaction with a variety of substrate analogs and through kinetic analyses. 1.

Gel Filtration Stirdies

Takahashi (3) reviewed the use of gel filtration on Sephadex (3-25 to examine the interaction of enzyme and nucleotide. Sat0 and Egami (72) first investigated the interaction of guanosine 2’phosphate and related compounds with RNase T I using a gel filtration method in which a mixture of the enzyme and a substrate analog is passed through a column of Sephadex G-25 to examine the extent of complex formation. Using this method, they found that the enzyme and 2’-GMP bind in approximately 1 : 1 molar ratio at pH 5.0 whereas the complex is scarcely formed at pH 7.2, and that guanosine 2‘-sulfate and the benzyl ester of guanosine 2’-phosphate do not bind appreciably to the enzyme at pH 5.0. Campbell and Ts’o (73) performed a similar study employing the same method, but using [2’-14C]GMP.They estimated the binding constants for various unlabeled compounds by a competition method with [2’-’‘C]GMP. 70. Ivanova, G. S . , Holy, A., Zelinkova, E., Bezborodova, S. I., AbrosimovaAmelyanchik, N . M., and Tatarskaya, R. I. (1974). Collecrion Czech. fhern. C o r n m ~ n 39, . 2986. 71. Yuki, R., and Yoshida, H. (1971). E 5 A 246, 206. 72. Sato, S., and Egami, F. (1965). 5iffChem.Z. 342, 437. 73. Campbell, M. K., and Ts’o, P. 0. P. (1971). EEA 232, 427.

450

KENJI TAKAHASHI AND STANFORD MOORE

The binding affinity decreases in the order: 2’-GMP > 3‘-GMP > 2‘-IMP > 2’-XMP > 2’-AMP > 5’-GMP > guanosine. In addition, 9(2’-hydroxyethy1)guanine 2‘-phosphate and its 4‘-hydroxybutyl isomer were found to bind to RNase TI almost as strongly as 2’-GMP. Takahashi (25, 7 4 ) performed quantitative and extensive studies using a gel filtration method based on the technique of Hummel and Dreyer (75), in which the enzyme, dissolved in a solution of a substrate analog, is passed through a Sephadex G-25 column equilibrated and eluted with the same substrate analog solution. The K d values obtained are summarized in Table 111. The binding is strongest around pH 5 , while the pH optima for the two steps of the catalytic process are at pH 7.5 and 7.2. The order of the binding strengths correlates with the knowledge of the groups in the substrate that facilitate catalysis ( 2 ) ,and with the information available on the protective effect on inactivation of the enzyme by carboxymethylation of Glu-58 (26, 72, 7 6 ) or by photooxidation of histidine residues (35). Guanosine binds to the enzyme fairly strongly; this shows that the guanosine portion is of primary importance for specific binding to the enzyme, although the phosphate portion further strengthens the binding. The binding of the phosphate group to the enzyme does not appear to be as specific as that of the guanosine portion, but its position is still important. The order of the binding strengths of guanylic acids, 2’-GMP > 3’-GMP > 5’-GMP, is similar to that of cytidylic acids to pancreatic RNase A (77). The fact that guanosine binds much more strongly to the enzyme than deoxyguanosine indicates the importance of the 2’-OH group of ribose for the binding of nucleosides to the enzyme. On the other hand, 9-(2’-hydroxyethy1)guanine 2’-phosphate binds to the enzyme quite strongly, although this compound has no ribose portion; a similar result was obtained by Campbell and Ts’o (73). The ribose ring may not be required for the binding of nucleotides to the enzyme, but the carbohydrate moiety is, of course, an obligatory element in the enzymatic action. In contrast to the fairly strong binding of guanosine to the enzyme, the other nucleosides examined, except 8-bromoguanosine, bind to the enzyme only very weakly or not at all under the conditions employed at pH 5.5. These results indicate that the N-1 and N-7 positions, and 2-amino and 6-0x0 (or hydroxy) groups on the purine ring are important for the strong binding to the enzyme. The interaction of the guanine portion with the 74. 75. 76. 77.

Takahashi, K. (1970). J . Biochern. ( T o k y o ) 68, 941. Hummel, J. P., and Dreyer, W. J . (1962). BBA 63, 530. Terao, T., and Ukita, T. (1967). BBA 149, 613. Anderson, D. G., Hammes, G. G . , and Walz, F. G . , Jr. (1968). Biochemistry 7, 1637.

TABLE 111 DISSOCIATION CONSTANTS OF RNASETl- A N D CM-RNASE T

RNase T 1

2'-GMP 3'-GMP

2'-9-GHEtPb 3'3'-GDP 5'-GMP 3'-AMP 5'-1-MeGMP Guanosine

5.5 2.9 3.7 4.7 5.5 7.2 7.8 8.4 5.5 5.5 5.5 5.5 7.7 5.5 2.8 3.7 4.7 5.5

l

-

S ANALOG ~ ~ COMPLEXES ~ ~ ~DETERMINED ~ ~ ~B Y T H E GEL FILTRATION METHOD"

Guanosine

6.5

8.7 7.6 110 250 620 21 18 68 300 540 1100 1300 270 120 120

7.7

8.4

21 17

8-Bromoguanosine Deoxyguanosine

Cm-RNase T1

3'-GMP

Guanosine

2',3'-cyclic GMP I

GPA

5.5

3.7 5.5 7.7 2.9 3.6 5.5 7.7 3.6 5.5 7.7 5.5 7.7 5.5 7.7

300 440 250 520 700 1700 280 56 18 86 180 97 170 58 250 71 250

From Takahashi (25). Temp. 25" Kd values for RNase Tl were above lo00 PM for 3'-CMP, 3'-UMP, 1-methylguanosine, NZ-methylguanosine, N ' , N 2dimethylguanosine, 6-thioguanosine, 7-rnetbylguanosine, inosine, xanthosine, adenosine, and guanine. 9-(2'-hydroxyethyl)guanine2'-phosphate.

452

KENJI TAKAHASHI AND STANFORD MOORE

enzyme appears to take place at several of these points. In line with the results of Ivanovaet al. (70) and Yuki and Yoshida (71), the C-8position does not appear to be involved in this interaction, since 8-bromoguanosine binds to the enzyme nearly as strongly as guanosine. Inosine and xanthosine bind to the enzyme only weakly, although the 2’,3’-phosphodiestersof these nucleosides are known to be hydrolyzed by the enzyme. The NH2 group on C-2 thus appears to contribute greatly to the binding, as has been pointed out by Campbell and Ts’o (73). This contribution represents about - 2.6 kcaymol toward AGO for binding (6). The pH dependence of the binding of 3‘-GMP and guanosine to the enzyme (3) indicates a role in the binding of at least two groups in the enzyme with pK, values of about 4 and 7, respectively, presumably a carboxyl and an imidazole group. The Glu-58 carboxymethylated, inactive RNase TI retains a considerable binding ability toward 3’-GMP, and moreover possesses almost the same binding ability toward guanosine over a wide pH range as that of the native enzyme (Table 111). This indicates that the carboxymethylation of Glu-58 affects mainly the binding of the phosphate portion, and only slightly the binding of the nucleoside portion, to the enzyme. The carboxylate anion of the y-carboxymethyl group on Glu-58 may interact with the site responsible for the binding of the phosphate portion of 3’-GMP, or cause an electrostatic repulsion against the phosphate anion of 3’-GMP, thus weakening the binding ability of the enzyme. Sawada et al. (78) investigated the interaction of 6-thioguanylic acid and its homologs with RNase TI using the gel filtration method of Hummel and Dreyer, together with UV-difference spectroscopy and CD spectroscopy. The order of the affinities for RNase TIat pH 5.6 is 2’(3’)-GMP, 2’(3‘), 5’-GDP > 2’(3’)-Ss GMP, 2’(3’), 5’-S6 GDP, 2’,3’-cyclic, 5’-S6 GDP > 5‘-S6 GMP > 2’,3’-cyclic S 6 GMP, S‘-guanosine. The binding ability toward 3’-GMP is lost by measures that unfold the protein, such as a temperature of 60” or solution in 8 M urea (25). Although alcohols are markedly inhibitory to the enzyme (59), no significant loss of binding ability toward 3‘-GMP has been observed (25) in the presence of 20% (v/v) methyl, ethyl, or n-propyl alcohol. ZnC12, a strong inhibitor of the enzyme, is also without effect on the binding ability at 1 mM concentration. These agents appear to affect directly the catalytic efficiency rather than the binding ability of the enzyme. 2. Spectral Data Sat0 and Egami (72) first investigated the interaction of RNase T I and 2’-GMP and related compounds by UV-difference spectral measurements. 78. Sawada, F., Samejima, T., and Saneyoshi, M. (1973). BBA 299, 596.

13. RIBONUCLEASE T I

453

The difference spectrum has a negative trough at around 250 nm and a positive peak near 290 nm. A mixture of 5’-GMP or 2’-AMP and the enzyme gave a less marked difference spectrum, whereas a difference was scarcely observed with guanosine 2’-sulfate, the benzyl ester of 2‘-GMP, guanosine, 2’-CMP, and 2‘(3’)-UMP. The change in absorbance at 290 nm by increasing the molar ratio of 2‘-GMP to RNase TI indicated that the nucleotide binds to the enzyme in an approximately I : 1 molar ratio. From the pH dependence of the difference spectrum, they concluded that the monoanionic form rather than the dianionic form of 2’-GMP is preferably bound by the enzyme, and that a group with a pKa value of about 6.0 to 6.5 participates in this binding, which might be attributed either to an imidazole group in the enzyme or to the secondary dissociation of the phosphate group of the nucleotide, or to both. Oshima and Imahori (68) reviewed their studies (79) and those of others on difference spectra observed upon the binding of various guanine derivatives. They investigated the difference spectra between acid and neutral solutions of 3‘-GMP and 9-methylguanine, and showed that these difference spectra are remarkably similar to the corresponding spectra observed for RNase TI-substrate analog complexes. They concluded that the N-7 nitrogen may be protonated by a proton-donating group in the enzyme upon formation of the enzyme-substrate analog complex. Upon carboxymethylation of Glu-58, the Kd value for 3’-GMP decreases nearly 8-fold (see Table IV). From this result, they suggested a possibility that Glu-58 may be the residue for the protonation of the N-7 of the guanine base. This assumption is contrary to that made by Takahashi that Glu-58 is primarily concerned with catalysis rather than with binding (25, 26, 74, 80). Epinatjeff and Pongs (81) made similar spectral studies. The temperature dependence of the binding constants was also measured. The pH dependencies of the different ligand-enzyme complex formations indicated several pKa values (2.5, 3.7,6.5, and 8.5). The pK, values of 2.5 and 6.5 were assigned to protonations of the ligand, and those of 3.7 and 8.5 to specific groups in the active site of RNase TI that are involved in the recognition of the guanine base. The results were taken to indicate that the N-1 and N-7 positions of the guanine base may be recognized by a basic group (possibly a histidine) and an acidic group (possibly a carboxylate) in the active site of the enzyme, and that the monoanionic phosphate moiety may interact with a basic group (either histidine or Arg-77) in the enzyme. 79. Oshima, T., and Imahori, K . (1971). J . Biochem. (Tokyo) 69, 987. 80. Takahashi, K . (1970). J . Biochem. (Tokyo) 67, 833. 81. Epinatjeff, C . , and Pongs, 0. (1972). E J B 26, 434.

TABLE IV DISSOCIATION CONSTANTS OF RNASETi- A N D CM-RNASET,-SUBSTRATE ANALOG COMPLEXES DETERMINED B Y UV-DIFFERENCE SPECTROSCOPY ~

Analog RNuse T,

2'-GMP 3'-GMP

PH 5.0 5.6 5.0

5.6 3'-dGMP 3'5'-GDP 5'-GMP 5'-dGMP APG CPG UPG Guanosine

Kd

5.0

lOOC

5.0

3.5 4.0 5.0 5.6 6.0 7.0

Guanosine

19°.d

5.0 5.6 5.0 5.0

Analog

6.9", 4.gd 9' 12' 83°*d 49' 115".d 30' 123", 64d

5.0 5.0

~

~~~~

(PM)

485' 769' 592b 493 292".b 350e 319b 327b

Deoxyguanosine

_

PH

Cm-RNuse T ,

2'-GMP 3'-GMP 3'-dGMP 5'-GMP 5'-dGMP Guanosine Deoxyguanosine

_

_

_

_

Kd

(P M )

8.0

5006

8.5

526b 11206

9.0 3.5 5 .O 5.6 8.5

9-Methylguanine

_

9.0 5.6 5.0 5.0 5.6 5.0 5.0 5.0 5.0 5.0

2000b 1890". 1820b 1900e 20006 1750b 3000" SOd 63d 90e 129d 37d 45d 145' 154b

13. RIBONUCLEASE T,

455

Walz and co-workers (82-86) have made more detailed UV-difference spectral studies. Their calculated K d values are summarized in Table IV together with those obtained by Oshima and Imahori (68, 79). There is qualitative but not quantitative agreement between the spectroscopically determined values and those measured by gel filtration (Table 111). The binding of guanosine and deoxyguanosine with RNase T1 can also be distinguished in terms of the wavelength for maximal difference absorbance between pH 5.0 and 7.0. With Cm-RNase TI, the binding constants and the nature of the difference spectra for guanosine and deoxyguanosine at pH 5.0 are the same. These results suggest that the discrete interaction of the guanosine 2’-hydroxyl group with RNase T, involves the y-carboxylate group of Glu-58 and an imidazolium group at the active site. Similar studies have been done at pH 5.0 with pGp, NpG (N = A, C, or U),dGpdN (N = A, C, G, T), dTpdG, pdNpdG (N = A or T), pdGpdN (N = A, G, or T), and c(pdGpdN) (N = A or G), and the characteristic difference spectrum and association constant for (1 : 1) RNase T I binding were determined for each ligand (85).The results indicate that the guanine moiety of each ligand is bound at the primary recognition site of the enzyme. Evidence for a specific enzyme subsite for binding of the adenine moiety of ApG and pdApdG is presented. Further, the binding of RNase T1 (from pH 3.0 to 8.5) and Cm-RNase T I (at pH 5.0) with a series of guanine nucleotides and some of their methyl esters was studied (86). At pH 5.0 the order of the affinities for the enzyme is 2’GMP S 3’-GMP % 5’-dGMP 1 3 ‘ - d G M P > 5’-GMP % 5’-GMP(Me) = 3’-dGMP(Me) > 5 ‘ dGMP(Me); that for Cm-RNase T1 binding is 5’-GMP 2 5’-dGMP = 2’-GMP 2 3‘-GMP > 3’-dGMP 15’-GMP(Me). The results suggest the existence of a phosphomonoester group binding locus at the active site that normally binds with a guanosine 3’-phosphate group by virtue of an interaction with the ligand 2’-hydroxyl group. On the other hand, the enzyme preferentially interacts with ligand 3’-phosphodiester groups, regardless of its interaction with the guanosine 2‘-hydroxyl group. In addition, the interaction of RNase T, with calf thymus DNA was investigated (83). The results indicate that the enzyme does not bind with doublestranded DNA, but binds with denatured DNA by direct interaction with exposed guanine residues. Pongs (87) investigated the binding of substrate analogs to RNase T, by 82. 83. 2, 1 1 . 84. 85. 86. 87.

Walz, F. G., Jr., and Hooverman, L. L. (1973). Biochemistry 12, 4846. Walz, F. G., Jr., Biddlecome, S., and Hooverman, L. L. (1975). Nucleic Acids Rrs. Walz, F. G., Jr. (1976). Biochemistry 15, 4446. Walz, F. G., Jr. and Terenna, B. (1976). Biochpmistrv 15, 2837. Walz, F. G., Jr. (1977). Biochemistry 16, 5509. Pongs, 0. (1970). Biochemistrv 9, 2316.

456

KENJI TAKAHASHI AND STANFORD MOORE

measuring their quenching effect on the tyrosyl and tryptophanyl fluorescence of the enzyme at pH 6.4. The wavelength of excitation used was 280 nm, and those of emission were 295 nm for tyrosine and 360 nm for tryptophan. Among the substrate analogs examined, 3’-GMP produced the greatest decrease in the tyrosyl and tryptophanyl fluorescence intensities, whereas 3’-CMP did not significantly affect the fluorescence intensities. Irie (88) investigated the quenching effect of some nucleotides on the tryptophanyl fluorescence of RNase T1 at pH 5.0, using an excitation wavelength of 295 nm and an emission wavelength of 320 nm. The results show that the order of the binding strengths and the estimated dissociation constants are 2’(3)‘-AMP(0.4 x 10-4M) > 5’-AMP(1.2 x 10-4M) > 2’(3’)-CMP(1.6 x 10-4M). The maximum of the emission spectrum remains at 320 nm after the formation of an enzyme-ligand complex, indicating that the binding of a nucleotide does not appreciably change the state of Trp-59. Oobatake et al. (56) reported from fluorescence measurements that the RNase TI-2’-GMP complex shows a transition temperature about 6“ higher than that for the free enzyme at pH 5.0. This is thought to be due to some fixation of amino acid side chains caused by the binding of 2’-GMP. Sander and Ts’o (89) reported that the circular dichroism spectrum in the 240-310 nm region of a mixture of RNase T1with 2’- or 3‘-GMP or some related substrate analogs, including 3’-dGMP and 9-(2‘-hydroxyethy1)guanine 2‘-phosphate and its 4’-hydroxybutyl isomer differs significantly from the algebraic sum of the circular dichroism of the enzyme and that of the nucleotide measured separately. Strong extrinsic Cotton effects are induced with positive and negative dichroic bands at 250 and 280 nm, respectively. The characteristic features of the difference spectrum suggest a ligand-induced exiton coupling effect, which they assume could arise from electronic interactions between the purine group and the aromatic tyrosines and/or tryptophan located at or near the active site of the enzyme. They showed that the formation of the RNase TInucleotide complex as indicated by the induced CD bands has a qualitative relation to the binding constants obtained by gel filtration techniques. Oshima and Imahori (68) also investigated by CD spectra the interaction of 3’-GMP and related substrate analogs with RNase T1. They showed that upon mixing with RNase T1, 3’-GMP, 2’-GMP, 5’-GMP, 8-bromoguanosine 2’(3’)-phosphate, guanosine, 8-bromoguanosine, and deoxyguanosine all give similar CD-difference spectra characterized by a peak around 250 nm and a trough at around 280 nm. The difference re88. Irie, M. (1970). J . Biochem. (Tokyo) 68, 31. 89. Sander, C., and Ts’o,P. 0. P. (1971). Biochemistry 10, 1953.

13. RIBONUCLEASE T,

457

sembles quantitatively the dichroic spectrum of 3’-GMP in acid. On the other hand, upon protonation of the guanine base the preferred conformation of guanosine is considered to be changed from the anti to the syn conformation (69 ). From these comparisons, they concluded that the guanosine ligand is fixed into the syn conformation in the complex with RNase T1. The dichroic spectrum of the mixture of RNase T1 and 2’(3’)AMP, however, is nearly identical with the graphical summation of the respective spectra, suggesting that the adenosine group is not restricted to a certain conformation in the complex, and that the aromatic residues in the enzyme do not reveal any significant change in the optical activity upon binding of adenylate. In nuclear magnetic resonance studies, Ruterjans et al. (90) and Ruterjans and Pongs (91) investigated the binding of 3’-GMP and guanosine to RNase T,. Upon addition of 3’-GMP to an RNase TI solution, major changes occur in the chemical shifts of the C-2 proton magnetic resonance signals of two histidine residues (which they designated A and C) and of the proton magnetic resonance signals of the aromatic amino acid residues. On the other hand, only the C-2 proton magnetic resonance titration curve for the third histidine (designated C) changes upon addition of guanosine to an RNase Tl solution. The curve is analogous to that for histidine C in the RNase TI-3’-GMP complex. This result was interpreted to mean that histidine C is located near the binding site of the guanine ring of 3‘-GMP, and presumably plays a role in the specific recognition of the guanine base. The downfield shift of the C-2 proton magnetic resonance signal of histidine A appears to be due to the interaction of this histidine residue with the phosphate group of 3’-GMP. Arata et al. (50, 92) have also studied the interaction of 3’-GMP with RNase T1 by NMR. Their analysis of the chemical shifts of the C-8 proton of 3’-GMP in the absence and presence of RNase T1 indicates that it is quite unlikely that 3‘-GMP is protonated in the enzyme-inhibitor complex at neutral pH. This conclusion is different from that drawn from ultraviolet difference spectral data at 290 nm by Oshima and Imahori (68, 7 9 ) and Epinatjeff and Pongs (81). The 31PNMR titration curve for 3‘-GMP observed in the presence of RNase T1 gives a pK, of 6.5 for the second dissociation of the phosphate group of the inhibitor. This means that at pH 5 . 5 , where 3’-GMP most strongly binds to RNase T1, the phosphate group of the inhibitor exists as the monoanion. This result is consistent with that reported by Sato and Egami (72), who first pointed out that the 90. Ruterjans, H., Witzel, H., and Pongs, 0. (1969).EERC 37, 247. 91. Ruterjans, H., and Pongs, 0. (1971). EJE 18, 313. 92. Arata, Y . , Kimura, S., Matsuo, H . , and Narita, K . (1976). EERC 73, 133.

458

KENJI TAKAHASHI AND STANFORD MOORE

monoanionic form of 3’-GMP binds to RNase T, more strongly than the dianionic form. 3. Enzyme Inhibition Irie (93, 94) reported that the inhibitory effects of mononucleotides on the activity of RNase T, toward 2‘,3’-cyclic GMP decrease in the order 2’-GMP > 3’-GMP > 5‘-GMP > 3’-CMP > 2’(3‘)-UMP. The preferable binding form of 2’-GMP with the enzyme is presumed to be the monoionic species from the pKi-pH profile. From the pKi-pH profiles of RNase T, with 2’-GMP as a competitive inhibitor and 2‘ ,3’-cyclic GMP or GpC as a substrate, three groups with pK, values of 3.4, 5.7, and 7.5, respectively, were suggested to be involved in the enzyme-substrate complex formation (63, 93). Pongs (95) measured the inhibitory effects of mononucleotides and various other compounds on GpC cleavage by RNase T,. White et al. (96) measured the inhibitory effects of guanylyl2’-5’ nucleotides on the RNase T1activity toward GpU. Both (2‘-5’)GpG and (2’-5‘)GpA are effective inhibitors, whereas (2‘-5’)GpU and (2’-5’)GpC have no inhibitory effect on RNase T1under the conditions employed. (2’-5’)GpG is the strongest inhibitor among these nucleotides and its Kfvalue at pH 7.5 was estimated to be 0.165 m M . C. STEADY STATEKINETICS Whitfeld and Witzel (97) first investigated the rate of cleavage of various dinucleoside (3’-5’)-phosphates by RNase T,. They showed that the relative rates decrease in the order (the values in parenthesis are relative rates at pH 7.4): GpCp (1100) > GpC (800) > GpA (550) > GpG (450) > GpU (250) > IpC (150) > XpC (10) > glyoxal-GpC ( 5 ) > 2‘,3‘cyclic GMP (2). This result shows that the neighboring nucleosides have a considerable effect on the rate of cleavage of 3‘-guanylyl phosphodiester bonds, and that the rate of hydrolysis of 2’ ,3’-cyclic GMP is extremely slow as compared with the rates of cleavage (i.e., transphosphorylation) of GpNs. Irie (63) determined the K m and V,,, values of RNase T1toward dinucleoside monophosphates, GpN (N = A, C, G, and U), as substrates at pH 7.5 and 5.0. The K m values of the four substrates are of a similar order of 93. 94. 95. 96. 97.

hie, M. (1%7). J . Biochern. ( T o k y o ) 61, 550. Irie, M . (1964).J . Biochern. ( T o k y o ) 56, 495. Pongs, 0. (1968). Thesis, Univ. Marburg, Marburg. White, M. D., Rapoport, S , , and Lapidot, Y. (1977). BBRC 77, 1084. Whitfeld, P. R., and Witzel, H. (1963). BBA 72, 338.

459

13. RIBONUCLEASE TI

A 3

4

5

6

7

8

9

PH FIG.3 . Plots of the logarithm of k,,,lK, and ka,for the RNase TI-catalyzed transesterification of GpA and GpG and theoretical curves for all four GpNs versus pH in 0.2 M buffer. (0-) GpA; (----) GpC; (A----)GpG; (. . .) GpU. Reproduced from Osterman and Walz (53).

magnitude whereas the V,,, values vary considerably. The V,, values decrease in the order GpC > GpG > GpA > GpU, while the Vma,/Km values decrease in the order GpC > GpA > GpG > GpU, which is qualitatively similar to the result obtained by Whitfeld and Witzel (97). From the PKm-pH and log V,,,,,-pH profiles using 2‘ ,3‘-cyclic GMP or GpC as a substrate, the presence of three dissociable groups with pK, values of 3.5, 5.7, and about 7.5 in the free enzyme, and pK, values of 3 . 7 , 6 . 7 , and about 7.4 in the enzyme-substrate complex were deduced. Yoshida and Otsuka (62) reported the K,,, and kcatvalues of RNase TI at p H 7.4 toward 2’,3’-cyclic GMP, 2’,3’-cyclic IMP, and RNA. Zabinski and Walz (98) and Osterman and Walz (53) have performed rigorous studies on the steady state kinetics of the RNase TI-catalyzed transesterification of GpN (N = C, U, A, and G ) substrates. The results obtained at 0.2 M ionic strength and 25” are shown in Fig. 3. The analysis was carried out on the assumption of four ionizable groups in the enzyme (Fig. 4) (53). The pH dependencies of kca,lK, for the four dinucleoside monophosphates are similar and suggest the involvement in binding a n d o r catalysis at the active site of two unprotonated groups on the free enzyme having apparent pK, values of 3.4 and 4.3 and two protonated groups having apparent pK, values of 7.5 and 8.1 (Table V). The group with pK, value of 3.4 appears to correspond to that of an unidentified carboxyl group and those with pK, values of 4.3, 7.5 and 8.1 to those of

460

KENJI TAKAHASHI AND STANFORD MOORE

EH

Xl H

EH

X"H

K q

K:]j E

Kgl

Xi

II

i

E

X"

FIG.4. A general pH dependent mechanism for the RNase TI-catalyzed transesterification of dinucleoside monophosphate substrates. EHt, Xj,Ht-+ XjHl, S, and P represent free enzyme, complex enzyme, substrate (i.e., GpX; X = A, C , G, or U),and product (i.e., 2,3,-cyclic GMP), respectively. Protons have been omitted for clarity. Reproduced from Osterman and Walz (53). TABLE V BEST-FITKINETIC PARAMETERS" Substrates Parameter

Units ~~~

~

sec-' M x 105 (M . sec)-l x

~~~~~

~

GpA

GpC

GpG

GpU

96 5.5 1.7 3.7 4.1 7.4 8.2 <2.5 3.3 8.2 19

350 16 2.2 4.0 4.1 7.7 8.3 <2.5 4.0 8.8 >9

62 2.7 2.3 3.2 4.3 7.7 7.6 C2.5 3.4 7.8 >9

38 2.2 1.8 2.9 4.6 7.3 8.1 <2.5 3.0 8.3 29

Average values

~

~

3.4 2 0.5 4.3 f. 0.2 7.5 2 0.3 8.1 -+ 0.3

~

" From Osterman and Walz (53). Mechanisms used are in Fig. 4. The data were obtained using 0.2 M buffer. Similar results were obtained using 0.02 M buffer. The mechanism in Fig. 4 is characterized by

and

where kcal/Km is the ratio of the pH independent value of the turnover number and the Michaelis constant, K I . KK. K;, and KB are macroscopic acid dissociation constants characterizing pertinent groups on the free enzyme and KX, K*,, K$, and KE are apparent dissociation constants that characterize the generalized scheme in Fig. 4.

46 I

13. RIBONUCLEASE T,

Glu-58, His-40, and His-92, respectively. The pH-independent values of k,,,lK, characterizing the four substrates are virtually identical, while the individual values for k,,, and K, range within an order of magnitude of each other and follow the sequence: GpC > GpA > GpG > GpU. Walz et al. (6) also performed kinetic studies on the RNase T,-catalyzed transesterification of 12 dinucleoside monophosphates, N'pN' (N' = A, C, and U; N2 = A, C, G, and U) as well as IpU and IpC at pH 5,25", and 0.2 M ionic strength and compared the kinetic parameters thus obtained with those obtained for GpNs (53, 98). The results indicate that ApN dinucleoside monophosphates are 106-foldless efficient when compared with GpN substrates, whereas, IpC and IpU are 10'- to 103-fold less efficient than the corresponding GpNs. No quantifiable kinetics were obtained with CpNs and UpNs. Values of k,,,lK, show the order NpC > NpU (N = A, G, and I), which appears to indicate the existence of a subsite for the leaving nucleoside group that binds cytidine. Osterman and Walz (99) have further studied the RNase T1-catalyzed transesterification of the trimeric substrates ApGpC and ApGpU [to (ApG > p + C)and (ApG > p + U)] in steady-state kinetic experiments performed at 25" and 0.2 M ionic strength over the pH range 3-9. The results indicate that the adenosine moiety of the trimeric substrate binds with an enzyme subsite and that catalysis can proceed via three parallel reaction paths that are governed by apparent pK, values of 5.2 and 7.7 in the enzyme-substrate complex. From these results they conclude that the mechanism of action on these trimeric substrates is different from that for dimeric substrates such as GpC and GpU. Further knowledge of the roles of subsites will ultimately be required for a fuller understanding of the mechanism of the action of the enzyme on polymeric substrates.

-

D. MECHANISM OF CATALYSIS The stereochemistry of the transesterification step of RNase T1 has been investigated by Eckstein et ul. (100). RNase Tl was found to hydrolyze only the endo isomer of the mixture of the two diastereomers of guanosine 2',3'-cyclophosphorothioate, without loss of sulfur. On methanolysis of the mixture of isomers by the enzyme, only the endo isomer reacts to give guanosine 3'-phosphorothioate 0 -methyl ester. Based on these results, the transesterification step of RNase T, has been suggested to follow a sterically in-line mechanism. 98. Zabinski, M., and Walz, F. G . , Jr. (1976). AEE 175, 558. 99. Osterman, H . L., and Walz, F. G., Jr. (1979). Eiochemisrry 18, 1984. 100. Eckstein, F., Schulz, H. H., Riiterjans, H . , Haar, W., and Maurer, W. (1972). 6iochrrnisrr.v 11, 3507.

462

KENJI TAKAHASHI AND STANFORD MOORE

L FIG.5 . A proposed mechanism for the action of RNase TI.Based on Takahashi (80).

In early chemical modification studies, the implication of one or two histidine residues (80, 101) and Glu-58 (23) in the active site of RNase TI was suggested. Taking these results into consideration, Takahashi (80) proposed a reaction mechanism, as shown in Fig. 5 , in which a histidine residue works in concert with Glu-58, one as a general base and the other as a general acid, in a similar mechanism as postulated for His-12 and His-119 in RNase A (102, 103). A similar mechanism was also suggested for RNase TI by Pongs (95). In the mechanism shown in Fig. 5 , Glu-58 acts as a general base and histidine as a general acid in the transphosphorylation step, while Glu-58 acts as a general acid and histidine as a general base in the hydrolysis step. A mechanism in which the roles of these two residues are mutually exchanged is also conceivable. The possibility cannot be excluded, however, that two histidine residues constitute the catalytic site, as in the case 101. Yamagata. S . , Takahashi, K . , and Egarni, F. (1962). J . Eiochem. (Tokyo) 52, 261. 102. Findlay, D., Herries, D. G . , Mathias, A. P., Rabin, B. R., and Ross, C. A. (1962). EJ 85, 152. 103. Roberts, G. C. K . , Dennis, E. A., Meadows, D. H . , Cohen, J. S., and Jardetzky, 0. (1969). P N A S 62, 1151.

13. RIBONUCLEASE TI

463

of RNase A. Further chemical modification studies have shown that His-40 and His-92 are involved in the active site (36, 371, and that Arg-77 may also be involved (104). Kinetic studies with synthetic substrates (53, 63, 93, 98) have shown the involvement of carboxyl group(s) and imidazole group(s) in binding and/or catalysis, while gel filtration (25, 7 4 )and UV-difference spectroscopic studies (81 ) have indicated the involvement of such groups in the binding of substrates, especially for recognition of the guanosine residue. Moreover, His-40 has been suggested to be part of the catalytic site from NMR studies coupled with tritium exchange experiments (50, 92). Judging from these results, it seems most likely that Glu-58 and His-40 are primarily involved in catalysis while His-92 participates in guanosine binding. This is consistent with the proposed reaction mechanism shown in Fig. 5 . Osterman and Walz (99) have recently described a modified model in which Glu-58 is replaced with the Glu-58His-92 pair. IV.

Research Applications

A. DETERMINATION OF SEQUENCES I N RNA The base specificity of RNase TI has rendered it a key enzyme in researches on the sequences of RNAs. Brownlee (105) has reviewed the literature to 1972, which began with the classic studies of Holley and his associates on tRNA in 1965 and extended to the sequence methods introduced by the Sanger laboratory for use with RNA labeled in vivo with 32P. Complete digestion with RNase T, yields oligonucleotides ending in guanosine 3’-phosphate; partial digestion is used to obtain information on the ordering of the segments. The methods for preparing TI digests summarized by Brownlee (105) are current. The procedures initiated by Szekely and Sanger (106) in 1969 for the use of enzymes to obtain in vitro 32P-labeledproducts for sequence analysis of RNAs that are not available in a biologically labeled form have been subject to review in 1979 in Silberklang et ul. (107). More rapid methods for sequencing RNA, which make use of the specificity of RNase T1 to identify the positions of G residues in the sequence, have been 104. Takahashi, K . (1970). J . Biochem. (Tokyo) 68, 659. 105. Brownlee, G. G . (1972). “Determination of Sequences in RNA” in “Laboratory Techniques in Biochemistry and Molecular Biology” Pocket edition of Part 1 of Vol. 3, pp. 1-265, North-HollandiAmerican Elsevier, Amsterdam. 106. Szekely, M . , and Sanger, F. (1969). JMB 43, 607. 107. Silberklang, M . , Gillum, A. M . , and RajBhandary, U. L. (1979). “Methods in Enzymology,” Vol. 59, p. 58.

464

KENJI TAKAHASHI AND STANFORD MOORE

developed by Simoncsits et ul. (108) and by Donis-Keller et ul. (109). Other recent examples of the use of the enzyme in sequence work include the studies of Fraser and Ziff (110) on RNA structures near poly(A) of adenovirus-2 mRNAs, of Davies et ul. ( 1 1 1 ) on large RNase Tl-resistant oligonucleotides from protamine mRNA, of Domdey and Gross (112) on RNA sequence determination in the nanogram range by a combination of in vitro labeling procedures, and of Robertson et ul. (113) on '2sI-labeled RNAs.

B. ADDITIONAL APPLICATIONS Although the widest use of RNase Tl has been in the sequence determination of RNA, the enzyme also has found use in other studies, as previously reviewed by Egami et ul. (65). The enzyme has been used to synthesize guanylylnucleosides, oligoguanylates, and other guanosine-containing oligonucleotides with (3'-5')-phosphodiester bonds (114 -121 ); the possible formation of 2'-5' linkages (122, 123) has been shown to be an artifact of isolation (124). 2',3'-cyclic GMP can be prepared as an intermediate by digestion of RNA with RNase TI, and 2',3'-cyclic IMP and 2',3'-cyclic XMP can be obtained by digestion of deaminated RNA (125). Various oligonucleotides 108. Simoncsits, A., Brownlee, G. G., Brown, R. S., Rubin, J. R., and Guilley, H. (1977). Nutrire (Loridon) 269, 833. 109. Donis-Keller, H., Maxam, A. M., and Gilbert, W. (1977). Nucleic Acids R e s . 4, 2527. 110. Fraser, N . , and Ziff, E . (1978). JMB 124, 27. 111. Davies, P. L., Dixon, G. H., Simoncsits, A., and Brownlee, G. G. (1979). Nucleic Acids Res. 7, 2323. 112. Domdey, H., and Gross, H. J. (1979). A d . Biochem. 93, 321. 113. Robertson, H. D., Dickson, E., Plotch, S. J., and Krug, R. M.,(1980). Nuckic Acids R e s . 8, 925. 114. Sato-Asano, K., and Egami, F. (1958). BBA 29, 655. 115. Scheit, K. K.,and Cramer, F. (1964). Tetruhedron Letr., p. 2765. 116. Sekiya, T., Furuichi, Y.. Yoshida, M.,and Ukita, T. (1968).J. Biochem. ( T o k y o ) 63, 514. 117. Grunberger D., Holy, A., and Sorm, F. (1968). Collection Czech. Chem. Commrrn. 33, 286. 118. Mohr, S. C., and Thach, R. E. (1969).JBC 244, 6566. 119. Rowe, M. J., and Smith, M. A. (1970). BBRC 38, 393. 120. Sato-Asano, K. (1960). J . Birrhem. ( T o k y o ) 48, 284. 121. Hayashi, H., and Egami, F. (1963). J. Biuchem. ( T o k y o ) 53, 176. 122. Podder, S. K., and Tinoco, I., Jr. (1969). BBRC 34, 569. 123. Podder, S. K. (1970). BBA 209, 455. 124. Omori, A., Yoshida, H., and Tamiya, N. (1974). J. Biochem. ( T o k y o ) 76, 117. 125. Sato-Asano, K., Fujii, Y., and Egami, F. (1959). Bull. Chem. Soc. Japan 32, 1068.

13. RIBONUCLEASE TI

465

ending in 2',3'-cyclic GMP or 3'-GMP may also be obtained by digestion of RNA. The enzyme may be used as a reagent for limited cleavage of an RNA chain in the structure-function studies of RNA. It may also be used to remove contaminating RNA from DNA or other biological samples.

V.

Other Guanine-Specific RNases

Several guanine-specific RNases like RNase T1 have been isolated from microorganisms, especially from fungi (see Ref. (2)). They include RNase U1 (Usfilago sphaerogena) (126-128), RNase N1 (Neurospora crassa) (129, 130), RNase Ch (Chalaropsis sp.) (131, 132), RNase from Aspergillus firmigutus (133), RNase F1 (Fusarium moniliforme (134, 135), RNase St (Streptornyces eryrlzreus ) (136), RNase Cz (Aspergillus clavutus ) (137), RNase I1 (an intracellular RNase of Aspergillus cluvarus) (138), RNase from Actinomyces aitreoverticillarus (139), RNase from Streptomyces aureofaciens (140 ), RNase from Penicillium brevicompactum (141), and RNase Pchl (Penicillium chrysogenum) (141). Like RNase T1, most of these guanyloribonucleases are extracellular enzymes and are generally thermostable, small proteins (MW 10,000-15,000), and specifically cleave the 3'-phosphodiester bonds via the corresponding 2' ,3'cyclic phosphates. The amino acid compositions of some of these RNases are shown in Table VI; the compositions resemble that of RNase T I , with the possible 126. Arima, T., Uchida, T., and Egami, F. (1968). BJ 106, 601. 127. Hashimoto, J., Uchida, T., and Egami, F. (1971). J . Biochem. (Tokyo) 70, 903. 128. Kenney, W. C., and Dekker, C. A. (1971). Biochemistry 10, 4962. 129. Takai, N . , Uchida, T., and Egami, F. (1966). BBA 128, 218. 130. Kasai, K . , Uchida, T., Egami, F., Yoshida, K . , and Nomoto, M. (1969).J. Biochem. (Tokyo) 66, 389. 131. Fletcher, P. L., Jr., and Hash, J. H. (1972). Biochemisfr-y 11, 4274. 132. Fletcher, P. L., Jr., and Hash, J. H. (1972). B i o c h r m i s f ~11, 4281. 133. Glitz, D. G., Angel, L., and Eichler, D. C. (1972). Biochemistry 11, 1746. 134. Omori, A , , Sato, S . , and Tamiya, N . (1972). BBA 268, 125. 135. Yoshida, H . , Fukuda, I . , and Hashiguchi, M. (1980).J. Biochem. (Tokyo),88, 1813. 136. Tanaka, K . (1961). J . Biochem. (Tokyo) 50, 62. 137. Morozova, V. G., Grishchenko, V. M., and Bezborodova, S. I. (1972). I z v . Akad. Nnrtk. SSSR Srr. B i d . p . 865. 138. Ivanova, G. S . , and Valiukaite, R. V. (1974). Mrkrobiologiya 43, 417. 139. Abrosimova-Amelyanchik, N . M., Tatarskaya, R. I . , Venkstern, T. V., Axelrod, V. D., and Baev, A. A. (1965). Biokhimiya 30, 1269. 140. Barova, M . , Zelinkova, E . , and Zelinka, J. (1971). BBA 235, 335. 141. Zhenodarova, S. M., Gulyaeva, V. I., and Bezborodova, S. I. (1976). Biaorg. Khim. 2, 1 1 1 1 .

TABLE VI THEAMINO ACIDCOMPOSITIONS OF SOME GUANYLORIBONUCLEASES" RESIDUESPER MOLECULE RNase from Amino acid

RNase T1 (14)

RNase U1 (127, 128)

RNase N1 ( 2)

RNase Ch (132)

15 6 15

16(15)

14 4 14 4 5 13 10 4 4 2

8 8 15 6 5 14

A . firmigatus (133)

RNase F, (135)

RNase C2 ( 144 )

RNase St (143)

15 10 10 8 6 14 12 4 5 0 3 0 8 4 0 1

11 3 13 6 4 14 8

11 8 3 14 8 13 5 2 6 0 3

~

Asp + Asn Thr Ser Glu + Gln pro G~Y Ala

KYS Val Met Ile Leu Tyr Phe TrP LY s His Ar&!

8 (9) 13 6 4 15 5 4 6

9 4 12

7 4 8 0 2 3

0 2 1 12 4 0 3 2 2

9 4 1 1 3 1

Total

103

104 ~~

Numbers in parenthesis are references. N.D., not determined.

5 4 9 5 1 3 3 3

9 4 5 1 3 5 10 6 3 4 3 1

107

110

14 5 12 8 5 11 11 4 6 0 3 3 10 4 1 1

N.D.* 4

2

2

2

2

0 2 4 10 3 1 1 3 3

102

104

90

5 9 5 0 2 2 6 102

13. RIBONUCLEASE Ti

467

exception of that of RNase St. A tentative amino acid sequence of RNase U, has been reported (142). The sequence is different from that of RNase T1 in about 60% of the total residues. However, it was found to contain the amino acid sequences corresponding to those in RNase T1involving His-40, His-92, Glu-58, and Arg-77, which have been implicated in the active site of RNase T,. The amino-terminal 18-residue sequence of RNase Ch has been reported (132) and shows a close homology with that of RNase T,. The amino acid sequence of RNase St has been determined (143); although the sequence is quite different from that of RNase T1, there is Arg at 77 and His at 92. Antibodies to RNase T I reacted weakly with Aspergillus fitmigutus RNase, but did not react with RNases U, and N1 (133). Iodoacetate or bromoacetate are known to inactivate RNase U, (127), RNase N l (127), Aspergillus fumigutus RNase (133), RNase Ch (132), RNase C2(144), and RNase Pch, (145). A 1 : 1 stoichiometry of the reaction was established for RNase U,, RNase Ch, and Aspergillus .furnigutus RNase. The residue in RNase U1 specifically modified by iodoacetate was shown to be a glutamic acid residue (146).Involvement of some histidine residues in the active site was demonstrated for RNase U1 ( / 4 7 ) ,RNase C2 (144), and RNase Pch, (145) by photooxidation studies. Further, implication of an arginine residue in the active site was suggested for RNase U,(146) and Asper~ilirrsfumi~ufIrs RNase (133)from the results of phenylglyoxal modification. UV absorption spectra of RNases U1 and N1 (127), CD spectra of Aspergillrw firmigatrrs RNase and its derivatives (133) and RNase St ( I d a ) , and the fluorescence spectrum of RNase C2 (149) have been reported. In the case of RNase St, the CD-spectral measurement was utilized to investigate the interaction between the enzyme and inhibitors. Substrate specificity has been investigated in comparison with RNase T, for RNase C2 (70, 141), RNase I1 (70), RNase from Actinomyces uureoverticillutus (701, RNase from S. uureoficiens (70), RNase from P. brevicornpucturn (141), and RNase Pch, (141). Kinetic studies have been performed with 142. Hashimoto, J., and Takahashi, K. (1974). J . Biochem. (Tokyo) 76, 1359. 143. Yoshida, N . , Sasaki, A., Rashid, M. A., and Otsuka, H. (1976). FEBS Lerr. 64, 122. 144. Grishchenko, V. M., Beletskaya, 0. P., and Bezborodova, S. I. (1975). Bioorg. Khim. 1, 1474. 145. Grishchenko, V. M., and Markelova, N. Yu. (1979). Biokhirniva 44, 1447. 146. Hashirnoto, J . , and Takahashi, K. (1977). J . Biochern. ( T o k y o ) 81, 1175. 147. Hashimoto, J . , Takahashi, K . , and Uchida, T. (1973). J . Biochem. ( T o k y o ) 73, 13. 148. Yoshida, N . , Kuriyama, K., Iwata, T., and Otsuka, H. (1971). BBRC 43, 954. 149. Grishchenko, V. M., Ernelyanenko, V. I . , Ivkova, M. N., Bezborodova, S. I . , and Burstein, E. A. (1976). Bioorg. Khim. 2, 207.

468

KENJI TAKAHASHI AND STANFORD MOORE

Aspergillus fumigcltus RNase (133), RNase St (62), and RNases U, and N, (127). Most of these guanyloribonucleases appear to be homologs of RNase TI with a similar active site characterized by the presence of a carboxyl group, presumably of a specific glutamic acid residue, reactive with iodoacetate or bromoacetate, and hence to function via similar mechanisms of substrate recognition and catalysis.

tRNA Processing Enzymes from Escherichia coli RYSZARD KOLE

SIDNEY ALTMAN

I . Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . 470 I1 . Ribonuclease P . . . . . . . . . . . . . . . . . . . . . . . . . 471 A . Introduction . . . . . . . . . . . . . . . . . . . . . . . . 471 B . Purification and Properties . . . . . . . . . . . . . . . . . . 471 C . Structure . . . . . . . . . . . . . . . . . . . . . . . . . . 472 473 D . Reaction Catalyzed . . . . . . . . . . . . . . . . . . . . . E . Biological Role . . . . . . . . . . . . . . . . . . . . . . . 475 F. Ribonuclease P from Other Organisms . . . . . . . . . . . . . 475 111. Ribonuclease 111 . . . . . . . . . . . . . . . . . . . . . . . . 476 A . Introduction . . . . . . . . . . . . . . . . . . . . . . . . . 476 B . Biological Role . . . . . . . . . . . . . . . . . . . . . . . 476 IV. Ribonuclease P2and Ribonuclease 0 . . . . . . . . . . . . . . . 477 A . Introduction . . . . . . . . . . . . . . . . . . . . . . . . 477 477 B . Purification and Properties . . . . . . . . . . . . . . . . . . C . Biological Role . . . . . . . . . . . . . . . . . . . . . . . 478 V. Ribonuclease D . . . . . . . . . . . . . . . . . . . . . . . . 479 A . Introduction . . . . . . . . . . . . . . . . . . . . . . . . 479 480 B . Purification and Properties . . . . . . . . . . . . . . . . . . 480 C . Reaction Catalyzed . . . . . . . . . . . . . . . . . . . . . D . Biological Role . . . . . . . . . . . . . . . . . . . . . . . 482 VI . Other Nucleases . . . . . . . . . . . . . . . . . . . . . . . . 482 VII . Concluding Remarks . . . . . . . . . . . . . . . . . . . . . . 483 469 THE ENZYMES. VOL . XV Copyright @ !, 1982 by Academic Press. Inc . All rights of reproduction in any form reserved . ISBN 0- 12- I227 13-4

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FIG. 1. Schematic illustration of the interaction of tRNA processing enzymes with tRNA precursor molecules in E . coli: The upper part of the sketch shows potential sites of action of various enzymatic activities (see text) on either mono- or polycistronic tRNA precursor molecules. Endoribonuclease action is indicated by an arrow perpendicular to the line showing the gene transcript, and exoribonuclease action by a parallel line. The lower part of the figure illustrates enzyme action on the 30 S transcript of a rRNA cistron isolated from RNase 111- cells. Events (and enzymatic activities) are shown that have been identified using the particular gene transcript portrayed in the figure. More events may exist than those shown, and some of the enzymatic activities indicated may be identical to each other.

I.

Introduction

The primary transcript of a tRNA gene undergoes several enzymecatalyzed transformations before it becomes a functioning tRNA molecule. The steps involved in the maturation of prokaryotic tRNA molecules are now quite well-defined. The known or suspected processing steps and the enzymatic activities involved are shown in Fig. 1. [For detailed treatments of tRNA biosynthesis several reviews can be consulted, Refs. (/5)l. Depending on its chromosomal environment, a tRNA gene is transcribed as either a monomeric or multimeric precursor molecule that is rapidly processed by both endo- and exonucleases. Further processing steps involve enzymatic modification of nucleotides and the addition of terminal CCA residues to processed precursor molecules. Some tRNA genes are cotranscribed with rRNA genes and thus the mature tRNA molecules must be excised from the long transcription unit (Fig. 1). Our understanding of tRNA processing in eukaryotes is less detailed than our knowledge of the processes in prokaryotes, but the general features of tRNA synthesis seem to be similar in all organisms. One impor1. Sbll, D., Abelson, J. N., Schimmel, P. R. (eds.) (1980). “Transfer RNA: Biological Aspects’’ Cold Spring Harbor Laboratory, Cold Spring Harbor, New York. 2. Altman, S. (1978). Intern. R e v . Biocliem. 17, 19. 3. Altman, S. (1981). Cell 23, 3. 4. Abelson, J. (1979). Annu. Re\*. Biochem. 48, 1035.

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47 1

tant difference between eukaryotes and prokaryotes is that some tRNA genes contain an intervening sequence that is transcribed into the tRNA precursor molecule and must then be spliced out during maturation of the precursor. Enzymes responsible for the splicing of tRNA precursors have been identified in yeast extracts and Xenopiprts lnevis extracts. In this chapter we discuss only the endo- and exonucleases that alter the size of tRNA gene transcripts. Other tRNA processing enzymes are discussed elsewhere in this volume. The emphasis of our discussion is on enzymes identified in extracts of E. coli since some of these have been very well-characterized; the same cannot be said for most eukaryotic tRNA processing activities. The data showing the involvement of some of the E. coli enzymes in tRNA processing (e.g., for RNases P, D, 111, and P4)are much stronger than they are for others (e.g., RNases Pz,O,Q,PIII, Y, and II), which have not been as well-characterized. II.

A.

Ribonuclease P

INTRODUCTION

RNase P is an endoribonuclease that generates the 5’ termini of all mature tRNAs in E. coli. An essential role for this enzyme in the biosynthesis of tRNA in E. coli has been demonstrated through the use of various mutants that affect the enzymatic activity. RNase P has an unusual structure in that it is a ribonucleoprotein. The active enzyme is made up of both protein and RNA. RNase P-like activity has been detected in several organisms, and in every case its properties are similar to those of RNase P from E. coli. B.

P U R I F I C A T I O N A N D PROPERTIES

Initial attempts to characterize RNase P (6-8) met with great difficulty, which in retrospect was due to the ribonucleoprotein nature of this enzyme. A total purification of both the protein and RNA moieties has been achieved (9, 9 a ) . The starting material for this purification scheme is an 5. Shimura, Y., and Sakano, H. (1977). I n “Nucleic Acid Protein Recognition” (H. J. Vogel, ed.), p. 293. Academic Press, New York. 6. Robertson, H . D., Altman, S . , and Smith, J . D. (1972). JBC 247, 5243. 7. Bikoff, E. K . , and Gefter, M. L. (1975). JBC 250, 6240; Bikoff, E. K . . LaRue, B. F., and Gefter, M . L. (1975). ihid p. 6248. 8. Guthrie, C., and Atchison, R. (1980). I n “Transfer RNA: Biological Aspects” (D. Soll, J. N . Abelson, and P. R . Schimmel, eds.), p. 83. Cold Spring Harbor Laboratory, Cold Spring Harbor, New York. 9. Kole, R . , and Altman, S. (1981). Biochernistr.y. 20, 1902. 9a. Baer, M . Personal communication.

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extract of E. coli prepared by grinding the cells with alumina. The crude extract is then chromatographed on DEAE-Sephadex using stepwise elution. [The assay for RNase P activity involves incubating subcellular fractions or column fractions with a 32P-labeledtRNA precursor molecule (6). Cleavage reaction products are then analyzed by polyacrylamide gel electrophoresis.] The fractions containing the peak of activity from the DEAE-Sephadex column are then passed over a Sepharose-4B column and the resulting peak of RNase P activity is then passed over a Sephadex G-200 column. As might be expected of a complex containing a nucleic acid, RNase P elutes from the DEAE-Sephadex column between 0.4 and 0.5 M NH,Cl. The Sepharose 4B column removes most of the contaminating rRNA and the Sephadex G-200 column removes most of the contaminating tRNA. Further purification of RNase P is achieved by chromatography on two consecutive n -octyl-Sepharose columns. The purest fractions resulting from such a scheme still contain at least two protein species, as determined by polyacrylamide gel electrophoresis, and two major RNA species as well as some smaller RNA breakdown products. Total purification of the RNase P polypeptide protein is achieved by chromatography of material from the scheme described above on CMSephadex column in the presence of 7 M urea, which dissociates the nucleic acid from the protein. In such a chromatography step the RNA is collected in the flow-through fractions but the protein is retarded and can then be eluted with a salt gradient. None of the fractions collected from the CM-Sephadex column, either in the presence of urea or when dialyzed against the appropriate buffer, show any enzymatic activity. However when one particular protein fraction, designated C5, is mixed in the urea buffer with an RNA species, designated M1 RNA [which has been previously purified by successive gel electrophoresis, (Ref. 9u, l o ) ] ,and the urea is dialyzed away from this mixture, the protein and RNA apparently come together to form an active enzyme complex. Thus RNase P can be reconstituted with a high yield from separated RNA and protein components (9, 10). C. STRUCTURE The two components necessary for reconstitution of active RNase P (9) are the C5 protein (molecular weight 17,500) and the M1 RNA (molecular weight 120,000, about 360 nucleotides). If active RNase P is pretreated with either a ribonuclease or a protease, enzymatic activity is

-

-

10. Kole, R., and Altman, S . (1979). PNAS 76, 3795.

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abolished (11). This functional test also demonstrates that the enzyme requires both an RNA and a protein component for its activity. Furthermore the bouyant density ofE. coli RNase P in CsCl is 1.71 g/cm3(12), and in Cs2S04it is 1.55 g/cm3 (13), thus manifesting again the properties of a ribonucleoprotein complex. The buoyant density results are consistent with a hypothetical composition of an enzyme with 1 M1 RNA molecule and 1 C5 protein molecule. However it is not possible to say whether the active enzyme complex contains one or more copies of each type of subunit in stoichiometric proportions. No RNA species other than M1 RNA has yet been found to yield active RNase P in a reconstitution reaction with C5 protein. Similarly no protein has been found yet that can substitute for the C5 protein in the reconstitution reaction (9, 10). Studies of temperature-sensitive mutants ofE. coli have shown that one such mutant affects the protein component of RNase P and another mutant (which maps at a different locus on the E. coli genetic map) affects the RNA component of the enzyme. In particular the metabolism of M1 RNA is abnormal in the second type of mutant (14). These observations lend support to the notion that the enzyme functions as an RNA-protein complex in vivo as well as in vitro,

D. REACTION CATALYZED The 5‘ termini of all mature tRNA molecules begin with a phosphate group. There is no sequence homology at the 5’ termini of tRNA molecules. Nucleotide sequence analysis of the products generated by RNase P action on the precursor to E. coli tRNATyror other tRNA percursors from various sources has shown that in every case the RNase P activity generates the correct 5’ terminus of the mature tRNA sequence. This statement, for example, applies to the action of E. coli RNase P on E. coli tRNA precursors, or to the action of eucaryotic RNase P on E. coli tRNA precursors, or to the action on their homologous precursors [reviewed in Refs. (2) and ( 3 1 . Thus RNase P is an endoribonuclease that produces 5’-phosphate groups and 3’-hydroxyl groups and does not recognize nucleotide sequences around its cleavage sites. Very few nucleotides are needed on the 5’ side of an RNase P cleavage site for the enzyme to 1 1 . Stark, B. C., Kole, R., Bowman, E. J . , and Altman, S. (1978). P N A S 75, 3717. 12. Altman, S . , Bowman, E. J . , Garber, R. L., Kole, R., Koski, R. A. and Stark, B. C. (1980).In “Transfer RNA: Biological Aspects” (D. SOU, J. N. Abelson, and P.R. Schimmel, eds.), p. 71. Cold Spring Harbor Laboratory, Cold Spring Harbor, New York. 13. Akaboshi, E., Guerrier-Takada. C., and Altman, S. (1980). BBRC 96, 831. 14. Kole, R . , Baer, M. F., and Altman, S. (1980). Ceil 19, 881.

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function. Indeed a precursor with only three extra nucleotides has been found in E. coli. Mutants in tRNA precursors that affect the ability of RNase P to cleave these precursors have been studied extensively in E. coli and bacteriophage T4. These mutants invariably disrupt the tertiary and/or secondary structure of the tRNA precursors in solution [discussed in detail in Ref. (2)l. It is, therefore, thought that RNase P recognizes the common tRNA conformation of its substrate molecules in solution as a step in finding the appropriate cleavage site on all its substrates. In this way the enzyme need not look for common nucleotide sequences near its cleavage sites. On another level of detail, all tRNA molecules have invariant nucleotides at certain positions in their sequences. These nucleotides are thought to be critical in determining the conformation of the tRNA molecules in the solution. Thus it may be hypothesized that M1 RNA, the RNA component of the enzyme, interacts through hydrogenbonding schemes with the invariant nucleotides in the substrates to facilitate correct enzyme orientation on its substrates (10, 12). This hypothesis remains to be checked. It is important to note that RNase P does not act with the same efficiency on all precursor substrates. The reasons for the differences in enzyme rates with different substrates are not known, but nucleotide modification, which may affect the details of the tRNA moiety conformation, may play a role in enzyme substrate rates. Synthetic precursor molecules have been constructed by using RNA ligase to join a radioactive tetranucleotide to the 5' end of a mature tRNA molecule. Such synthetic substrates provide an assay for RNase P activity based on acid solubilization of radioactivity which is less cumbersome than gel analysis. (S. Nishikawa and D. Soill, personal communication). In addition to tRNA precursors, highly purified E. cob RNase P can cleave in vitro a precursor to 4.5 S RNA. This precursor accumulates at high temperatures in RNase P thermosensitive mutants of E. Cali . A biological role for this RNA is not known. The rate of cleavage is less than 10% of the precursor to tRNATYrand thus may be a secondary reaction to RNase P (9). Several other RNA molecules besides tRNA precursors have been tried as substrates for RNase P but none have been found to function in this manner. For example, rRNA, tRNA, various messenger RNAs and single-stranded RNAs fail to function as RNase P substrates (9, 11).

The requirements for the RNase P reaction, regardless of the source of the enzyme, are similar. That is, the enzyme requires magnesium ion and a monovalent cation (NH,+, K+, Na+). The pH optimum is near 8. The reaction is inhibited by tRNA or rRNA (9, 11, 13, 15, 16). TheK,for tRNA 15. Garber, R. L., and Altman, S. (1979). Cell 17, 389. 16. Bowman, E. J . , and Altman, S. (1980). BBA 613, 439.

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is about 10-6 M while the K ,,for the reaction with tRNATyras substrate is about M (164.

E. BIOLOGICAL ROLE In E. coli mutants that have thermosensitive RNase P function, precursors to forty o r more tRNAs accumulate at restrictive temperatures. If some of these precursors are exposed to RNase P in vitro, products are generated that comigrate with mature tRNA species in electrophoresis experiments, and/or have been shown by nucleotide sequence analysis to be identical to the corresponding mature tRNAs. This statement is restricted to the maturation of the 5‘ termini of tRNA molecules. Multimeric precursors are also found that can then be cleaved by RNase P, and possibly other enzymes, to generate either monomeric precursors, which still have extra 3’-nucleotides, or the fully mature tRNA molecules [reviewed in Refs. (2, 5 , 1 7 ) ] . It is clear, however, that the role of RNase P in generating the 5‘ termini of mature tRNA molecules is an essential one.

F. RIBONUCLEASE P FROM OTHERORGANISMS RNase P-like activities can be detected in extracts of several organisms by simply exposing the extracts to a 32P-labeledtRNA precursor from E. coli. In fact, it has been found that RNase P-like activity from any organism thus tested can make the precise, required cleavage in E. coli tRNA precursors as well as in homologous precursors, where they are available. RNase P-like activities have been identified in extracts of B. subtilis (18), and in many eukaryotic extracts; for example, in human tissue culture cells (19, 20), veal heart (13), chick embryonic tissue (161, green monkey kidney cells (21),yeast ( 2 / a ) , X . luevis (22),and silk worms (15). It is not surprising that substrate specificity of the enzyme is conserved throughout evolution because the distinctive feature of all the substrates appears to be the conformation of the tRNA moiety of the tRNA 16a. Stark, B. C . , PhD. thesis, Yale University, New Haven, Connecticut. 17. Shimura, Y., Sakano, H., Kubokawa, S., Nagawa, F., and Ozeki, H. (1980). I n “Transfer RNA: Biological Aspects” (D. SOU, J. N. Abelson, and P. R. Schimmel, eds.), p. 43. Cold Spring Harbor Laboratory, Cold Spring Harbor, New York. 18. Gardiner, K., and Pace, N. R. (1980). JBC 255, 7507. 19. Ferrari, S., Yehle, C. D., Robertson, H. D., and Dickson, E. (1980). PNAS 77 2395. 20. Koski, R. A , , Bothwell, A. L. M., and Altman, S. (1976). Cell 9, 101. 21. Altman, S . , and Robertson, H. D. (1973). Molec. Cell Biochern. 1, 83. 21a. Kline, L., Nishikawa, S. , and Soll, D. (1981). JBC 256, 5058. 22. Cortese, R., Melton, D., Tranquilla, T., and Smith, J. D. (1980). I n “Transfer RNA: Biological Aspects” (D. Soll, J. N. Abelson, and P. R. Schimmel, eds.), p. 287. Cold Spring Harbor Laboratory, Cold Spring Harbor, New York.

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precursor molecules. It is very likely that this structure is conserved throughout evolution. In addition, the ribonucleoprotein nature of the RNase P complex also seems to have been conserved throughout evolution. Reconstitution of RNase P from RNA and protein subunits has been demonstrated with E . subtilis RNase P (18), and functional inactivation of RNase P by pretreatment with ribonuclease and protease has been demonstrated with activities from veal heart ( l 3 ) , human tissue culture cells (20), and yeast ( 2 1 ~ )The . bouyant densities of RNase P from the mammalian tissues has also been shown in Cs2S04gradients to be intermediate between that of protein and RNA (13). The similarities in substrate specificity, ion requirements, and enzyme structure in all RNase P activities studied indicate that the role this enzyme has in tRNA biosynthesis is a general one in all organisms. 111.

Ribonuclease 111

A. INTRODUCTION Escherichia coli RNase I11 is known to play a role in the processing of rRNA transcripts in E. coli. Since some tRNAs are cotranscribed with rRNAs, it seems likely that RNase I11 plays an indirect role in the maturation of tRNAs also. In this section we describe the evidence that RNase I11 is involved in the processing ofE. coli tRNAs. RNase 111-like activities have been detected in eukaryotes but there is no evidence yet that this enzyme participates in the biosynthesis of eukaryotic tRNAs (The biological role, purification, and characterization of RNase I11 is described in Chapter 15, this volume.)

B. BIOLOGICAL ROLE In wild-type E. coli. RNase I11 participates in the processing of 16 and 23 S rRNAs. It may, therefore, be expected that tRNA sequences that are cotranscribed with the ribosomal RNAs would be released in large fragments from a polycistronic transcript from rRNA gene regions of the E. coli chromosome. In fact, Lund et al. (23) and Apirion et a f . (24) have shown that this is the case. In a strain of E. coli that lacks RNase I11 activity, a 30 S transcript accumulates that contains all the ribosomal 23. Lund, E., Dahlberg, J. E., and Guthrie, C. (1980). In “Transfer RNA: Biological Aspects” (D. Son, J. N. Abelson, and P. R. Schimrnell, eds.), p. 123. Cold Spring Harbor Laboratory, Cold Spring Harbor, New York. 24. Apirion, D . , Ghora, B. K., Plautz, G., Misra, T. K., and Gegenheimer, P. (1980). In “Transfer RNA: Biological Aspects” (D. 5611, J. N . Abelson, and P. R. Schimmel, eds.), p. 139. Cold Spring Harbor Laboratory, Cold Spring Harbor, New York.

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RNA sequences and the “spacer” tRNAs. Purified RNase 111 can cleave this large precursor in v i m to generate smaller rRNA and tRNA precursor molecules that contain extra sequences both at the 5’ and 3’ termini of their sequences. The tRNA precursors can then be treated with RNase P to generate the mature 5’ termini of the tRNA molecules. It is important to point out that the RNase 111 strains of E. coli are viable and thus RNase I11 is not essential for the maturation of either these particular tRNA molecules or rRNA. There appear to be other endonucleases in E. coli that can assume the role RNase I11 normally plays in wild-type cells. A direct role for RNase I11 in the biosynthesis of bacteriophage T4 tRNAs has been described by McClain (25). The function of a particular suppressor tRNA is reduced at least 1W-fold in an RNase 111- strain of E. coli when compared to the wild-type. The synthesis of other TCencoded tRNAs is not affected by the RNase I11 mutation. The specific role that RNase I11 plays in the synthesis of this particular T4 tRNA has not been elucidated. IV.

Ribonuclease P2 and Ribonuclease 0

A.

INTRODUCTION

Crude extracts of E. coli manifest endoribonuclease activities that are capable of cleaving multimeric tRNA precursors (that is precursors containing more than one tRNA sequence) in the regions between the mature tRNA sequences. These partially processed precursors, which may then be processed to monomeric precursors that contain extra nucleotides at their 5‘ and 3‘ termini, are presumably further processed by enzymes like RNase P and an exonuclease acting at the 3‘ terminus of a precursor molecule. The endoribonuclease activities responsible for the intercistronic cleavages have been studied by two groups and named RNase P2 and RNase 0. Since both enzymes are not very well characterized, and different substrates have been used in the studies by each group, it is not clear whether the two activities represent the same o r different enzymes. Data suggest that RNase 0 maybe identical to RNase 111, whereas RNase P2may be distinct from RNase 111. B.

P U R I F I C A T IAONND PROPERTIES

1. RNase P 2 (26) RNase P2has been identified in 0.5 M NH4Cl washes of ribosomes in crude extracts of E. coli. The activity has been further purified by 25. McClain, W. H. (1979). BBRC 86, 718. 26. Schedl, P., Roberts, J . , and Primakoff,P. (1976). Cell 8, 581.

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chromatography on DEAE-Sephadex and phosphocellulose columns. At this stage of purification RNase Pzis free of RNase P, RNase 111, and 3' exonuclease activity. This enzyme (as does RNase 0, RNase 111, and RNase P) produces 5'-phosphate and 3'-hydroxyl groups in intercistronic regions in tRNA precursor molecules. Its complete substrate range has not been described.

2. RNnse 0 (5, 27) RNase 0 activity can be detected in the ribosome pellet of crude extracts of E. coli and in the SlOO supernatant of the ribosome pellet. The S 100 material has been passed through successive DEAE-cellulose columns and overa hydroxylapatite column. The resulting activity is very unstable. The enzyme has a broad pH range, 7.5-10, but is inactive below pH 7. It is stimulated by magnesium and manganese ions but is inhibited by monovalent cations at a concentration of 0.1 M , and by double-stranded RNA. This last characteristic is also shared by RNase 111. Although RNase I11 primarily cuts double-stranded RNA substrates, it is known to be able to cleave single-stranded RNA under low salt conditions. Thus it is not entirely clear that RNase 0 activity is free of RNase 111.

C. BIOLOGICAL ROLE Since no known mutants of E. coli affect the activity of either RNase Pz or RNase 0, it is difficult to establish the true role of these enzymes in the in vivo metabolism of tRNA. However, some data indicate that these enzymes may be responsible for early processing steps in the maturation of tRNA sequences contained in polycistronic transcripts. Under restrictive temperature conditions, no mature tRNA sequences are detectable during pulse-labeling in E. coli mutants thermosensitive for RNase P. Many tRNA precursors accumulate under these conditions but multimeric precursors appear to be less stable than monomeric precursors. This indicates the presence of a nucleolytic activity that is not inactivated under the restrictive conditions. Many of the monomeric precursors have ribonucleoside monophosphates at their 5' termini, suggesting that they are not primary transcripts but products of processing reactions. It is presumed that the nucleolytic activity responsible for these early processing reactions may be either RNase 0 or RNase Pz.In fact, when some multimeric precursors are incubated with partially purified RNase 0 or a crude extract of E. coli, products the same size as some monomeric precursors are generated. These, in turn, can be further processed by RNase P and 3'-exonucleases. Some dimeric tRNA precursors are resis27. Shimura, Y., Sakano, H., and Nagawa, F. (1978). EJB 86, 267.

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tant to both RNase 0 and RNase P, activity but can be processed by RNase P. From these data it has been concluded that some precursors are processed in a necessary order that may be RNase P action prior to RNase 0 or P2 action, or in some cases RNase 0 or RNase P, action prior to RNase P action. All these events have been observed only in v i m (5, 17, 28). Thus comments about the relevance of these ordered reactions to in \ j i t ’ o events are purely speculative. There is conflicting evidence regarding the distinction, or lack of it, of RNase 0 from RNase Pz.The in vitro reactions these enzymes carry out appear to be very similar. However, crude extracts of an E. coli strain that lack RNase I11 activity have no activity against multimeric tRNA precursors. This observation suggests that RNase 0 and RNase I11 may be related ( 5 ) . In an E. coli strain that both lacks RNase I11 and is temperaturesensitive for RNase P activity, under restrictive conditions, some cleavage is still observed of the large 30 S transcript of ribosomal RNA cistrons and some tRNA precursors (29). Thus a residual nucleolytic activity that is capable of producing smaller tRNA precursors is present in the strain. These products resemble, in gel electrophoretic mobility, products produced by RNase p,. Since no detailed information is available concerning the precise nucleotide sequences of the products, and the characterization of RNase Pzis not complete, further experimentation is necessary to show whether or not RNase Piis indeed distinct from RNase 111. V.

Ribonucleare D

A.

INTRODUCTION

Most tRNA precursors that accumulate in RNase P temperaturesensitive strains of E. coli carry extra nucleotides at both their 5‘ and 3‘ termini. Studies in vitro with crude extracts of E . coli indicate that 3’nucleotides are removed by an exonuclease activity (17, 26, 27). Similar results have been found in cases where eukaryotic tRNA precursors with extra 3’-nucleotides have been studied (15, 20). There are several constraints on the properties of the 3’-exonuclease that could be involved in the processing of the 3’ termini of tRNA precursors [reviewed in Ref. (XI)]. It has been shown that E.coli mutants defec28. Sakano, H . , and Shimura, Y. (1978). J M B 123, 287. 29. Robertson, H. D., Pelle, E. G . and McClain, W. H. (1980). In “Transfer RNA: Biological Aspects” (D. SOU, J. N . Abelson, and P. R. Schimmel, eds.), p. 107. Cold Spring Harbor Laboratory, Cold Spring Harbor, New York. 30. Chosh, R. K. and Deutscher, M. P. (1980). In ”Transfer RNA: Biological Aspects” (D. SOH, J. N . Abelson, and P. R. Schimmel, eds.), p. 59. Cold Spring Harbor Laboratory, Cold Spring Harbor, New York.

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tive in tRNA nucleotidyltransferase are not defective in tRNA biosynthesis. Therefore the nuclease that removes the extra residues must stop at the CCA sequence common to all tRNAs. The processed tRNA molecules can then be aminoacylated. (Eukaryotic and some T4 tRNA genes do not have CCA sequences encoded. Thus extra nucleotides must be removed to the point where nucleotides found in the mature sequence start and the CCA sequence must then be added.) The order of action of nucleases on tRNA precursors, that is whether or not the 3' terminus or the 5' terminus is matured first, seems to depend upon the particular tRNA sequence involved. However, using data from in vivo experiments, one can make predictions about the mode of action of 3'-nucleases on particular tRNA precursor molecules. Several 3'-exonucleases have been described in the literature and implicated in the processing of tRNA: RNase D (SO, S l ) , RNase I1 (26), RNase Q, RNase Y (5, 17), RNase PI (7), and a nuclease found in a strain called E. coli BN (32). RNase D, which has been extremely well characterized, seems to have the features necessary for the tRNA processing enzyme, whereas some of the other enzymes may be either identical to RNase D, or lack some of the features one would wish to see in the canonical 3'-exonuclease. B. PURIFICATION A N D PROPERTIES (31, 33-34a) An ,3100 extract of E. coli has been chromatographed on two consecutive DEAE-Sephadex columns and hydroxylapatite. At this stage of purification RNase D is completely devoid of RNase I1 activity. Further purification steps include gel filtration on ultragel AcA44 (Biorad) and chromatography on Affi-Gel Blue. This procedure yields homogeneous enzyme as judged by staining of protein bands in polyacrylamide gels. The molecular weight of RNase D is about 40,000. The enzyme has a pH optimum between 9 and 10 and requires magnesium ion for activity. Monovalent cations at concentrations up to 0.1 M have only minor effects on activity. C. REACTION CATALYZED Three types of substrates were used for in vitro assays of RNase D. On all three substrates RNase D acts in an exonucleolytic fashion, releasing 31. Cudny, H . , and Deutscher, M. P. (1980). PNAS 77, 837. 32. Seidman, J. G., Schmidt, F. J . , Foss, K., and McClain, W. H. (1975). Cell 5, 389. 33. Ghosh, R . K . , and Deutscher, M. P.(1978). JBC 253, 997. 34. Ghosh, R. K . , and Deutscher, M. P. (1978). Ntrclric Acids Res 5, 3831. 34a. Cudny, H. personal communication.

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48 1

5'-ribonucleotides. Three of the substrates are synthetic, and have been made by incubating bulk tRNA with rabbit liver tRNA nucleotidyltransferase in the presence of labeled ribonucleoside triphosphates. The substrates obtained have the structure tRNA-CU, tRNA-CCA, and tRNA-CCACC with the last nucleotide labeled with I4C. The assay for RNase D consists of measuring the amount of released acid-soluble radioactivity (30.35).A fourth substrate used to assay RNase D activity is the E. coli tRNATYrprecursor 32P-labeledin vivo (36). All four substrates including tRNA with a mature CCA terminus were hydrolyzed by RNase D, indicating that the enzyme has no absolute sequence specificity. The rate of hydrolysis of mature tRNA (tRNA-CCA) is less than 3% of that for tRNA with extra nucleotides at the 3' terminus. The hydrolysis of tRNA-CU (that is, with the CCA sequence absent) is even slower (30). These properties are to be expected of an enzyme involved in tRNA maturation in viva . When synthetic tRNA precursor, tRNA-CCACC, was treated with RNase D and simultaneously assayed for its amino acid-acceptor activity, about 60% of acceptor activity was regenerated compared to the original tRNA-CCA. Under these conditions about 80% of the extra residues have been removed. In contrast, when purified RNase I1 is tested in the same system, no restoration of the amino acid acceptor activity is observed (31). It is thought therefore that RNase D stops its action at the CCA sequence (or slows down considerably), whereas RNase I1 proceeds into the mature tRNA sequence. RNase D can also hydrolyze the precursor to tRNATY'(with extra nucleotides present at both termini). The action of the enzyme is much more efficient when RNase P is also present to remove the extra 5' nucleotides from the precursor. From this natural precursor, RNase D generates a product containing the mature tRNA sequence but with lower efficiency than that observed for synthetic precursors (S. Altman, unpublished). It is interesting to note that the tRNATYrprecursor which accumulates in RNase P thermosensitive mutants has extra nucleotides at both ends (36). This seems to indicate that in vivo RNase P action is required before efficient removal of nucleotides at the 3' end can occur, just as has been observed in virro . RNase D can also hydrolyze tRNA with altered structures. For example, tRNA treated with phosphatase and snake venom phosphodiesterase (the 5'-phosphate and several nucleotides from the 3' terminus are removed) is hydrolyzed by RNase D about idfold faster than intact tRNA. Similarly, heat-denatured tRNA is more susceptible to RNase D action 35. Deutscher, M. P., and Ghosh, R. K. (1978). Nirc./eic Acids Res. 5 , 3821. 36. Altrnan, S . (1971). Nrrrrrre Neil, B i d . 229, 19.

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RYSZARD KOLE AND SIDNEY ALTMAN

than intact tRNA. Poly(A) * poly(U), E. coli ribosomal RNA, and doublestranded poly(A) . poly(U) are all resistant to RNase D action (33). However, RNase D degrades the 5' terminal fragment released from tRNATYr precursor by RNase P action (30).

D. BIOLOGICAL ROLE Although lethal mutants of E. coli that lack RNase D activity have not been found, the characteristics of this enzyme found in in vitro experiments make it a good candidate for a necessary 3' processing exonuclease in tRNA biosynthesis. The lack of absolute specificity of RNase D i n v i m (that is, its small but detectable rate of activity against CCA sequences) may not be relevant to its in vivo function. In vivo it is possible that after tRNA is processed by RNase D, it is rapidly aminoacylated and rendered resistant to further degradation. VI.

Other Nucleases

In addition to the enzymes described above, several other activities have been observed in the processing of tRNA precursor molecules in vitro. However, it is difficult to assess the properties and biological function of many of these poorly characterized enzymes. It is possible that some of them are identical to the enzymes we have already described. In addition to RNase D, five different activities have been implicated in processing at the 3' termini of tRNA molecules: RNase Q, RNase Y (5, l 7 ) , RNase I1 (26), RNase P3 (7), and an RNase activity that is absent from E. coli BN (32). It is not known whether RNase P3 and the BN enzyme are the endo- or exonucleolytic activities. Escherichiu coli BN is viable but defective in the processing of certain T4-coded tRNA precursors. Data indicate (360) that E. coli BN may contain a mutant form of RNase D. Both RNase Q and RNase P3 remove extra nucleotides from the 3' termini of tRNA precursors in vitro. However, these enzymes are insufficiently characterized to say whether or not they are identical to, or distinct from, RNase D or each other. Although RNase I1 copurifies with RNase D through several steps of various purifications schemes, it can be finally separated from RNase D by chromatography on DEAE-Sephadex and hydroxlapatite columns. RNase I1 does not appear to stop its exonucleolytic action when it reaches the CCA sequence of tRNA precursors (31). So this enzyme and a similar one, RNase Y, appear not to have essential roles in tRNA biosynthesis. In addition to RNase 0, RNase Pz,and RNase 111, other activities that 36a. Deutscher, M. P., personal communication.

14. tRNA PROCESSING ENZYMES FROM E . co/i

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are endoribonucleolytic have been described that may participate in intercistronic processing of tRNA precursors. They are RNase P4(37), fraction IV (7), RNase F, and RNase E (24,38).RNase P4is capable of cleaving an in vitro-generated precursor to E. coli tRNATYr,about 8 nucleotides downstream from the CCA sequence. Such an endonucleolytic event seems to be also necessary in vivo to generate a tRNATYrprecursor and function from a synthetic tRNATYrsuppressor gene (37). Thus a strong case can be made for the necessary involvement of RNase P4 in the biosynthesis of at least tRNATYrin E. coli. Whether or not it is involved in the processing of other tRNA precursors near their 3' termini is not known. The biochemical characteristics of RNase P4and the other nucleases mentioned in this paragraph are not sufficiently well known to determine whether any of these activities are identical to each other or to some of the other endoribonucleases mentioned here. VII.

Concluding Remarks

While the general scheme of tRNA biosynthesis is reasonably well understood for all organisms, the particular enzymes involved in the processing events have not been well-characterized. This is mainly because of the difficulty in isolating sufficient amounts of natural substrates for extensive enzymatic studies. Furthermore, it seems that some of these enzymes may be present in vivo at relatively low levels. The use of synthetic substrates (for example, the synthetic precursors used in the studies of RNase P and RNase D) may facilitate future studies of enzymatic mechanisms. The most significant question to be answered in the study of tRNA processing enzymes is the nature of substrate recognition mechanisms. That is, how do they recognize their substrates with such exquisite precision that they never make errors in their cleavage reactions? We can already guess that in most cases the enzymes recognize conformational features of their substrates rather than details of nucleotide sequences or the characteristics of specific nucleotides. ACKNOWLEDGMENTS We thank many of our colleagues for providing reprints and preprints of their work and for interesting discussions. In particular, we acknowledge the help of the members of our laboratory. Work performed in the laboratory of S.A. was supported by U.S. Public Health Services Grant GM-19422 and National Science Foundation Grant PCM 79-04054. 37. Sekiya, T., Contreras, R . , Takeya, T., and Khorana, H. G. (1979). JBC 254, 5802. 38. Ghora, B. K., and Apirion, D. (1979). JBC 254, 1951; Misra, T. K. and Apirion, D. (1979). ihid.. 11154.

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Ribonuclease III JOHN J. DUNN

I . Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . 11. Purification . . . . . . . . . . . . . . . . . . . . . . . . . . 111. Structure . . . . . . . . . . . . . . . . . . . . . . . . . . . IV. Cleavage o f Double-Stranded RNA . . . . . . . . . . . . . . . .

V. Cleavage of Single-Stranded RNA A. Primary Sites . . . . . . . . B . Secondary Sites . . . . . . . VI. Related Eukaryotic Enzymes . .

I.

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

485 486 487 489 490 491 495 497

Introduction

RNase I11 ofEscherichirr coli is the endonuclease responsible for the first steps in the post-transcriptional processing of E. coli ribosomal RNA ( I -3). Ordinarily, cutting of the primary transcript is very rapidin viva and closely coupled with transcription, and the uncut precursor RNA, termed 30 S preribosomal RNA, is not observed in cells that have normal levels of RNase 111. RNase I11 was first characterized as an endoribonuclease that specifically degrades double-stranded RNAs of either natural or synthetic origin (4). Its role as a processing enzyme was discovered when it was found that extracts of E. coli contain an endoribonuclease capable of processing a 1. 2. 3. 4.

Dunn, J. J . , and Studier, F. W. (1973). P N A S 70, 3296. Nikolaev, N . , Silengo, L., and Schlessinger, D. (1973). P N A S 70, 3361. Ginsburg, D., and Steitz, J . A. (1975). JEC 250, 5647. Robertson, H . D., Webster, R. E . , and Zinder, N . D. (1968).JBC 243, 82. 485 THE ENZYMES, VOL. XV Copyright Q 1982 by Academic Press, Inc. All rights of reproduction in any form reserved. ISBN 0-12-122715-4

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JOHN J. DUNN

large in virro transcript of bacteriophage T7 DNA to produce RNAs identical in size to those found in infected cells (5). During the purification of this T7 “sizing factor,” it became apparent that it copurified with RNase 111. In contrast to its specific cutting of 30 S preribosomal RNA and T7 RNA, purified RNase I11 seems to degrade double-stranded RNA rather nonspecifically (4. 6). Double-stranded RNA is an effective competitive inhibitor of RNase 111cleavage at processing sites, suggesting that specific cleavage sites in single-stranded RNAs may have some double-stranded character ( I , 6). Support for this idea has come with the delineation of the nucleotide sequence surrounding sites known to act as processing signals for RNase I11 (7-12). Enzymatic activities that degrade double-stranded RNA have been identified in a variety of eukaryotic tissues, and some of these enzymes may play roles in the processing of precursor RNAs. This chapter focuses primarily on RNase I11 of E. coli since it is the most thoroughly studied.

11.

Purification

RNase I11 appears to be the only enzyme in extracts of E. coli that selectively degrades double-stranded RNA (4, 13, 14). Therefore, doublestranded RNase activity, rather than processing activity, is usually monitored during the enzymes purification. The original purification procedure of Robertson et al. ( 4 ) was improved by using an RNase I-deficient strain of E. coli as starting material, and by the introduction of affinity chromatography on columns of poly(1) . poly(C) covalently linked to agarose ( I . 5 ) . In the absence of divalent cations, which RNase I11 requires for enzymatic activity, the enzyme binds strongly to the poly(1) poly(C) column and is eluted only when the monovalent salt concentration, usually NH4C1, reaches 1 M or higher. With some lots of poly(1) * poly(C) agarose, the RNase I11 activity originally bound to the column elutes near 1 M salt, whereas with other lots

-

5 . Dunn, J. J., and Studier F. W. (1973). PNAS 70, 1559. 6. Robertson, H. D . , and Dunn, J. J . (1975).JBC 250, 3050. 7. Rosenberg, M., and Kramer, R. A. (1977).PNAS 74, 984. 8. Robertson, H. D., Dickson, E . , and Dunn J. J . (1977). PNAS 74, 822. 9. Oakley, J . , and Coleman, J. (1977). PNAS 74, 4266. 10. Young, R. A., and Steitz, J. A. (1978). PNAS 75, 3593. 1 1 . Bram, R. J . , Young, R. A . , and Steitz, J. A. (1980). Cell 19, 393. 12. Dunn, J. J . , and Studier, F. W. (1981). J M B 148, 303. 13. Kindler, P., Keil, T. U., and Hofschneider, P.H. (1973). Molec. G e n . Genet. 126, 53. 14. Nikolaev, N . , Silengo, L., and Schlessinger, D. (1973). JBC 248, 7967. 15. Dunn, J. J. (1976).JBC 251, 3807.

15. RIBONUCLEASE 111

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salt concentrations between 1.5-2 M are required to completely elute the enzyme. Enzymatic activity is routinely assayed by measuring the release of acid-soluble counts from [3H]poly(A-U) synthesized by E. toli RNA polymerase using poly(dA-dT) as template (4). One unit of enzyme is defined as the amount of enzyme that will solubilize 1 nmol of acidprecipitable polynucleotide phosphorus per hour under standard conditions. The most highly purified preparations of the enzyme have activities in the range 45,000-90,000 u n i t s h g protein. In addition to divalent cations (0.005-0.1 M Mg2+ or 0.05-0.1 mM Mn*+)RNase 111 has an absolute requirement for monovalent cations ( 4 ) . NH4+,Na', and K + all stimulate digestion of double-stranded RNA, with maximum activity being observed between 0.15-0.3 M . Ca2+cannot replace Mg2+or Mn2+.Cuz+,Fe3+,HgZf,and Co2+are all reported to inhibit enzymatic activity (16). Hydrolysis of poly(1) poly(C) and processing of 30 S preribosomal RNA are totally inhibited by the intercalation agent ethidium bromide at a concentration of 1.5 mM (17). RNase I11 binds tightly to ribosomes at low to moderate ionic strengths, and in the purification procedure of Robertson et al. this association is used as an initial fractionation step ( 4 ) . Washing the ribosomes with 0.2 M salt seems to release all the RNase I11 activity without significantly impairing their ability to suppart protein synthesis ( 4 ) . The low-salt-washed ribosome fraction usually employed in ribosome binding experiments contains considerable amounts of RNase 111. 111.

Structure

The molecular weight of the purified enzyme was determined originally by sucrose gradient centrifugation to be between 45,000 and 55,000 ( 4 ) . Chromatography on Sephadex G-100 is consistent with a molecular weight of 50,000. Polyacrylamide gel electrophoresis of the purified enzyme in the presence of sodium dodecyl sulfate gives one band of protein (Fig. 1) with a molecular weight of approximately 25,000 (15, I8), suggesting that the native enzyme is a dimer. If double-stranded RNA is incorporated into the sodium dodecyl sulfate gel prior to electrophoresis, RNase I11 activity can be detected in sitir after removal of the sodium dodecyl sulfate by diffusion. Upon staining the gel with ethidium bromide, activity appears as a dark band on a fluorescent background of ethidium bromide 16. Paddock, G . V., Fukada, K . , Abelson, J . , and Robertson, H . D. (1976).Nucleic Acids R e s . 3, 1351. 17. Nikolaev, N . , Birge, C. H . , Gotoh, S . , Glazier, K . . and Schlessinger, D. (1975). BrooXlin\~c.riS y m p . B i d . 26, 175. 18. Darlix, J . L. (1975). EJB 51, 369.

FIG. 1. SDS-polyacrylamide gel electrophoresis of fractions during the purification of RNase 111. Portions of the following samples were analyzed by electrophoresis on a 15%

489

15. RIBONUCLEASE 111

bound to undigested double-stranded RNA (19). Only one band of activity is observed in both crude extracts and highly purified enzyme preparations, and this band has the same electrophoretic mobility as the protein component visualized by staining the purified enzyme with Coomassie brilliant blue. An RNase 111-deficient mutant of E. coli AB301-105, has been isolated by Kindler et 01. (13). The mutation, mc-105, maps near 55 min on the genetic map of E. coli, and isogenic RNase 111' and RNase 111- strains of E. c d i are now available for studying thein vivo role of RNase I11 (20-22). The RNase 111- strains are useful not only in the isolation of 30 S preribosomal RNA but also in identifying other RNA processing enzymes that normally act on the RNase 111cleavage products of 30 S RNA (23). In the absence of RNase I11 cleavages, these other enzymes apparently act more slowly on the ribosomal RNA transcript to form an alternate pathway for the maturation of ribosomal RNAs ( / ) .

IV.

Cleavage of Double-Stranded RNA

RNase I11 cleavage of double-stranded RNA has been characterized with regard to the type of end groups produced and the size of the products. Exhaustive RNase 111digestion of double-stranded RNA produces a mixture of oligonucleotides having 5'-phosphate and 3'-hydroxyl termini ( 6 , 2 4 ) .The final size distribution, as determined by homochromatography and end group analysis, is rather narrow with the median size being about 15 bases long. The most straightforward interpretation of this result is that the final size of the products reflects the requirement for a stable doublestranded structure. It is not known with certainty if RNase I11 can cleave between all combinations of bases or if particular combinations are cleaved more readily than others. To try to elucidate this, end group analysis has been performed on RNase I11 digests of polyoma double19. 20. 21. 22. 23. 24.

Rosenthal, A. L., and Lacks, S. A . (1977). Atrtrl. Biochem. 80, 76. Studier, F. W. (1975). J . Bocteriol. 124, 307. Apirion, D . , and Watson, N. (1975). J . Bocreriol. 124, 317. Bachmann, B . J . , Low, K . B., and Taylor, A. L. (1976). Bacreriol. Abelson, J. (1979). Annu. Rev. Biochem. 48, 1035. Crouch, R. J . (1974). JBC 249, 1314.

Reil.

40, 116.

polyacrylamide slab gel containing 0.1% sodium dodecyl sulfate ( I S ) : (a) high-speed supernatant; (b) ammonium sulfate fraction; (c) flow-through of DEAE-cellulose column; (d) flow-through of poly(1) . poly(C) column; (e) 2 M NHICl eluent of poly(1) poly(C) column. Prior to electrophoresis the samples were added to sodium dodecyl sulfate sample buffer and placed in a boiling water bath for 2 min.

490

JOHN J. DUNN

stranded RNA (6). It was concluded that RNase 111cleavage in exhaustive digests of this RNA is nearly sequence-independent although a slight enrichment for A and U at the 5' termini was noted. On the other hand, analysis of the products formed during RNase 111 digestion of the doublestranded RNAs of Penicillium chrysogenirm virus led Edy er d. (25) to conclude that cleavage of this RNA is a two-step process. During the initial phase, cleavage tends to be specific and 20-25 distinct bands are obtained after polyacrylamide gel electrophoresis. Phosphorylation of the 5' ends of the RNA fragments with polynucleotide kinase revealed a preponderance of A and U at the 5' termini. More extensive digestion caused all the bands to disappear. At present it is not known if RNase I11 cleaves both strands of double-stranded RNA in a concerted fashion or if the cleavage of double-stranded RNA is the result of the accumulation of independent single-strand cuts. RNA hybridized to DNA is not degraded by purified RNase 111 (6, 24) and, as expected, neither poly(A) nor poly(U) alone is a substrate for the enzyme. It has been reported that RNase 111cleaves the poly(U) strand of poly(A) * poly(U) much faster than it does the poly(A) strand (26). However, the conditions used (10 mM Mg") are known to promote triple helical formation, which might have affected the rate of degradation (27). V.

Cleavage of Single-Stranded RNA

The cleavage patterns obtained after treatment of single-stranded RNAs with RNase I11 at different concentrations of enzyme and monovalent salt revealed that there are two types of RNase I11 cleavage site (15). One type, primary sites, corresponds to sites normally cut by the enzyme in rivo. These are also the preferred cleavage sites in vitro and they are the only sites cut readily at monovalent salt concentrations around 0.25 M. The other type, secondary sites, is cut much less efficiently than primary sites under all conditions tested. The ability of RNase I11 to cleave at secondary sites depends strongly on the ionic strength of the reaction mixture, increasing as the ionic strength decreases. As a rule, cleavage at primary or secondary sites does not release small, acid-soluble oligonucleotides, indicating that RNase 111 makes a single cut rather than a cluster of cuts (28). 25. 26. 27. 28.

Edy, V. G . , Szekely, M., Loviny, T., and Dreyer, C. (1976). EJB 61, 563. Bishaeye, S . , and Maitra, U . (1976). BBRC 73, 306. Suryanarayana, T., and Burma, D. P. (1975). BBA 407, 459. Dunn, J. J . , and Studier, F. W. (1975). Brookhaivn Symp. B i d . 26, 267.

15.

RIBONUCLEASE 111

49 1

A. PRIMARY SITES The first primary sites to be identified were those in bacteriophage T7 early RNA ( 5 ) . Synthesis of this 7000-base-long RNA is initiated at a cluster of three promoters forE. coli RNA polymerase near the left end of the T7 DNA, and continues rightward until a transcription termination signal is reached 19% of the way along the DNA molecule (5, 29. 30). RNase I11 cuts this large RNA at five primary sites to generate the T7 early messenger RNAs, and the cuts made by the purified enzyme in v i m are known to be identical to those madein vivo (Fig. 2) (31,32).The effects of deletion mutants of T7 on cleavage demonstrate that cleavage at any one site is an independent event (i.e., the lack of one or more sites does not effect the other cleavages). These deletion mutations also aid in determining the position of the five primary sites and help in defining the minimum length of RNA surrounding each site needed for normal enzyme-substrate recognition. The nucleotide sequence of the entire early region of T7 has been determined and, by analysis of the sequences at the ends of the individual early RNAs, the precise positions of the five primary cleavage sites and the number of cuts made are now known (12). While a few sequence elements are conserved in all five sites, the most constant feature found is the potential for extensive base-pairing that can fold the RNA into a localized stem and loop structure. Each stem contains two base-paired regions separated by an internal bubble within which a single phosphodiester bond in the 3' half is broken to generate the 5'-phosphate and 3'-hydroxyl termini of the processed RNAs. The size of the base-paired regions and internal bubbles vary from site to site; however, none of the T7 sites has more than 12 uninterrupted base-pairs in either the upper or lower stem portion, and none has more than five bases in its loop. This is in striking contrast to the giant stem and loop structures deduced (10, / I )for two RNase 111 cleavage sites in 30 S preribosomal RNA. One hairpin loop contains the entire 16 S ribosomal RNA molecule, the other the 23 S ribosomal RNA. In addition, both loop structures are closed by regions of more extensive uninterrupted base-pairing (up to 26 base-pairs in the case of 16 S ribosomal RNA) than is observed in T7 (Fig. 3). Within each double-helical stem region RNase I11 cleaves twice, once in the 5' 29. 135. 30. 31. 32.

Millette, R . L., Trotter, C. D., Herrlich, P., and Schweiger, M. (1970). CSHSQB 35, Minkley, E. G.. and Pribnow, D. (1973). J M E 77, 255. Kramer, R. A., Rosenberg, M., and Steitz, J . A . (1974). J M B 89, 767. Rosenberg, M . , Kramer, R. A., and Steitz, J . A. (1974). J M E 89, 777.

FIG.2. Effect of monovalent salt concentration on the specificity of cleavage of T7 early RNA by RNase 111. T7 early RNA was incubated with RNase I11 for 20 min at 37" in 5% sucrose, 0.02 M Tris-HCI (pH 7.9), 0.005 M MgCl,, 0.1 rnM EDTA, 0.1 mM dithiothreitol,

15. RIBONUCLEASE 111

493

/

\

ribosomal 23s

G AAUCA G

A U

G

c

c

C=G U-A C=G G*U C=G U.G G=C G=C A-U A- U

c

u

U

A

,"-

c A

A G-U A-U U-A A-U G=C U-A G=C A-U G-U

.......

u

5' GC 0.3 RNA

u

AC .......3' 0.7 RNA

RNA

cc

AGUG G C=G G=C A-U A-U U-A U-A G=C G=C A-U G-U U-A G=C U-A U-A +G=C G=C G=C C=G U-A U-A C=G U-G A-U C=G A-U A-U A- U G*U U*G G G 5 ' .......AA CG .......3'

~

FIG.3. Nucleotide sequences and secondary structures predicted to form surrounding two primary RNase I11 cleavage sites. Shown on the left is the cleavage site between the gene 0.3 and 0.7 messenger RNAs of T7 (7). On the right is the stem-loop structure encompassing 23 S ribosomal RNA ( 1 1 ) . The arrows indicate where the cuts are made. and N H C I as indicated. Each incubation mixture contained, in a final volume of 50 p1, 20,000 cpm T7 early [3'P]RNA and 1.5 units of RNase 111. Equal portions of each sample were then electrophoresed on a 2% polyacrylamide plus 0.5% agarose gel and on a 3-2095 polyacrylamide gradient gel. The RNA applied to the tracks marked control was from an incubation mixture that received no RNase 111. The positions of the individual early messenger RNAs, 0 . 3 , 0 . 7 , I , 1 . 1 , and 1.3, plus the three initiator RNAs, 11,I,, and I,, are indicated to the right of each gel pattern. Similar patterns were obtained using NaCl or KCI in place of NH4CI. The normalin rivo pattern is obtained at salt concentrations between 0.15 and 0.3 M (5, IS).

494

JOHN J. DUNN

half and once in the 3' half, to release the portions of the RNA destined to become 16 and 23 S ribosomal RNAs. The double cleavages are staggered by two base-pairs, a configuration noted by Bram et al. (II), that would place the bonds to be cleaved close to each other on the same side of the double-stranded stem regions. It is not known if the double cleavages are concerted or if they are sequential. A double cleavage also occurs, albeit rather inefficiently, within the region between genes 1.1 and 1.3 of T7 to release a 29-base-long fragment called F5 (8, 28). Of all the T7 primary sites, this region comes closest to resembling a double-stranded ribosomal RNA stem structure in that it has a stretch of 16 potential base-pairs interrupted by a single CU mismatch. When there is a double cleavage within this region it is also staggered by two bases; however, cleavage in the 5' half of the site seems to be inefficient; only the cleavage in the 3' half behaves in all respects as a primary cleavage. Separating the cleavage sites that generate the 16 and 23 S ribosomal RNAs are spacer regions containing sequences for transfer RNAs (in four of the seven ribosomal RNA operons tRNAB'" is present; in three of the operons tRNA:le and tRNA9la are both present) (33-36). I n v i m , RNase I11 seems to cleave once in the 437-base-long spacer region to produce a smaller 290-base-long dimeric tRNA precursor (37). This cleavage occurs in a region of the transcript that appears to lack extensive, uninterrupted base-pairing . Potential RNase I11 cleavage sites are thought to be present near the 5' ends of hpt-RNA (38),lac messenger RNA (39), and the messenger RNA coding for the /3 subunit of E. coli-RNA polymerase (40). RNase I11 also processes bacteriophage T3 early RNA (41). The enzyme can cut some 6x174 transcripts (42) and it is reported to have a role in producing an RNA primer need for ColElDNA replication (43). 33. Deonier, R . C . , Ohtsubo, E., Lee, H. J., and Davidson, N . (1974).J M B 89, 619. 34. Lund, E., Dahlberg, J. E., Lindahl, L . , Jaskunas, S. R., Dennis, P. P., and Nomura, M. (1976). Cell 7, 165. 35. Lund, E . , and Dahlberg, 3. E. (1977). Cell 11, 247. 36. Young, R. A , , Macklis, R., and Steitz, J. A. (1979).JBC 254, 3264. 37. Lund, E., Dahlberg, J. E., and Guthrie, C. (1980). In "Transfer RNA: Biological Aspects" (D. So11, J. N. Abelson, and P. R . Schimmel, eds.), p. 123. Cold Spring Harbor Laboratory, Cold Spring Harbor, New York. 38. Lozeron, H. A., Dahlberg, J. E., and Szybalski, W. (1976). Virology 71, 262. 39. Cannistrado, V. J . , and Kennell, D. (1979).Nntrtre (London) 277, 407. 40. Post, L . , Strycharz, G . D . , Nomura, M., Lewis, H . , and Dennis, P. (1979). P N A S 76, 1697. 41. Anderson, C. W., Atkins, J. F., and Dunn, J. J. (1976). PNAS 73, 2752. 42. Kapitza, E. L., Stukacheva, E. A . , and Shemyakin, M. F. (1976).FEES Letr. 64,81. 43. Conrad, S. E., and Campbell, J. L. (1979). Cell 18, 61.

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495

Exactly how RNase I11 recognizes a primary site and chooses which phosphodiester bond(s) to break remains unclear. The enzyme acts directly on purified primary transcripts; auxiliary factors are not required. It seems clear from the analysis of two mutations of T7 that interfere with RNase I11 cleavage that the specific structure of the RNA is important for cleavage (44). No sequence element(s) common to all known primary sites has been identified. However, limited sequence homologies between the primary sites in T7 early RNA and the stem regions in 30 S preribosomal RNA have been noted (11). Whether these regions of homology contribute to the specificity of cleavage remains to be established. It has been argued that the mere lengths of base-paired regions are insufficient to properly align the enzyme (6). In this regard it is interesting to note that additional sites of cleavage have been reported in the ribosomal RNA stems (11, 37, 45). The stems themselves can be isolated from 30 S preribosomal RNA by using single-strand specific nucleases. When incubated with RNase I11 the isolated stems are cut at the known preferred cleavage sites, and also at the additional cut sites (46). In the case of the T7 sites, which have less extensive double-stranded character, additional cleavages have not been observed. Perhaps the ribosomal stems have additional cleavages because they more closely resemble completely double-stranded RNA. B.

SECONDARY SITES

When T7 early RNA is incubated with high levels of RNase 111, cuts in addition to those at the five primary sites are observed. The ability of the enzyme to cleave these secondary sites increases as the monovalent salt concentration of the incubation mixture is decreased (Fig. 2). However, even at low ionic strength primary sites are still the preferred sites of cleavage and they are the sites cut initially. At a constant enzyme to substrate ratio, the size range of the cleavage products depends on the salt conditions chosen for cleavage, demonstrating that the optimal ionic conditions vary for different secondary sites. The reproducibility of the patterns obtained with T7 early RNA as well as with 16 and 23 S ribosomal RNA under a n y given set of conditions suggests that secondary sites occur at specific locations on the RNAs. It also suggests that RNase I11 might be useful for producing fragments from many RNAs in much the 44. Studier, F. W., Dunn, J. J . , and Buzash-Pollert, E. (1979). Micrmi Winter Symp. 16, 261. 45. Lund, E . , and Dahlberg, J. E. (1979). P N A S 76, 5480. 46. Robertson, H . , Pelle, E. G . , and McClain, W. H. (1980). In “Transfer RNA: Biological Aspects” ( D . SOH, J . N . Abelson, P. R . Schimmel, eds.), p. 107. Cold Spring Harbor Laboratory, Cold Spring Harbor, New York.

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JOHN J. D U N N

same manner as specific DNA fragments are produced using restriction endonucleases. Most RNAs tested that are not natural substrates for RNase I11 seem to lack primary sites, but many different RNAs can be cut at secondary sites by using the appropriate ionic strength and enzyme concentration. Studies with poliovirus RNA show that although it lacks the equivalent of primary sites, it contains numerous secondary sites because it is readily cut by RNase I11 at NHsCl concentrations below 0.1 M (47). Cleavage is clearly not random as reproducible, and different cleavage patterns are obtained with the genomic RNAs of poliovirus types 1, 2, and 3 when they are incubated under conditions that limit cleavage to only a few secondary sites. Cleavage at these sites has been used to help map the location of deletions of defective interfering particles of poliovirus in the RNAs (481, and to generate subgenomic fragments for use in mapping poliovirus gene products (49). Less stringent incubation conditions have been used to produce an RNA fragment that contains the first 90 or so nucleotides of poliovirus RNA still linked at its 5' end to the small protein VPg (50). Other eukaryotic viral RNAs known to be cut by RNase I11 are those from Rous sarcoma (51, 52), vesicular stomatitis (53),influenza (Desselberger and Dunn, unpublished), and brome mosaic (Burr and Dunn, unpublished) viruses. Adenovirus messenger RNA (54), HeLa heterogeneous nuclear RNA ( 5 3 , and 45 S preribosomal RNA (17) are also substrates. Thus, it seems likely that any large RNA probably can be cut by RNase I11 provided the proper conditions are used. Attempts to cut small, sequenced RNAs, such as 5 S and tRNAs, to gain insight into the mechanism of secondary site selection have been for the most part unsuccessful. Small RNAs are not usually cleaved. One notable exception is the 140-base-long Species I RNA synthesized after infection of E. coli with bacteriophage T4. Although this RNA does not appear to be cut in vivo by RNase 111, it can be cut in vitro provided the salt concentration is below 0.03 M (16). This RNA, which has some structural features in common with tRNAs, is cut in two places to produce 47. Nomoto, A., Jacobson, A., Lee, Y. F., Dunn, J. J . , and Wimmer, E. (1979).JMB 128, 179. 48. Nomoto, A., Lee, Y. F., Babich, A . , Jacobson, A., Dunn, J. J., and Wimmer, E. (1979). J M B 128, 165. 49. Stewart, M . , Crouch, R. J . , and Maizel, J. V., Jr. (1980). Virology 104, 375. 50. Harris, T. J. R . , Dunn, J. J . , and Wimmer, E. (1978). Niicleic Acids Res. 5, 4039. 51. Leis, J. P., McGinnis, J., and Green, R. W. (1978). Virology 84, 87. 52. Darlix, J . L., Spahr, P. F.. and Bromley, P.A. (1978). Virology 90, 317. 53. Wertz, G . W., and Davis, N . L . (1979). J . Virol. 30, 108. 54. Westphal, H . , and Crouch, J. R. (1975). PNAS 72, 3077. 55. Robertson, H. D., and Dickson, E. (1975). Erookhuven Symp. B i d . 26, 240.

15. RIBONUCLEASE I11

497

fragments 19, 48, and 73 nucleotides long. Both cleavages occur within a region of presumed secondary structure. Each side of the paired structure is cut once and the cuts are again staggered by two bases. Bacteriophages T2 and T6 specify Species I RNAs that have base substitutions near the potential cleavage sites. Interestingly, one of these substitutions would decrease the stability of the base-paired region surrounding the cleavage sites, and seems to hinder cutting at only one of the two potential cut points. All cleavages by RNase 111, including those at secondary sites, release RNAs with 5’-phosphate and 3’-hydroxyl termini. There is every indication that secondary cuts generate unique ends, and that RNase 111 can be used to produce RNA fragments suitable for use with rapid RNAsequencing techniques (56). The observation that lowering the monovalent salt concentration increases the number of potential RNase I11 cleavage sites in a singlestranded RNA seems paradoxical, since double-stranded structures (the apparent recognition sites for RNase 111) should be destabilized. However, the patterns of Fig. 2 were obtained using 5 mM Mg2+, which stabilizes base-pairing. Perhaps decreasing the monovalent salt to Mg2+ ratio changes the overall structure of the RNA sufficiently to allow the enzyme to find potential sites that are inaccessible or not even present at higher ratios. Another possibility is that the interaction between enzyme and substrate increases at low ionic strength. This might allow primary as well as secondary sites to bind RNase I11 more efficiently, resulting in a greater probability that both will be cleaved. It is also possible that the enzyme is directly affected by changes in salt concentration, leading to alteration of its conformation or its monomer-dimer equilibrium with resulting changes in its specificity.

VI.

Related Enzymes from Eukaryotes

For some time it has been thought that double-stranded regions are important in determining the specificity of ribosomal and messenger RNA processing in eukaryotic cells. Intercalating agents, such as ethidium bromide, block processing of 45 S preribosomal RNA in vivo (57), as does incorporation of base analogs that weaken base-pairing (58, 5 9 ) . Doublestranded regions are known to be present in heterogeneous nuclear RNA 56. Donis-Keller, H . , Maxam, A . , and Gilbert, W. (1977). Nucleic Acids Reg. 8, 2527. 57. Synder, A. L., Kann, H. E., and Kohn, K. W. (1977). J M E 58, 555. 58. Tavitian, A., Vretsky, S. C . , and Acs, G. (1%8). EBA 157, 33. 59. Wilkinson, D. S., and Pitot, H. (1973). JBC 248, 63.

498

JOHN J. DUNN

but they are no longer found when the RNA is processed into poly(A) containing cytoplasmic messenger RNA (60). The double-stranded regions in 45 S preribosomal RNA can be cleaved by RNase I11 from E. coli to produce RNAs with sizes similar to intermediates observed in vivo ( 6 1 ) , and it has been postulated that enzymes capable of cutting doublestranded RNA might be involved in eukaryotic RNA processing (62-67). Several RNase 111-like activities have been partially purified from the cytoplasmic and nuclear fractions of various eukaryotic cells. Rech et ul. (68) starting with the cytosol of Krebs I1 mouse ascites cells, succeeded in purifying about 650-fold an activity that degrades poly(G) . poly(C). Like RNase 111from E. coli, this enzyme does not bind to DEAE-cellulose but does bind to CM-cellulose. A latent RNase 111-like activity has been found associated with heterogeneous nuclear RNA-protein particles prepared from HeLa cells (69). Only after the particles were chromatographed on DEAE-cellulose or the RNA digested with pancreatic RNase or RNase TI was hydrolysis of exogenous poly(A-U) detectable. An RNase 111-like activity can be extracted by low salt plus EDTA treatment of nucleoli from mouse ascites cells (70). This enzyme, which is thought to have a role in processing 45 S preribosomal RNA, can also be found associated with RNA-protein particles (80 and 50 S preribosomal particles). In this case, the double-stranded RNase activity associated with the particles can be assayed directly. A similar activity has been purified from calf thymus nuclei (71). The purified eukaryotic enzymes all require divalent cations (usually Mg2+around 2 mM) for maximal activity. Moderate concentrations of monovalent salts tend to inhibit enzymatic activity. Double-stranded RNA is digested to yield oligonucleotides with 5’-phosphate termini and its cleavage is inhibited by ethidium bromide. Affinity chromatography on poly(1) . poly(C) agarose has been used in the purification of the RNase 111-like activity from the nucleoli of ascites cells (70). However, unlike 60. 61. BBRC 62. 63. 64. 65. 66. 67. 68. 69. 70. 71.

Jelinek, W., and Darnell, J. E. (1972). PNAS 69, 2537. Gotoh, S., Nikolaev, N . , Battaner, E., Birge, C. H., and Schlessinger, D., (1974). 59, 972. Weinberg, R . A., and Penman, S. (1970). J M B 47, 169. Molloy, G. G . , Jelinek, W., Salditt, M., and Darnell, J. E. (1974). Cell I, 43. MacNaughton, M., Freeman, K . B . , and Bishop, J. 0. (1974). Cell 1, 117. Munoz, R. F., and Darnell, J. E., (1974). Cell 2, 247. Price, R. P., Ranson, L., and Penman, S. (1974). Cell 2, 253. Williams, J. G., and Penman, S. (1975). Ce/l 6, 197. Rech, J . , Cathala, G . , and Jeanteur, Ph. (1976). Nucleic Acids Res. 3, 2055. Rech, J., Brunel, C., and Jeanteur, Ph. (1979). BBRC 88, 422. Grummt, I., Hall, S. H., and Crouch, R. J. (1979). EJB 94, 437. Kenzo, O . , Groner, Y., and Hurwitz, J. (1977). JBC 252, 483.

15.

499

RIBONUCLEASE I11

RNase I11 from E. coli, the ascites enzyme binds less strongly to poly(1) . poly(C) and is eluted by buffer that contains 0.45 M NH4CI. When characterized with regard to substrate specificity, most eukaryotic RNase 111-like activities seem to be less specific for double-stranded RNA than is the enzyme from E. coli (68-72). All digest single-stranded RNA to some extent, and many can degrade RNA hybridized to DNA. An exception is the enzyme from calf thymus (71), which lacks detectable activity against RNA hybridized to DNA. This enzyme cleaves 45 S preribosomal RNA into fragments that approximate the size of eukaryotic ribosomal RNA, and it also cuts duck reticulocyte heterogeneous nuclear RNA to produce mainly 16 and 10 S fragments. During cleavage of either substrate, acid-soluble material is not released. It remains to be established if the broader specificity of the enzymes purified from other sources is caused by contamination with additional RNases, or if lower specificity is an intrinsic property of these RNase 111-like activities. Although many lines of evidence suggest that RNase 111-like activities may be involved in processing eukaryotic RNAs, unequivocal data concerning any one enzyme and substrate are lacking. Recent studies by Ferrari et ul. (73) suggest that endonucleases with no apparent activity against double-stranded RNA may be involved in processing HeLa 45 S preribosomal RNA and heterogeneous nuclear RNA. Presumably, the utilization of cloned DNA fragments will eventually make it possible to determine precisely if apparent processing by RNase 111-like activities in ritro correspond to those that occur ii? v i i w .

ACKNOWLEDGMENTS Research for the author's studies reported in this chapter were carried out at Brookhaven National Laboratory under the auspices of the United States Department of Energy.

72. Busen, W . , and Hausen, P. (1975). EJB 52, 179. 73. Ferrari, S., YeNe, C. 0.. Robertson, H. D., and Dickson, E. (1980). P N A S 77,2395.

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RNases, I, 11, and IV of Eschichia coli V. SHEN

D. SCHLESSINGER

I. Introduction . . . . .

.

.

.

. , . . . . . . . . . . , . A. Intracellular Location, Purification, and Properties . . B. Mechanism of Action . . . . . . . . . . . . . . . C. Biological Role and Possible Application . . . . . . . 111. RNase I1 of Esckericliicr coli . . . . . . . . . . , . , . A . Purification and Properties . . . . . . . . . . . . . B. Mechanism of Action . . . . . . , , , . , , . . . C. Biological Role and Possible Application . , , , , , . IV. RNase IV of Escherickici coli . . . , . , , , , , , . , A. Purification and Properties . . . . . , . , . . , . . B. Mechanism of Action . . . . . . . , . . . . , . . C. Biological Role and Possible Application . . . . . . . 11. RNase I of E.schrrichirr coli

I.

,

. . . ,

, . , . . . . .

. . . . .

. . . . . . . . . ,

,

.

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

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

501

503 503 505 505 506 506 508 510

512 512 513 5 15

Introduction

RNase I, 11, and IV are the most venerable of the known E. coli RNases, but have no known indispensible functions in the cell. These enzymes were only briefly mentioned earlier in this series (I). RNase I of E. coli was first discovered as a latent ribonuclease associated with ribosomes (2). This tight association was later found to be an 1. Uchida, T., and Egami, F. (1971). “The Enzymes,” 3rd ed., Vol. IV, p. 204. 2. Elson, D. (1959). BBA 36, 372. 50 1 THE ENZYMES. VOL. X V Copyright 0 I982 by Academic Press, Inc. All rights of reproduction in any form reserved. ISBN 0-12-122715-4

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artifact arising during cell disruption: its normal intracellular location is in the periplasmic space. Various essential biological roles for RNase I in cell physiology have been proposed ( 3 ) ,but all are unlikely since naturally occurring and laboratory mutant strains that are defective in RNase I (4) show no apparent change in RNA metabolism. RNase I1 ofE. coli was discovered as a K'-activated phosphodiesterase ( 4 , 5 )that was also ribosome-associated in extracts. It is the predominant exoribonuclease seen in extracts, with RNase 11-deficient mutants showing much lower levels of RNase activity than wild-type (6, 7). It has been purified to homogeneity by several groups (8-11), and many of its physical properties have been well-studied. The action of RNase I1 seems to be highly dependent on RNA conformation with different optimal ionic conditions for different substrates (810). No correlations can be drawn within vivo function, however. Roles in rRNA asd tRNA processing, and in mRNA degradation, have been proposed (5, 12-22); but the enzyme seems most likely to be only a salvage enzyme active in RNA turnover in stressed cells. RNase IV of E. coli has been much less studied. It cleaves bacteriophage R17 RNA at limited sites and was useful in mapping genes along the RNA (23). It has also been used in studies of 5 S rRNA and Rous sarcoma virus RNA structure. Its biological role is unknown. 3. Datta, A . K., and Niyogi, S . K. (1976). Progr. Nircleic Acid Res. M o l . B i d . 17, 271. 4. Wade, H . E. (1961). BJ 78, 457. 5. Spahr, P. F., and Schlessinger, D. (1963). JBC 238, PC2251. 6. Castles, J. J., and Singer, M. F. (1968). BBRC 33, 715. 7. Nikolaev, N., Folsam, V., and Schlessinger, D. (1976). BBRC 70, 920. 8. Spahr, P. F. (1964). JBC 239, 3716. 9. Singer, M. F., and Tolbert, G. (1965). Biochemistry 4, 1319. 10. Gupta, R. S., Kasai, T., and Schlessinger, D. (1977). JBC 252, 8945. 11. Leineweber, M., and Philips, G. R. (1978). J . Anal. Biochem. 517, 419. 12. Corte, G., Schlessinger, D., Longo, D., and Venkov, P. (1971). JMB 60, 325. 13. Yuki, A. (1971). J M B 56, 435. 14. Yuki, A. (1971). JMB 62, 321. 15. Schedl, P., Roberts, J., and Primakoff, P. (1976). CelI 8, 581. 16. Kitamura, N., Ikeda, H., Yamada, Y.,and Ishikura, H. (1977). EJB 73, 297. 17. Birenbaum, M., Schlessinger, D., and Ohnishi, Y. (1980). J . Bacferiol. 142, 327. 18. Sekiguchi, M., and Cohen, S. S . (1963). JBC 238, 349. 19. Sekiguchi, M., and Cohen, S. S. (1964). JMB 8, 638. 20. Andoh, T., Natori, S., and Mizuno, D. (1963). BBA 76, 447. 21. Futai, M., Anraku, Y., and Mizuno, D. (1966). BBA 119, 373. 22. Castles, J. J., and Singer, M. F. (1969). JMB 40, 1. 23. Jippsen, P. G. N., Steitz, J. A , , Gasteland, R. F., and Spahr, P. F. (1970). Nature (London) 226, 230.

16. RNase I, 11, AND 1V OF E. coli

503

Most studies of these enzymes have been done in E. coli. However, RNase I from Salmonella typl~imririirmhas also been studied (24). RNases with similar modes of action have also been found in ryegrass (25), tobacco leaves (26), and pea leaves (27) [the plant enzymes have been referred to as ribonuclease T,]. Activities similar to E. coli RNase I1 have also been purified from S . typhimurium (28) and Lactobacillus plantarum (29). No enzyme similar to RNase IV of E. coli has been reported from any other organism. II.

RNase I of Escherichia coli

A. INTRACELLULAR LOCATION, PURIFICATION, AND PROPERTIES 1. ttitracellular Location

RNase I was first detected as a latent RNase because its activity is not expressed unless it is released from ribosomes by urea, high salt, EDTA, or other reagents that modify ribosome structure (2). The enzyme was found selectively in 30 S ribosomes (30), but some enzyme activity with similar properties was also found in the cell debris and soluble proteins of cells after cold shock (31, 32). The location of the enzyme in vivo was resolved when it was found that carefully prepared spheroplasts are devoid of RNase I-like activity (33, 34). The enzyme can be released quantitatively into the medium when cells are treated with membrane-active peptide antibiotics that penetrate only the periplasmic space (35). Furthermore, purified RNase I can bind strongly to ribosomes in vifro (33). Therefore, the intracellular location of RNase I was unequivocally assigned to the periplasmic space, the association with 30 S ribosomes occurring only after cell disruption. 24. 25. 26. 27. 28. 29. 30. 31. 32. 33. 34. 35.

Chakraburtty, K., and Burma, D. P. (1968). JBC 243, 1133. Schuster, L., Khorana, H. G., and Heppel, L. A. (1959). BBA 33, 452. Reddi, K. K . (1958).BBA 28, 386. Markham, R., and Strominger, J. L. (1956). EJ 64,460. Ray, R. K . , and Burma, D. P. (1970). BBA 212, 102. Logan, D. M., and Singer, M. F. (1968). JBC 243, 6161. Spahr, P. F., and Hollingworth, B. R. (1961). JBC 236, 823. Anraku, Y., and Mizuno, D. (1965). E B R C 18, 462. Neu, H. C., and Heppel, L. A. (1964). BBRC 17, 215. Neu, H. C., and Heppel, L. A . (1964). P N A S 51, 1267. Neu, H. C., and Heppel, L. A. (1964). JBC 239, 3893. Teuber, M., and Cerny, G. (1970). FEES Lett. 8, 49.

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2. Purijication The standard assay of RNase I of E. coli is performed in 0.1 M Tris buffer, pH 8.1, at 25" for 25 minutes with 0.6% yeast RNA as substrate. One unit of RNase I is defined as the amount of enzyme that, under these conditions, produces 0.1 absorbancy unit of acid-soluble material at 260 nm. Purification of RNase I begins from E. coli ribosomes (30). The ribosomes are first incubated in 6 M urea and then dialyzed against 0.1 M Tris buffer that contains 6 M urea to promote complete digestion of the rRNA. When the protein solution is dialyzed against water, the bulk of the ribosomal protein precipitates. The soluble proteins were fractionated with ammonium sulfate fraction (40-65% saturation) and absorbed to Amberlite CG-50 type 11. After it is eluted by 0.2 M phosphate buffer, pH 7.2, the active fraction is further purified on Amberlite CG-50 type I11 and eluted with 0.1 M phosphate buffer, pH 7.14, that contains 0.12 M NaC1. The purification factor starting from ribosomes is about 500-fold (30). The RNase I from S. typhimurium has been purified by Sephadex GlOO and DEAE fractionations combined with repeated (NH&S04 fractionation (24).

3. Physical Properties RNase I was extensively studied by Spahr and Hollingworth (30). It appears to be a low-molecular-weight, basic protein with a single subunit. The pH optimum in 0.1 M Tris buffer was found to be 8.1; at pH 7.5 and 8.5, the activity was approximately 85% of that at pH 8.1. RNase I was stable for 15 min at 100" at pH 3.1, but all activity was lost after heating at pH 8.8. The enzyme was stable from pH 3.9 to 8.0 at 4" for at least 4 days; at pH 9.8, only 8% of activity was lost after the same time at 4".The enzyme can be stored after dialysis against water at -20". NaCl, KC1, and NaF enhance enzyme activity, with an optimum at 0 . 2 M , whereas Mg2+is slightly inhibitory. Denatured DNA seems to inhibit RNase I activity by competitively binding to the enzyme (36). RNase digestion of all synthetic polynucleotide is stimulated by polyamines, although the degree of stimulation varies with substrate [poly(C) > poly(A) = poly(U) (37)l. The differential stimulation of the digestion of different polynucleotides by polyamines decreases as the monovalent cation concentration is increased (37). Some properties of a similar enzyme from S. typhimurium have also 36. Mukhopadhyay, A . K., and Burma, D. P. (1969). BBA 190, 232. 37. Kumagai, H., Igarashi, K . , Yoshikawa, M., and Hirose, S. (1971).J . Biochern. 81, 381.

505

16. RNase I, 11, AND IV OF E . coli

been investigated. This enzyme shows some differences in cation stimulation. The relative rates of hydrolysis of homopolymers are in the order poly(U) > polyCC) or poly(A) $- tRNA. The specificity for single-strand substrates was inferred from the observation that double-stranded poly(A)-poly(U) formed in high concentrations of mixed poly(A) and poly(U) is inhibitory to RNase I activity (24).

B. MECHANISM OF ACTION Partial digestion of RNA by RNase I produces 2’,3‘-cyclic adenylic, cytidylic, guanylic, and uridylic phosphates as major acid-soluble products. When digestion is carried to completion, nucleoside 3’monophosphates are generated. As expected from these results, purified enzyme slowly converts cyclic nucleotides to 3’-mononucleotides. Thus, the enzyme can cleave all internucleotide bonds in RNA, giving rise to 2’,3‘-cyclic nucleotides, which are in turn more slowly hydrolyzed to the 3’-mononucleotides. Also, the dialyzable fragments released early in the digestion of rRNA showed a preponderance of A and a relative deficit of C residues, suggesting a possible preference for cleavage at A and U residues (30).

c.

BIOLOGICAL ROLEA N D

POSSIBLE

APPLICATION

1. A Scavenger Role?

Protein synthesis in extracts from RNase I + and RNase I- strains, directed by natural mRNA or poly(U), is identical (38), and mRNA metabolism in a naturally occurring RNase I- strain (MRE600) is also unaffected (38, 39), suggesting that RNase I is not involved in mRNA degradation. Ribosomal RNA is stable during exponential growth of RNase I-containing strains (40). Slow rRNA turnover occurs in stressed cells (starved for Mg2+or carbon), and large rRNA fragments are transiently formed (41). It has been proposed that RNase I could enter these cells from the periplasmic space after an alteration or rupture of the cytoplasmic membrane in the stressed cells, and could participate in the turnover processes. In this regard, a speculative analogy could compare the sequestered location of RNase I to the ‘‘latent’’ lysosomal RNase that 38. Gesteland, R . F. (1%7). J M B 16, 67. 39. Cammack, K . A., and Wade, H. E. (1965). BJ 96, 671. 40. Pato, M . L., and Meyenburg, K. V. (1970). CSHSQB 35, 497. 41. Kaplan, R . , and Hartstein, E. (1976). JBC 251, 1147.

506

V. SHEN AND D. SCHLESSINGER

may be involved in rRNA turnover in eukaryotic cells. However, the rate of rRNA turnover is unchanged in a mutant that is deficient in RNase I, and oligonucleotides and 3'-mononucleotides characteristic of RNase I action are not observed'(42). Even in a mutant in which turnover of rRNA proceeds 10-fold faster than in the wild-type, the rate is the same in RNase I+ and RNase I- derivatives (43). Thus, although RNase I may participate in salvage or turnover processes, other nucleases can apparently substitute for it. 2. Application When cells are ruptured by many means (see above), RNase I is bound almost quantitatively to 30 S ribosomal subunits. The 30 S subunits are stable at Mg*+concentrationsgreater than 1 mM (44). However, when the Mg2+concentration is lower, or various monovalent cations or polyarnines are added, the rRNA in the subunits can be digested by RNase I to variable extents. Limited digestion generates nucleoprotein particles of various sizes, which are thought to result from breaks in more exposed single-stranded regions of rRNA (44-46). The small amount of RNase I attached to 50 S ribosomes will partially degrade their 23 S rRNA, even in 10-20 mM Mg", with the attendant release of four ribosomal proteins (L4, L10, L7, and L12) (47). These ribosome fragments produced by RNase I have been used to infer some features of RNA secondary structure, but other approaches have been found superior to this method (48, 49). Ill. RNase II of Escherichia coli

A.

PURIFICATION A N D

PROPERTIES

1. Puri'jication The standard RNase I1 assay is usually performed in 10 mM Tris-HC1, pH 7.5, 100 mM KCI, 1 mM MgC12, with 15 pg/ml of poly(U) at 30". One 42. Cohen, L., and Kaplan, R. (1977). J . Bncteriol. 129, 651. 43. Ohnishi, Y. (1974). Generics 76, 185. 44. Suryanarayana. T., and Burma, D. P. (1975). BBRC 65, 708. 45. Ghosh, S., and Burma, D. P. (1976). Ind. J . Med. Res. 64, 923. 46. Ghosh, S., and Burma, D. P.(1976). Ind. J . Med. Res. 64, 1680. 47. Raziuddin, Chateryi, D., Ghosh, S., and Burma, D. P. (1979).JBC 254, 10575. 48. Woese, C. R., Magrum, L. J . , Gupta, R., Siegel, R. B., Stahl, D. A,, Kop, J., Crawford, N . , Brosins, J . , Gutell, R.,Hogan, J. J., and Noller, H. F. (1980). Nucleic Acids Res. 8, 2275. 49. Ross, A., and Brimacombe, R. (1979). Nutiire (London) 281, 271.

16. RNase I , II, AND IV OF E . coii

507

unit of enzyme was defined as the amount of enzyme that renders 1 pg of poly(U) alcohol-soluble in 1 hr (8). The enzyme was partially purified by Spahr (8) and Singer and Tolbert (9). Later it was purified to homogeneity by Guptaer ul. (10). Purification is easier starting from ribosomes, but the yield is better from ribosome-free cytoplasm (8). Starting from a mixture of a 0.2 M NH4Cl wash of ribosomes and cytoplasm, it can be purified to homogeneity by 40-60% (NH&S04 fractionation, elution from DEAE at 0.2 M salt, Sephadex G150 chromatography, and 0.1 M phosphate elution from hydroxyapatite. The increase in specific activity is only 270-fold. However, the enzyme is easily inactivated, so the specific activity in purified fractions may be underestimated. 2. Physical Properties The enzyme activity requires both divalent (Mg'+ or Mn") and monovalent cations (K+ or NH:) for maximum activity. The nature of the anion also affects the activity. Thus, when enzyme is assayed at 10 m M Tris, pH 7.5, 1 mM Mg2+,and a series of different potassium salts (all 0.1 M in K+ ions), the activity relative to KCl for various anions is: acetate, 1.62; phosphate, 1.45; fluoride, 1.1; sulfate, 0.95; bicarbonate, 0.5; nitrate, 0.25; iodide, 0.05; and thiocyanate, 0.005 (8). The pH optimum is found to be between 7.0 and 8.0; at pH 9.0 and 6.0, 55 and 4% of the optimal activity was observed. The molecular weight estimation from Sephadex, sucrose gradients, nondissociating polyacrylamide gels, and dissociating gels varies from 68,000 to 85,000 and is consistent with a single protein subunit ( 8 4 0 ) . An independent report finds three subunits of MW 40,000,33,000, and 26,000 (If). Further studies are required to clarify this point. The purified enzyme is highly unstable on storage, though its stability can be improved by adding dialyzed bovine serum albumin and storing it in small portions at -80". Enzyme activity is inhibited by EDTA, singlestranded DNA, and small-to-medium size oligonucleotides (presumably, these small fragments can bind without being enzymatically degraded). RNase I1 activity is also inhibited by tRNA, but a reported inhibition by ATP (50) was probably an artifact resulting from the conversion of released nucleotides to an alcohol-insoluble form in the presence of ATP (51). Under the standard assay conditions mentioned above, but with varying concentrations of poly(U) (from 0.2 to 45 pg), the K , was found to be 4.88 x lov3g per liter, 7.5 x 10-8M (8). 50. Venkov, P., Schlessinger, D., and Longo, D. (1971). J . Brrrreriol. 108, 601. 51. Holmes, R.,and Singer, M. F. (1973). JBC. 248, 2014.

508

V. SHEN AND D. SCHLESSINGER

B. MECHANISM OF ACTION From the 3’-OH chain end, RNase I1 catalyzes the processive reaction ~ , degradation to a limiting value ofn = 3 to 5 . (pX), + pX + ( P X ) ~ - with 1. Single-Strand Specificity

With different synthetic homopolymers, the rate of enzyme cleavage is in the order poly(A) > poly(U) > poly(C) > yeast RNA. Homopolymers with substantial secondary structure, like poly(G) and poly(I), are not significantly hydrolyzed. The rate of hydrolysis of two poly(UG) copolymers (containing 43 and 63% G, respectively) are 50 and 2% of the hydrolysis of poly(A), indicating that RNase I1 is sensitive to secondary structure. Additional experiments indicating this sensitivity include a study of hydrolysis of poly(U) at different temperatures in the presence of 2,6-diaminopurine ribonucleotide. At low temperature, two strands of poly(U) interact with this nucleoside to form a triple helix, which protects poly(U) from hydrolysis by RNase 11. As the temperature is raised to disrupt the helix, poly(U) is again susceptible to RNase I1 hydrolysis (9). Similarly, a double-stranded poly(A)-poly(1) complex is resistant to RNase I1 hydrolysis; but when poly(C) is allowed to displace poly(A) from the complex, poly(A) becomes again susceptible to hydrolysis (9). It is safe to conclude that RNase I1 has a specificity for single-stranded RNA with no significant base specificity. 2 . Different Monovalent Cation Optima Within Differenl Substrates

When the KC1 concentration is varied in the presence of the same divalent cation concentration, poly(U), poly(A), poly(C), T4 mRNA, and pulse-labeled E. cofi RNA show optima at 250, 100, 50, 75, and 10-100 mM, respectively (Fig. 1). Different substrates show a differential sensitivity to hydrolysis with K + compared to Na+ as the monovalent cation (10). Whether the monovalent cation affects the secondary or tertiary structure of RNA molecules is not clear. 3. A 3’ j.5 ’ Exonuclease The primary products of the hydrolysis of polyribonucleotide are the corresponding 5’-mononucleotides. Using the polymer ( A p ) d , for example, adenosine will appear as a product only if hydrolysis starts at the 5’-hydroxyl end of the polymer. Intermediate hydrolysis products [(Ap)JA, (Ap).& (Ap)&] are found, but no adenosine is detected. Therefore, the direction of hydrolysis is from 3’ to 5‘ (52). It is of interest that 52. Nossal, N. G . , and Singer, M. F.(1968). JBC 243, 913.

509

16. RNase I, 11. AND IV OF E . coli

ADDED MONOVALENT CATION [mW

FIG. 1 . Monovalent cation requirement of RNase I1 with different substrates [ E . coli pulse-labeled RNA, T4 mRNA, and poly(U)]. The effect in increasing concentration of monovalent cations (LiCl, NaCI, KCI, NH4CI) was assayed using purified RNase 11. The trhscisstr is broken to indicate enzyme activity in the absence of added monovalent cations. From Gupta et rrl. ( 1 0 ) .

the enzyme hydrolyzes (Ap), at the same rate as (Ap)&, unlike snake venom phosphodiesterase or polynucleotide phosphorylase, which act only slowly on a phosphorylated 3' terminus. RNase I1 can attack both polynucleotides and oligonucleotides; only dinucleotides and trinucleotides are resistant to digestion.

4. Processive Mode of Degradation By using oligonucleotides labeled at the 3' terminus, the percentages of labeled material and ultraviolet-absorbing material are identical up to 100% hydrolysis. This result contrasts sharply with that found for snake venom phosphodiesterase and various E. coli DNA exonucleases that work by Michaelis and Menten kinetics. It is most easily interpreted as

510

V. SHEN AND D. SCHLESSINGER

SEPHADEX G-I00 RNAOS~

04

0

n

0%

03 02

$

ai

N 10

%

0 c

c

::

05

n

a

10%

04

~40%

03 02 01

20

40

60

80

100

120

ml

FIG. 2. Processive degradation of poly(A) by RNase 11. The products of poly(A) produced by RNase I1 and venom phosphodiesterase were chromatographed on Sephadex G-100. The amount of enzyme and times of reaction were no enzyme, 0 min (0);0.45 units, 17 min (0);0.45 units, 80 min (A). The percentage of material rendered acid-soluble is shown. The position of the peak of (Ap) is shown for reference. From Nossal and Singer (52).

reflecting a processive degradation, in which the enzyme repeatedly attacks a single polyribonucleotide chain, hydrolyzing it to small resistant oligonucleotide before releasing it. This interpretation is further corroborated by the results of sizing analysis, since after a partial digestion by RNase 11, no intermediate-size products are seen [see Fig. 2 and Ref. ( 5 2 ) ] , as expected for processive action. The significance of this processive mode of degradation is unclear. ROLEA N D POSSIBLE APPLICATION C. BIOLOGICAL 1. A Scavenger Role

RNase I1 was found originally as the major nuclease that destroyed poly(U) in a cell-free protein synthesis reaction (5). It was later implicated in degradation of T2 mRNA, T6 mRNA, and pulse-labeled E. coli mRNA

16. RNase I. 11. AND IV OF E . coli

51 1

in v i m . The specificity for single-stranded substrates and its 3‘exonuclease activity have also suggested a possible role in mRNA degradation. This ran counter to the later findings that (1) the overall direction of mRNA degradation is from 5’ to 3’, and (2) a nascent mRNA without an exposed 3’ end is also susceptible to degradation in vhw. The hypothesis of a concerted action of some endonuclease(s) and RNase I1 has been suggested several times, but, to date, the putative endonuclease(s) remains elusive, and mutants temperature-sensitive in RNase I1 (53) have shown no unequivocal change in mRNA metabolism (Lee,the chemical half-life of total cellular mRNA is actually shorter when RNase I1 is inactivated at nonpermissive temperature). Thus, any role of RNase I1 in mRNA degradation appears to be dispensible. Eschevichiu coli N464, an RNase 11-temperature-sensitive mutant strain, cannot synthesize mature ribosome or mature rRNA at nonpermissive temperatures. Partially purified RNase I1 from this strain was shown to cleave precursor 16 S rRNA into two fragments, one with the same electrophoretic mobility as mature rRNA, and the other about 100 nucleotides long. Similar results were reported using preribosomal particles, suggesting that RNase I1 plays a role in rRNA processing (12-14). However, it seemed unlikely that the exonucleolytic RNase I1 could carry out such an endonucleolytic cleavage. It was later found that the endonuclease activity could be separated from the exonuclease RNase I1 genetically (54); the suggestion has been made that strain N464 may contain a second mutation in the endonuclease (54). All of the processing steps studied that generate proper termini of rRNA appear to be endonucleases ( 5 3 , eliminating a critical role for RNase 11. When an E. coli tRNATYrprecursor molecule that contains extra nucleotides at the 3’ end was digested with a “partially” purified RNase I1 preparation in v i m , an intermediate precursor having the same nucleotide sequence as the precursor isolated from the cells was produced (15, 16). However, this activity was later separated out as RNase PIII, distinct from RNase 11. RNase I1 may be involved in the trimming of T4 tRNA precursors in viva, since an altered maturation pattern of these tRNAs is seen in RNase 11-deficientmutants (17); but another ribonuclease (RNase D) has been found to trim the precursor tRNA more precisely, and also restores the amino acid-accepting capacity in vitro. Further, the gene required for proper tRNA maturation(56 ) has been separated from RNase I1 53. 54. 55. 56.

Kinscherf, T. G., and Apirion, D. (1975). Molec. Gen. Ge~rer.139, 357. Mayhack, B . , Meyhack, I . , and Apirion, D. (1974). FEES. Lett. 49, 215 Hayes, F., and Vasseur, M. (1976). EJB 61, 433. Ghosh, R . K . , and Deutscher, M . (1978). N i d e i c . A d . 5 Rrs. 5, 3831.

512

V. SHEN AND D. SCHLESSINGER

(Y.Ohnishi, work in progress). (This work is discussed in Chapter 14 in this volume.) RNase I1 was also suggested to participate in the degradation of stable RNA under stress (57). However, the final products of RNase I1 digestion, 5'-mononucleotides, are still present in a strain with altered RNase 11, and the rate of RNA turnover is unaffected in a mutant that is deficient in RNase I1 (42). Therefore, again any action of RNase I1 can be substituted by other ribonuclease. Additional exonuclease activity has been characterized from wild-type cells and RNase 11-deficient mutants (58). All the earlier studies on roles of RNase I1 have been superseded by the discovery of more specific RNases that tend to have some isolation properties similar to RNase 11, but have more defined action on their substrates. Although RNase I1 is a major exonuclease in crude extracts, it probably does not have an indispensable role in RNA metabolism. 2. Application The instability of purified RNase I1 preparations has limited its use as a 3'-exonuclease. However, its failure to degrade the 5'-oligonucleotides of RNA chains, and its preference for RNA sequences with less secondary structure make it potentially useful in studies of RNA structure and conformation. IV.

RNase IV of Escherichia coli

A.

PURIFICATION A N D PROPERTIES

RNase IV is an endonuclease that cleaves single-stranded phage R17 RNA at limited, well-defined sites. It was obtained from an RNase Istrain (59). Nucleic acids were removed from the 100,000g supernatant fraction of a crude extract by precipitation with streptomycin sulfate and protamine sulfate. Ammonium sulfate precipitation (60% saturation) produced a precipitate that was redissolved, dialyzed in 10 mM Tris, pH 7.5, adsorbed to DEAE-cellulose, and eluted with 0.1 M KCl. This preparation, RNase IV, was lyophilized and stored frozen (59); the same preparation was active after 10 years of storage, having been thawed and refrozen twice in the interim. Like RNase I, RNase IV requires no ion supplementation, distinguishing it from RNases I1 and I11 as well as from polynucleotide phosphorylase. The enzyme is distinct, howe.ver, from RNase I, since it is 57. Lennette, E. T., Mayhack, B., and Apirion, D. (1972). FEBS Lett. 21, 286. 58. Kasai, T., Gupta, R. S . , and Schlessinger, D. (1977). JBC 252, 8950. 59. Spahr, P. F., and Gasteland, R. F. (1968). PNAS 59, 876-883.

513

16. RNase I, 11, AND IV OF E . coli U G

A C * G G . C

U ' A G - C G--A U C * G G . C U G --a , G . C G--U/ AAUCAGGCAAC CUCAACC ACUCAG ...

.

Ip

...

\I

/

FIG.3. Possible secondary structure for an intercistronic region in R17 RNA, showing RNase IV cleavage sites. The arrows indicate the positions of scissions that occur when R17 RNA is digested with RNase IV. The initiation codon (AUG) for the coat protein and three possible termination codons preceding it are indicated in bold lettering. From Adam er al. (60).

prepared from an RNase I-deficient strain and does not cleave poly(U). It is of interest that short incubations of either RNase I or RNase IV with R17 RNA yield similar large fragments. However, RNase I then cleaves further to produce a heterogeneous collection of fragments, whereas RNase IV gives rise only to the two initial large pieces from R17, MS2 or f2 RNA. RNase IV hydrolyzes RNA such as 16 and 23 S rRNA very slowly to large oligonucleotides, but releases no acid-soluble fragments from poly(&, poly(U) or poly(C). DNA is apparently not a substrate.

B. MECHANISM OF ACTION RNase IV cleaves a number of RNAs at limited sites by an unknown mechanism of recognition. 15 S and 22 S fragments are produced from 26 S R17 RNA. The 1300- to 1400-nucleotide 15 S piece yields a 5'-pppGp residue in amounts almost equivalent to the intact RNA, and therefore contains the 5' 40% of the molecule; the 2000- to 2100-nucleotide 22 S piece contains the 3' 60% of the molecule. The coding sequence for the phage gene A protein was found in the 5' fragment. Originally the cleavage was placed roughly at the start of the coat protein gene, but translation in v i m yielded coat protein from both the 5' and 3' fragments ( 2 3 ) , suggesting that the cleavage is not always at the same site. More detailed fingerprint analyses later showed that RNase IV could cleave at least five sites [see Fig. 3 and Ref. ( 6 0 ) ] , one of them located on the 5' side of the ribosome initiation complex in the coat protein gene. The multiple cleavage sites in a small region, and the nearby cleavages produced by limited action of RNase I or IV (see 60. Adam, J. M., Cory, S., and Spahr, P. F. (1972). EJB 29, 469.

514

V . SHEN AND D. SCHLESSINGE.

above), make it likely that accessibility or some other feature of RNA conformation are more important than sequence specificity in determining the sites of cleavage for the enzyme. Because the secondary structure of RNA and the features recognized by RNase IV both remain unknown, it is hard to investigate one by the other. However, some efforts have been made. With the intact RNA an amber mutant early in the coat protein gene prevents translation of the distal synthetase gene; whereas, from the RNase IV fragments of phage RNA that contains only these two genes, the distal synthetase gene can be translated. This “relief of polarity” in the fragmented RNA led to the suggestions that RNase IV destroys a specific conformational interaction between the coat protein and synthetase genes (611, and that RNase IV recognizes this specific conformation. Furthermore, subsequent electron microscopic studies of native R17 RNA and the two fragments suggested that at the proper Mg2+concentration, RNase IV cleaves at a complex looped structure that includes initiation sites for both coat protein and synthetase (62, 63). However, the enzymatic cleavage reaction is usually carried out in 5 m M EDTA, with no Mg2+added-a condition in which little higher order RNA structure was seen in the electron microscope (63). Thus, the “special” conformation putatively recognized by RNase IV remains poorly defined. Further support for the notion that RNase IV recognizes some RNA conformation comes from studies of its action on Rous sarcoma virus RNA. RNase IV does not cleave the complex 70 S viral RNA at all. However, with the two identical 35 S RNA subunits of the 70 S RNA as a substrate, RNase IV produces a heterogeneous population of RNA fragments with a mean length of 50 to 75 nucleotides (64). Since the RNA sequence in the subunits is obviously the same as that in the 70 S dimer, the differential sensitivity of the subunits is clearly related to RNA conformation. It is interesting to note that RNases I11 and IV can cleave the same proposed double-stranded loop from one specific T 1 oligonucleotide, but at different sites (64). However, because the enzyme fails to cleave many large TI marker oligonucleotides from different places in the genome, the requirement for structure remains unknown and is probably not simple. The different susceptibilities of Rous sarcoma and R17 RNAs are also puzzling; the much more extensive activity with the sarcoma virus RNA 61. 62. 63. 64.

Gesteland, R . F., and Spahr, P. F. (1969). CSHSQE 34, 707. Jacobson, A. B. (1976). P N A S 73, 307. Jacobson, A. B., and Spahr, P. F. (1977). J M B 115, 279. Darlix, J . L., Spahr, P. F., and Bromley, P. A. (1978). Virology 90, 317.

16. RNase I , 11, AND IV OF E . coli

515

occurs even at enzyme to RNA ratios 50-fold lower than used with R17 RNA. Still another indication of the importance of RNA structure for susceptibility to RNase IV comes from studies with 5 S rRNA. A single cleavage point was found in 5 S RNA in the presence o r absence of Mg2+ions at position G41 in a region thought to be a single-stranded loop (65). In summary, the experiments on site recognition by RNase IV are intriguing, but thus far they only provide a reminder that the structure of RNA is poorly understood, making it hard to understand the features recognized by enzyme probes. ROLEA N D POSSIBLE APPLICATION C. BIOLOGICAL The knowledge of this enzyme is too fragmentary to permit any guess as to the probable function, but its interesting activity at a limited number of sites in a number of RNA species suggests that it may have a role as a structure-related site-specific nuclease. The enzyme may have additional uses in studies of RNA structure, and in the preparation of defined RNA fragments. Already the limited fragmentation of R17 RNA to translatable fragments has provided a convincing gene order of 5'-A protein-coat protein synthetase. Also, cleavage by RNase IV provides one of several methods (65) to obtain two subfragments from 5 S rRNA.

65. Bellemore, G . , Jordan, B. R . , and Monier, R. (1972). J M B 71, 307

This Page Intentionally Left Blank

Polynucleotzde Phosphorylase U . 2. LITTAUER

H . SOREQ

I . Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . I1 . Properties . . . . . . . . . . . . . . . . . . . . . . . . . . . A Occurrence and Intracellular Distribution . . . . . . . . . . . B . Purification . . . . . . . . . . . . . . . . . . . . . . . . . C . Molecular Weight of Whole Enzyme and Its Subunits . . . . . . D . Amino Acid Composition and Isoelectric Point . . . . . . . . . E . Immunological Analysis . . . . . . . . . . . . . . . . . . . F. Metal Ion Requirements . . . . . . . . . . . . . . . . . . . G . Stability and Sensitivity to Proteolytic Enzymes . . . . . . . . H . Inhibitors . . . . . . . . . . . . . . . . . . . . . . . . . I . Oligonucleotide Primers and Inhibitors . . . . . . . . . . . . J . Activators and Polyamines . . . . . . . . . . . . . . . . . . I11 . The Reactions Catalyzed . . . . . . . . . . . . . . . . . . . . A . Polymerization . . . . . . . . . . . . . . . . . . . . . . . B . Nucleoside Diphosphate-P, Exchange . . . . . . . . . . . . . C . Phosphorolysis . . . . . . . . . . . . . . . . . . . . . . . D . “Transnucleotidation” . . . . . . . . . . . . . . . . . . . . IV. Attributed Physiological Functions . . . . . . . . . . . . . . . . V. Research Applications . . . . . . . . . . . . . . . . . . . . . A . Polynucleotide Synthesis . . . . . . . . . . . . . . . . . . . B . Synthesis of Oligonucleotides with a Defined Sequence . . . . . C . Polymerization of Deoxyribonucleotides . . . . . . . . . . . . D . Conjugation to Insoluble Matrix . . . . . . . . . . . . . . . E . Synthesis of Radiolabeled Nucleotides and Fingerprinting of Oligonucleotides . . . . . . . . . . . . . . . . . . . . . . F. Synchronous Phosphorolysis as an Analytical Tool . . . . . . . G . Probe for the Regulatory Function of the 3’-OH Region of RNA . H . PNPase-Directed Labeling of the 3’-OH End of Polynucleotides .

.

518 519 519 520 522 523 524 525 525 528 529 529 530 531 534 535 537 537 539 539 543 545 546 547 548 550 553

517 THE ENZYMES. VOL . XV Copyright 0 1982 by Academic Press. Inc . All rights of reproduction in any form reserved ISBN C-IZ-l22715-4

518 1.

U. 2. LITTAUER AND H . SOREQ

Introduction

Polynucleotide phosphorylase (PNPase, polyribonucleotide :orthophosphate nucleotidyltransferase, EC 2.7.7.8) was discovered by Grunberg-Manago and Ochoa during the course of a study of the mechanism of biological phosphorylation in Azotobmter vinelundii (I -3). The enzyme catalyzes the reversible reaction formulated as follows [Eq. (l)]: Mg’+

~IPPN

(PN)n + nPi

(1)

Studies of the nature of ribonucleotide incorporation into nucleic acids led to a recognition of the same reaction inEscherichia coli extracts ( 4 , 5 ) .The enzyme was also isolated from Micrococcirs luteus (formerly classified as M. lysodeikticus) (6, 7), and subsequently has been shown to be widely distributed among bacteria (8). PNPase was the first enzyme to be discovered that can catalyze the formation of polyribonucleotides with a 3‘,5‘phosphodiester bond. In the forward reaction long polyribonucleotides are synthesized from various ribonucleoside diphosphates, with elimination of inorganic orthophosphate. Each of the four common ribonucleoside diphosphates can serve separately as a substrate for the polymerization reaction, leading to the formation of homopolymers. Polymerization of a mixture of nucleoside diphosphates that contain different bases results in the formation of a random copolymer, and the enzyme does not require a template and cannot copy one. Under suitable conditions the enzyme will also catalyze the elongation of a primer oligonucleotide with a free 3’-terminal hydroxyl group [Eq. (2)] as follows: R

+

Mgz+

n(ppN)

R(PN)~+ n P ,

(2)

where R represents the oligonucleotide primer, having at least two nucleoside residues and a free 3’-terminal hydroxyl group. In the reverse reaction, the enzyme catalyzes the breakdown of polyribonucleotides by phosphorolytic cleavage of the internucleotide I. 2. 3. 4. 5. 6. 7. 8.

Grunberg-Manago, M., and Ochoa, S. (1955). FP 14, 221. Grunberg-Manago, M., and Ochoa, S. (1955). JACS 77, 3165. Grunberg-Manago, M., Ortiz, P. J., and Ochoa, S. (1956). BBA 20, 269. Littauer, U. 2. (1956). FP 15, 302. Littauer, U. Z., and Kornberg, A. (1957). JBC 226, 1077. Beers, R. F., Jr. (1956). FP 15, 13. Beers, R. F., Jr. (1956). Nntirre (London) 177, 790. Grunberg-Manago, M. (1%3). Proyr. Nucleic Acid R r s . 1, 93.

5 19

17. POLYNUCLEOTIDE PHOSPHORYLASE

bonds. The phosphorolysis reaction proceeds in a stepwise fashion starting from the 3’-OH terminus of the polyribonucleotides to liberate NDPs. PNPase also catalyzes an exchange reaction between 32P-labeled inorganic phosphate and the P-phosphate of nucleoside diphosphates [Eq. (3)l Ribonu~leoside-”P-~~P + .’‘P

Mgs ‘

R i b o n u c l e ~ s i d e - ~ ~ P+- ~31P ~P

(3)

All these reactions have served as a basis for the assay of enzyme activity. Several excellent review articles summarize the extensive work carried out with this enzyme (8-15). In the following sections we summarize the current knowledge regarding the enzyme, its purification, properties, and the various reactions it catalyzes. In particular, we will emphasize the wide range of research applications that are in use with this enzyme. II.

A.

Properties

OCCURRENCE

AND

INTRACELLULAR DISTRIBUTION

PNPase is widely distributed among different aerobic, anaerobic, and halophilic bacteria [cf. Ref. ( I S ) ] . It was also isolated from Brevibacterium (161, B. stearc~tliermophilirs(17), Thermris crqirnticus (or Tliermus thermophilw) (17. 18), and the photosynthetic bacterium Rhodospirillum rubrum ( 1 9 ) . Achromobacter sp. KR 170-4 (20) and the bacteroid form of Rhiwbium meliloti (21) seem to be relatively rich sources for the enzyme. The properties of PNPase seem to differ somewhat in various bacterial species. 9. Steiner, R. F., and Beers, R. F., Jr. (1961). “Polynucleotides, Natural and Synthetic Nucleic Acids.” Elsevier, Amsterdam. 10. Grunberg-Manago, M. (1961). “The Enzymes,” 2nd ed., Vol. V, p. 257. 11. Grunberg-Manago, M. (1962). ARB 31, 301. 12. Grunberg-Manago, M. (1963). Progr. Biopfivs. M o l w . B i d . 13, 175. 13. Singer, M. F. (1966). I n “Procedures in Nucleic Acid Research” (G. L. Cantoni and D. R. Davies, eds.), p. 245. Harper and Row, New York. 14. Thang, M. N. (1969). Bull. Soc. Cliim. B i d . 51, 1407. 15. Godefroy-Colburn, T., and Grunberg-Manago, M. (1972). “The Enzymes,” 3rd ed., Vol. 7, p. 533. 16. Yang, H. H.. Thayer, D. W., and Yang, S. P. (1979). Appl. Environ. Microhiol. 38, 143. 17. Wood, J. N., and Hutchinson, D. W. (1976). N/rc./ric Acids R e s . 3, 219. 18. Hishinuma, F., Hirai, K., and Sakaguchi, K. (1977). EJB 77, 575. 19. Soe, G., and Yamashita, J . (1980). J B 87, 101. 20. Rokugawa, K., Katoh, Y., Kuninaka, A., and Yoshino, H. (1975). A g r . B i d . Chrm. 39, 1455. 21. Hunt, R. E., and Cowles, J. R. (1977). C O NMicrobid. . 102, 403.

520

U. Z. LITTAUER AND H. SOREQ

PNPase is found in the soluble fraction of many bacterial cells ( 2 2 , 2 3 ) . Ribosomes ofE. coli contain some enzyme activity; however, most of the PNPase can be removed by repeated washing. About 10% of the total activity remains attached to washed ribosomes, probably bound to mRNA ( 2 2 ) . Some activity is also found in membrane vesicles isolated from E. coli cells ( 2 4 ) . In Streptococcus fuecalis ( 2 5 ) , S . pyogenes ( 2 6 ) , and Halohacterium cutirubrirm ( 2 7 ) , however, the enzyme is found in the cell membranes. PNPase has also been detected in wheat roots (28), and partially purified from healthy and tobacco mosaic virus (TMV)-infected tobacco leaves. However, its localization within the plant cell is uncertain (29). Partial purification of PNPase from the blue-green alga Anacysris nidufans has also been described (30). Similar activities have been reported in animal cells [cf. Refs. (15, 31)], although the results could be due to a combination of other enzymes ( 3 2 ) . Enzymatic activity that catalyzes the phosphorolysis of polyribonucleotides to NDPs has been partially purified from guinea pig liver nuclei. Unlike bacterial PNPase, the animal enzyme does not appear to catalyze the synthesis of polynucleotides (33). In addition to being associated with the nuclear membrane from rat liver cells (34, 35), PNPase activity is associated with the inner membrane of their mitochondria ( 3 4 ) . Enzymatic activity has also been detected in the endoplasmic reticulum of ribosome fraction from regenerating liver cells (35-37). B. PURIFICATION Bacterial PNPase has been purified from a wide variety of sources [reviewed in Ref. (15)l. Improved isolation procedures have increased 22. 23. 24. 25. 26. 27. 28. 29. 30. 31. 32. 33. 34. 35. 36. 37.

Kimhi, Y., and Littauer, U. Z. (1967). Biochemistry 6, 2066. Owen, P., and Salton, M. R. J. (1977). J B 132, 974. Owen, P., and Kaback, H. R. (1979). Biochemistry 18, 1413. Abrams, A., and McNamara, P. (1962). JBC 237, 170. Kessler, R. E . , and van de Rijn, I. (1979).Infect. Zmmrrn. 26, 892. Peterkin, P. I., and Fitt, P. S. (1971). BJ 121, 613. Kessler, B., and Chen, D. (1%4). BBA 80, 533. Brishammar, S. , and Juntti, N. (1974). ABB 164, 224. Capesius, I., and Richter, G . (1967). Z.Nfiturfvrschg. 22b, 204. Fitt, P. S . , and See, Y.P. (1970). BJ 116, 309. Smellie, R. M. S. (1963). Progr. Nucleic Acid Res. 1, 27. See, Y.P., and Fitt, P. S. (1970). BJ 119, 517. See, Y.P., and Fitt, P. S. (1971). FEBS Lett. 15, 65. Delvig, A. A. (1978). Biokhimiu 43, 579. Delvig, A. A., and Mardachev, S. R. (1975). Biokhimiu 40, 1246. Delvig, A. A., Tarasov. A. P.. and Debov, S. S. (1976). Biokhimiu 41. 2201.

17. POLYNUCLEOTIDE PHOSPHORYLASE

52 1

both the yield and purity of the enzyme. Essentially homogeneous preparations have been obtained from E. coli (38-42) M . luteus (43, 44), A . vineiondii (45, 461, C . pedringens (47), B . srrarothermoplzilus ( 1 7 ) , Thermus thermophilus (17, 18), and Rhodospirillum rubrum (19). Afinity chromatography on columns of poly(A)-Sepharose (48),p-aminophenyl oligo(dT)-Sepharose (491, RNA-Sepharose (42), poly(1)-agarose (50), Blue-Dextran-Sepharose (50, 51 ) and poly(U)-Sepharose (44) have yielded substantial purification of the enzyme. The effectiveness of these methods depends on prior removal of nucleic acid contaminations from the crude enzyme preparations. Phenylmethylsulfonyl fluoride has been included in solutions used for PNPase purification because of the sensitivity of the enzyme to proteolytic degradation (41, 44). Purified enzyme preparations from M. luteus (44) and B. stenrothermophilus ( 1 7 ) are virtually free of contaminating nucleic acids. Escherichia coli PNPase purified by different procedures contains low levels of bound oligonucleotides (39, 41, 42), as does the enzyme from R. rubrum (19). Most of the bacterial PNPase preparations are primer-independent forms and catalyze de novo polymerization in a processive fashion. With these enzyme forms the rate of the polymerization reaction is only slightly stimulated by oligonucleotides. Early purification of M. lufeus PNPase yielded primer-dependent preparations, in which the polymerization reaction is almost completely dependent on the presence of oligonucleotides ( 1 3 , 5 2 ) . However, with subsequent batches of cells, only the primer-independent form (form I) could be obtained (44, 5 3 ) . Primer-dependent form (form T) can be derived from the inde38. Williams, F. R., and Grunberg-Manago, M. (1964). BBA 89, 66. 39. Kimhi, Y.,and Littauer, U. Z. (1968). JBC 243, 231. 40. Kimhi, Y.,and Littauer, U. Z. (1968). “Methodsin Enzymology,” Vol. XIIB, p. 513. 41. Portier, C., van Rapenbusch, R., Thang, M. N., and Grunberg-Manago, M. (1973). EJB 40, 77. 42. Soreq, H., and Littauer, U. Z. (1977). JBC 252, 6885. 43. Letendre, C. H., and Singer, M. F. (1975). Nucleic Acids Res. 2, 149. 44. Barbehenn, E. K., Craine, J. E., Chrambach, A., and Klee, C. B. (1982). JBC 257, 1007.

45. Gajda, A. T., Zaror de Behrens, G., and Fitt, P. S. (1970). BJ 120, 753. 46. Mii, S. (1977). J B 81, 899. 47. Guissani, A. (1978). Biochimie 60,755. 48. Lehrach, H., and Scheit, K. H. (1972). Hoppe Seyyler’s Z. Physiol. Chem. 353, 731. 49. Smith, J. C., and Eaton, M. A. W. (1974). Niicleic Acids Res. 1, 1763. 50. Drocourt, J. L., Thang, D. C., and Thang, M. N. (1978). EJB 82, 35.5. 51. Thang, M. N., Drocourt, J. L., Chelbi-AEx, M. K., Thang, D. C., Lubochinski, J., Ruet, A., Sentenac, A., Gangloff, J., and Dirheimer, G. (1979). Cmlloq. Inserm. Afiniry Chromatogr. 86, 303. 52. Singer, M. F., and Guss, J. K. (1962). JBC 237, 182. 53. Klee, C. B. (1967). JBC 242, 3579.

522

U . Z. LITTAUER AND H. SOREQ

pendent form by limited tryptic digestion (54,55). Following trypsin digestion, A. vinelandii PNPase also develops primer requirement (45). PNPase preparation purified from B. stearothermophilus (I 7) and T. thermophilus (18) show primer dependency probably due to endogenous proteolysis.

c.

MOLECULAR WEIGHT OF WHOLE ENZYME ITS SUBUNITS

AND

The molecular weight of the whole enzyme has been determined by sedimentation equilibrium, gel filtration, sucrose gradient centrifugation, and gel electrophoresis under nondenaturing conditions. In the latter method, enzyme activity can be visualized after electrophoresis by incubating the gels in the presence of ADP and Mg‘+, followed by staining the poly(A) formed in situ with acridine orange in the presence of lanthanum chloride (56). Other in sitir methods for visualizing active enzyme molecules have also been published (23, 45, 57). The physicochemical properties of E. coli form A (4f142,58) and M. luteus form I PNPase are similar (44).The molecular weight of purified E. coli PNPase determined by sedimentation equilibrium ranges between 230,000 +- 20,000 (42) and 216,000 I+_ 20,000 (4f), as compared to 237,000 24,000 for the M. iuteus enzyme (44). A value of 252,000 has been calculated from a Stokes radius of 6.4 nm and a sedimentation constant of 8.9 S for theE. coli enzyme (58). The observed frictional ratio is 1.52 (58). The E. coli enzyme is composed of three identical subunits of a molecular weight ranging between 84,000-95,000 ( 4 I , 42, 48, 58, 5 9 ) . Support for the a3 structure of E. coli PNPase arises from ultrastructural observations. Under the electron microscope, the enzyme appears as a triangle with a central hole. The diameter of these molecules was calculated to be 85 A (60). In crude cell extracts PNPase displays microheterogeneity . Sucrose gradient sedimentation, gel filtration, and gel electrophoresis all show the presence of higher level components ( M , 39, 4f.43, 44,54,58). These forms arise from the association of an additional polypeptide subunit (41. 58) or the presence of bound nucleic acids that

*

54. Klee, C. B. (1969). JBC 244, 2558. 55. Klee, C. B. (1971). I n “Procedures in Nucleic Acid Research” ( G . L. Cantoni and D. R. Davies, eds.), Vol. 2, p. 896. Harper and Row, New York. 56. Thang, M. N . , Thang, D. C., and Leautey, J. (1967). C . R . Acad. Soc. ( P a r i s ) 265, 1823. 57. Fitt, P. S ., Fitt, E. A., and Wille, H. (1968)’. BJ 110, 475. 58. Portier, C. (1975). EJE 55, 573. 59. Portier, C. (1975). FEBS Lett. 50, 79. 60. Valentine, R. C., Thang, M. N . , and Grunberg-Manago, M. (1969). J M B 39, 389.

17.

POLYNUCLEOTIDE PHOSPHORYLASE

523

induce a conformational change in the enzyme (44). PNPase from E. coli can be isolated in two active forms, A or B, having molecular weights of 252,000 and 365,000, respectively. The A form has an a3type structure, whereas the B form has two types of chains, a (MW 86,000) and /3 (MW 48,000). The exact proportion of the a and p subunits is not yet clear, and PNPase B form has been assigned a structure of a& (or a3Pn). The B form is obtained by keeping the ionic strength at 0.2 M during the purification of the enzyme on a Sephadex G-200 column, whereas at lower salt concentrations the /3 subunit tends to dissociate and the enzyme reverts to the A form. All the catalytic activity of PNPase resides in the a subunits, whereas the p subunit is inactive and does not alter the enzymatic properties of the whole enzyme. The role of the p subunit therefore remains to be determined (58). In addition to the main 252,000 MW form, E . coli B and K12 extracts contain 25 and 596, respectively, of a low molecular weight (100,000) PNPase. The 100,000 MW form catalyzes the phosphorolysis reaction but is unable to catalyze the polymerization of NDPs. The 100,000 form differs from the main 252,000 MW enzyme in that it can only phosphorolyze short-chain polymers and requires higher Mg2+ ion concentrations. PNPase preparations from E. coli Q13 and 1 1 13 mutants are particularly rich in this defective enzyme, and about 80% have a molecular weight of 100,000. In addition, about 20% of the mutant PNPases have a molecular weight of about 200,000. Unlike the 100,000 MW form, or the wild-type enzyme, this additional form requires Mn2+for NDP polymerization and has a higher K , for poly(A) phosphorolysis (61). Clostridirrm per-ingens PNPase also appears in two forms, a& and a3, with molecular weights similar to that of the E. coli enzyme (47). PNPase from A . vinelmdii has an apparent molecular weight of 200,000 (46). But the R. ruhrum enzyme has an MW of 160,000, and appears as a dimer of two subunits of 76,000 (19). PNPase from B . stearothermophilus (17)is a tetramer of 51,000, and that from T. thermophilus (18) shows three subunits of 92,000, 73,000, and 35,000, which may result from limited proteolysis of the enzyme.

D. A M I N OACID COMPOSITION

AND

ISOELECTRIC POINT

The amino acid composition of the E. coli PNPase (42, 62) is similar to that of the M . lrrteirs enzyme (43.44). Although there are some differences in the reported cysteine and tryptophan values for the E. coli enzyme (42, 61), it is likely that it contains 3 Cys and 3 Trp residues per mole of subunit 61. Thang, M. N . , Thang, D. C . , and Grunberg-Manago, M. (1969) EJB 8, 577. 62. Portier, C. (1975). Biocliirnie 57, 545.

524

U. Z. LITTAUER AND H. SOREQ

of 84,000 MW, as does the M. lureus enzyme (44). Of the 3 Cys residues of the E. coli subunit, only one group is exposed, and is found to react with dithionitrobenzene, whereas 2 groups are “masked” and react only after denaturation with 1% sodium dodecyl sulfate (42). This property may explain the insensitivity of the enzyme to -SH reagents (39). The UV difference spectrum of the M. luteus enzyme suggests that 6 Tyr residues (out of 17), and perhaps 1 Trp residue (out of 3), are buried in the interior of the protein and become exposed upon treatment with 6 M guanidineHCl(44). One Trp and three Tyr residues are lost during the conversion of M. luteus form I to form T, which may explain why form T has lost both the ability to bind oligonucleotides with high affinity and to catalyze de novo synthesis of poly(A) (44). The a and /3 subunits of E. coli PNPase appear to be unrelated to each other, and differ in their amino acid composition as well as in their cyanogen bromide cleavage peptides (62). The N-terminal amino acid sequence of the E. coli a form suggests that the a chains are all identical and terminate with Met-Leu-?-Pro-Phe (62). Methionine is also the only amino acid found at the N-terminus of B. srenrothermophilus enzyme (17). In contrast, neither the I nor the T form of the M. luteus enzyme contains detectable free amino end groups. Since the primer-dependent T form is obtained by limited trypsin digestion, it is suggested that proteolysis removes a peptide at the carboxy end of the molecule (44). In situ staining of isoelectric focusing gels revealed an isoelectric point of 6.1 for PNPase from E. coli (42) and M . luteus (44), whereas the enzyme preparations isolated from B. stearothermophilus (17) and T. thermus (18) focus at pH 4.1 and 4.3, respectively.

E.

IMMUNOLOGICAL ANALYSIS

Antibodies against purified PNPase from E. coli B were shown to react with the enzyme in a double diffusion test and in immunoelectrophoretic analysis. The enzyme, complexed with its antibodies, retains its polymerization properties, and the antigen-antibody complex can be visualized by autoradiography of the polynucleotide formed in situ by the enzyme (63). Double precipitation bands were obtained with enzyme purified according to Williams and Grunberg-Manago (38, 63). However, rabbit antibodies elicited against homogeneous E. coli PNPase, following the affinity chromatography step, displayed a single precipitation band (42). No serological relationship exists between E. coli PNPase and either the core enzyme or the L+ subunit of E. coli RNA polymerase (42), in 63. Uriel. J., Thang, M. N., and Berges, J. (1969). FEES Lett. 2, 321.

525

17. POLYNUCLEOTIDE PHOSPHORYLASE

contradistinction to an earlier suggestion (64). The ribosomal S 1 protein, which contaminates E. coli PNPase, is also unrelated to PNPase (42). F. METALION

REQUIREMENTS

Many studies indicate that Mg" is required for the reactions catalyzed by PNPase and that it can be partially replaced by Mn'+ (5, 15).Free Mgz+ ions bind to E. coli PNPase with a K,,, of 5 x M (65). Other cations, such as Co2+,Ni", Cd2+,Cu2+,and Zn2+,but not Ca'+, may also replace Mg2+in PNPase reactions, although with quite different efficiencies (19, 39, 66). Polymerization of GDP with E. coli PNPase, however, proceeds efficiently in the presence of Mn" at 60" (67). The polymerization reaction with a mutant PNPase from E. coli 413 requires Mn2+rather than Mg'+ (68), and Mn2+will stimulate more efficiently than Mg'+ the polymerization reaction with PNPase from Achromohucter (20). If, indeed, PNPase plays a role in the nucleolytic degradation of RNA (69), the inability of Ca2+to replace Mg2+in the phosphorolysis reaction with E. coli PNPase may partially contribute to the protective effect that Ca2+exerts in vitro on various types of mRNA (70). However, at low Ca2+concentration, of about 5 p M , there is a threefold activation of the polymerization reaction with B. steurothermophilus enzyme (17). At suboptimal Mg" concentrations, both the formation of polymers from NDPs and the NDP-Pi exchange reaction occur only after an initial lag period. In the presence of polynucleotides or short oligonucleotides, this lag period is almost abolished (38, 71-73).

G.

STABILITY A N D SENSITIVITY TO PROTEOLYTIC

ENZYMES

Purified E. coli PNPase is unstable above 55", and is rapidly and irreversibly inactivated at 65" (15, 39, 42). The M. I u t e ~ s(74) and the C. per64. Ohasa, S . , Tsugita, A., and Mii, S . (1972). Nature N e w B i d . 240, 39. 65. Williams, F. R . , Godefroy, T., Mery, E . , Yon, J . , and Grunberg-Manago, M. (1964). BBA 80, 349. 66. Babinet, C., Roller, A., Dubert, J. M., Thang, M. N., and Grunberg-Manago, M. (1965). BBRC 19, 95. 67. Thang, M. N., Graffe, M . , and Grunberg-Manago, M. (1965). EEA 108, 125. 68. Hsieh, W. T., and Buchanan, J. M. (1967) PNAS 58, 2468. 69. Kaplan, R . , and Apirion, D. (1974).JBC 249, 149. 70. Cremer, K . , and Schlessinger, D. (1974). JEC 249, 4730. 71. Ochoa, S . , and Mii, S. (1961). JBC 236, 3303. 72. Mii, S . , and Ochoa, S. (1957). EBA 26, 445. 73. Singer, M. F., Heppel, L. A., and Hilmoe, R. J. (1957). BBA 26, 447. 74. Brenneman, F. N., and Singer, M. F. (1964). EBRC 17, 401.

526

U . Z. LITTAUER AND H. SOREQ

fringens enzyme (75) are less stable than E. coli PNPase. The enzyme is stabilized against heat inactivation by the presence of NDPs, but not by NMPs, NTPs, or DNA. Substrate oligonucleotides with free 3'-OH termini can also exert this protective effect, whereas oligonucleotides with blocked 3' ends do not affect the rate of heat inactivation (76). Heatdenatured E. coli PNPase can be renatured. Following heating at 100" for 1 min the precipitate is dissolved in 6 M guanidine-HC1followed by dialysis. About 25-30% of the original enzyme activity is recovered by this procedure, and the reassociated enzyme reverts to its original quaternary as structure (41). High concentrations (>3.0 M) of urea have also been shown to cause inactivation of E. coli PNPase. In this case as well, the presence of substrates protects the enzyme against the inactivation process (77). PNPase is sensitive to proteolytic digestion. Earlier studies revealed differences in subunit structure and catalytic properties of the enzyme when isolated from various bacterial sources. It now appears that these differences mainly result from endogenous proteolytic digestion in various enzyme preparations, and that the properties of the various intact enzyme preparations are similar. Degradation by endogenous proteases of PNPase from M. luteus (53),A . ngilis (45), C. pegringens (47, 78), and E. coli (79, 80), or degradation with chymotrypsin or trypsin (45, 47, 54, 57, 79-81) yield very close gel electrophoretic patterns. Limited proteolysis of the enzyme supports the view that the catalytic center and the polynucleotide binding subsite (82-86) are distinct and dispersed over the enzyme surface. Storage of E. coli PNPase for extended periods at 4" results in limited proteolysis of the enzyme. The proteolyzed PNPase has a reduced molecular weight of 175,000 with an a$ structure (a' = 65,000). The endogenous proteolysis induces changes in both the phosphorolysis and polymerization reactions (79, 80). The K , 75. Fitt, P. S . , Dietz, F. W., Jr., and Grunberg-Manago, M. (1968). BBA 151, 99. 76. Lucas, J . M., and Grunberg-Manago, M. (1964). BBRC 17, 395. 77. Harvey, R. A . , Godefroy, T., Lucas-Lenard, J., and Grunberg-Manago, M. (1967). EJB 1, 327. 78. Guissani, A., and Grunberg-Manago, M. (1969). BBRC 35, 131. 79. Thang, M. N . , Dondon, L., and Godefroy-Colburn, Th. (1971). Biochimie 53, 291. 80. Guissani, A . , and Portier, C. (1976). Nucleic Acids Res. 3, 3015. 81. Fitt, P. S . , and Wille, H. (1969). BJ 112, 497. 82. Chou, J . Y . , and Singer, M. F. (1970). JBC 245, 995. 83. Thang, M. N . , Guschlbauer, W., Zachau, H. G . , and Grunberg-Manago, M. (1967). J M B 26, 403. 84. Thang, M. N . , Harvey, R. A., and Grunberg-Manago, M. (1970). JMB 53, 261. 85. Godefroy, T. (1970).EJB 14, 222. 86. Chou, J. Y . , Singer, M. F., and McPhie, P. (1975). JBC 250, 508.

17. POLYNUCLEOTIDE PHOSPHORYLASE

527

to for a polynucleotide in the phosphorolysis reaction shifts from M , indicating that proteolysis causes a loss of the polynucleotide binding site. The proteolyzed enzyme shows a much more stringent requirement for an oligonucleotide primer in the polymerization reaction but is not stimulated by polynucleotides. Because of the loss of polynucleotide binding sites, phosphorolysis of poly( A),U with the proteolyzed enzyme proceeds with a partially nonprocessive mechanism, as opposed to the processive phosphorolysis displayed by native enzyme. It has been assumed (80) that the polymerization mechanism as well will no longer be purely processive, and that the mean length of the polymers synthesized will be shorter than that observed for polymers obtained with native enzyme. The proteolyzed enzyme also fails to bind to polynucleotide-agarose or Blue Dextran-agarose columns (SO, 5 1 ) . As with native enzyme (87), phosphorylation of proteolyzed E. coli PNPase by cyclic AMP-dependent protein kinase can replace the stimulating effect of oligonucleotides in the polymerization reaction proteolyzed (79). Similar changes in the properties of E . coli PNPase were also produced by incubating the enzyme with isolated bacterial proteases (79, 88). Native PNPase from M . luterrs is primer-independent, catalyzes de n o w polymerization in a processive fashion, and is only slightly stimulated by oligonucleotides. Limited trypsin digestion of the native enzyme alters its polymerization activity without affecting its ability to phosphorolyze polynucleotides. The trypsinized enzyme (form T) catalyzes the elongation of primer by a random mechanism and is stimulated up to 20-fold by oligonucleotides (54, 57, 89 -91 ). Restoration of primerindependence to form T can be obtained by treatment with Pmercaptoethanol. Reconversion to primer dependence is achieved by reaction with sulfhydryl inhibitors, suggesting that the alteration in the enzyme properties is correlated with the modification of sulfur-containing amino acids (54, 92). Enhancement in primer requirement also appears in PNPase from Azorobacter tinehzdii upon mild treatment with trypsin or aging of the enzyme (45, 49, 93). Restoration of the reduced activity, but not the loss of primer requirements, is caused in this case as well by P-mercaptoethanol. 87. Thang, M. N . , and Meyer, F. (1971). FEES Lc,tr. 13, 345. 88. Regnier, Ph., and Thang, M. N. (1972). Biocliirnie 54, 1227. 89. Moses, R . E., and Singer, M . F. (1970). JBC 245, 2414. 90. Klee, C. B . , and Singer, M. F. (1968). JBC 243, 923. 91. Fitt, P. S . , and Fitt, E. A. (1967). BJ 105, 25. 92. Klee, C . B . , and Singer, M. F. (1968). JBC 243, 5094. 93. Gajda, A. T., and Fitt, P. S. (1969). BJ 112, 381.

528

U . Z. LITTAUER AND H . SOREQ

PNPase from C. perfringens is highly susceptible to proteolysis and is obtained as a mixture of variable proportions of native and proteolyzed forms. Under the action of either endogenous proteases or trypsin, two enzymatic forms are obtained that differ in their catalytic properties from each other and from the initial enzyme. One of the proteolyzed species catalyzes polymerization only in the presence of poly(A) or polylysine, whereas the other phosphorolyzes oligonucleotides but not polynucleotides (47). In contrast to the native enzyme, the proteolyzed enzyme requires P-mercaptoethanol and polylysine for efficient polymerization activity (47, 7 8 ) .

H. INHIBITORS Several chemical agents have been shown to block the catalytic activity of PNPase from various biological sources. Some of these, such as 6-azauridine or 5-fluorouridine diphosphates (94), as well as phosphonic acid analogs of ADP (95, 96) or analogs produced by periodate oxidation (97),appear to react with the active site and inhibit the exchange, the phosphorolysis, and the polymerization activities of the enzyme. Inhibitory reaction has also been noticed for deoxynucleoside diphosphates (98, 99). Other inhibitors, such as acridine orange, appear to inhibit the polymerization reaction via their interaction with the primer oligonucleotide (100, f 01). The catalytic activity of PNPase from B. amyloliquefuciens has been reported to be inhibited by heparin, rifamycin SV, and synthetic polynucleotides (102); the polymerization reaction catalyzed by E. coli PNPase is effectively inhibited by oligophosphates of pyridoxal, the percentage of inhibition being higher with longer chains of phosphate moieties bound to the pyridoxal core (103). 94. Skoda, J., Kara, J . , Sormova, Z., and Sorm, F. (1959). BBA 33, 579. 95. Simon, L. N . , and Myers, T. C. (l%l). BBA 51, 178. %. Godefroy-Colburn, T., and Setondji, J. (1972). BBA 272, 417. 97. Smrt, J., Mikhailov, S. N . , Hynie, S. , and Florentev, V. L. (1975). Collect. Czech. Chem. Commun. 40, 3399. 98. Lucas-Lenard, J . , and Cohen, S. S. (1966). BBA 123, 471. 99. Bon, S., Godefroy, T., and Grunberg-Manago, M. (1970). EJB 16, 363. 100. Beers, R. F.,Jr.. Hendley, D. D., and Steiner, R. F. (1958). Nature (London) 182, 242. 101. Beers, R. F., Jr. (1%0). JBC 235, 726. 102. Erickson, R. J . , and Grosch, J. C. (1977). J B 130, 869. 103. Mamaeva, 0. K., Karpeiskii, M. Ya., Karpeiskii, A. M., and Bibilashvili, R. Sh. (1979). Molek. B i d . 13, 811.

17. POLYNUCLEOTIDE PHOSPHORYLASE

I.

529

OLIGONUCLEOTIDE PRIMERS A N D INHIBITORS

The de n o w polymerization of NDPs, particularly at low Mg2+concentration, is preceded by a lag period, which may be overcome by the addition of polynucleotides or short oligonucleotide primer molecules with a free 3’-hydroxyl group (38,39, 71-73). These primers also accelerate the exchange of phosphate moieties by the enzyme (39, 73, 104). The oligonucleotide primers have been shown to be incorporated into the polymer synthesized by PNPase from M. luteus (105), and their effect was found to be maximal in the polymerization of GDP, which proceeds with difficulty and at a slow rate in the absence of such primers (106). When blocked with a 3’-terminal phosphate moiety, oligonucleotides act as inhibitors of the polymerization and the exchange reactions (71, 72, 107). The inhibition is temperature-dependent and may be overcome by addition of a complementary polynucleotide, which hybridizes with the oligonucleotide inhibitor and prevents its binding to the enzyme (108). The strong binding of blocked polynucleotides has been exploited to develop affinity chromatography procedures to purify the enzyme (42 ).

J. ACTIVATORS A N D POLYAMINES The polymerization reaction catalyzed by PNPase has been reported to be activated by several agents. Potassium, sodium, and lithium salts have been shown to affect the K, values of the M. luteus enzyme (109). A basic polypeptide that enhances the ADP-Pi exchange reaction has been isolated from E. coli extracts (5, 39). In the presence of this heat-stable activator, the optimal Pi concentration for ADP-Pi exchange shifts from 2 to about 0.65 mM. At low phosphate concentrations, the activator causes up to 3- to 6-fold stimulation of the exchange reaction withE. coli PNPase, but has no effect on the rate of polymer formation or the phosphorolysis of poly(A). The activation of the exchange reactions with NDPs other than ADP is much lower than with ADP. Spermine and spermidine (0.1- 1.0 mM) also activate the ADP-Pi exchange (twofold), whereas poly-L-lysine and poly-L-ornithine M ) hardly affect the reaction, and at higher concentrations cause inhibition (89). A basic protein from A. vinelandii 104. 105. 106. 107. 108. 109.

Beers, R . F., Jr. (1961). JBC 236, 2703. Singer, M . F., Heppel, L. A . , and Hilmoe, R . J. (1960). JBC 235, 738. Brenneman, F. N . , and Singer, M. F. (1964). JBC 239, 893. Beers, R . F., Jr. (1959). Nufitre (London) 183, 1335. Heppel, L . A . (1963). JBC 238, 357. Beers, R . F., Jr. (1957). Nutitre (London) 180, 246.

530

U. Z. LITTAUER AND H. SOREQ

causes a lag phase in the NDP polymerization reaction. Preferential repression of polymerization of UDP is observed with polylysine and of ADP with polyarginine. A lag phase is also caused by polylysine in the ADP-Pi exchange reaction with the A . vinekcindiii enzyme (46). The stimulating activity of polylysine and other polyamines on the ADP polymerization with proteolyzed C. perfringens enzyme has been noted (47, 75, S l ) , and is probably due to charge effects on the purified protein (1 10). A different mode of activation of the polymerization reaction is exerted by acridine orange, which forms complexes with the newly synthesized polynucleotides in the reaction mixture and, changing the equilibrium constants, drives the reaction toward further polymerization (100. 101). Yet another mechanism of activation has been observed with ATP, which improves the yields of polynucleotides synthesized with crude extracts from Azotobcicter vinelcindii (1 11). Activation by ATP has been suggested to function via the phosphorylation of PNPase by CAMP-dependent protein kinase (87). It should, however, be noted that these studies were mostly carried out with partially purified enzyme preparations, and that the effects observed could result from combined changes in the activities of contaminating enzymes.

111.

The Reactions Catalyzed

PNPase catalyzes the phosphorolysis of long-chain polynucleotides in a processive mechanism [also denoted progressive, see Ref. (IS)],whereby the enzyme does not dissociate from the polymeric substrate during the degradation process. Thus, the enzyme appears to degrade one polymer chain to completion prior to releasing a small resistant oligonucleotide and initiating phosphorolysis of another chain (15, 83, 90). In contrast, short oligonucleotides are degraded by a random nonprocessive mechanism [also denoted synchronous, see Ref. ( I S ) ] in which the enzyme dissociates from the substrate after hydrolysis of each nucleotide (82, 112). In addition, PNPase catalyzes the de nuvo polymerization of NDPs to polynucleotides in a processive mechanism, whereas elongation of oligonucleotide primers may occur by a nonprocessive mechanism [cf. Ref. (15)]. The diversity of mechanisms seems to be due to the existence of two classes of 110. Fitt, P. S., and Wille, H. (1969). BJ 112, 489. 1 1 1 . Shiobara, Y., and Itagaki, K. (1963). J B 54, 317. 112. Singer, M. F., Hilmoe, R. J., and Grunberg-Manago, M. (1960). JBC 235, 2705.

17. POLYNUCLEOTIDE PHOSPHORYLASE

53 I

binding sites in the molecule (84-86, 113-115). The first site, subsite I, is the catalytic center of the molecule and includes the mononucleotide, inorganic phosphate, and oligonucleotide binding domains. Subsite I binds the 3’-OH terminus of the growing polynucleotide or oligonucleotide. When polynucleotides are long enough, they can reach a second site, subsite 11, which is not involved in the phosphorolysis of oligonucleotides. Subsite I1 probably includes several polynucleotide binding domains (116). It has also been suggested that subsite I1 includes a lysinerich area that may act as a regulatory site (46). The residence time of the polymer in subsite I is short, whereas in subsite I1 it is long and corresponds to a very strung affinity of the enzyme for polynucleotides (lO-*M to M ) . This dual attachment to the enzyme allows long polynucleotides to snap back to a reactive position after removal of one nucleotide residue and thereby be degraded in a processive manner. Thus, binding to subsite I1 is responsible for the marked enhancement in the binding of polynucleotides. Oligonucleotides, not being anchored at subsite 11, are lost into the solution after the reaction and released from subsite I, and may then be replaced by another substrate molecule, leading to a nonprocessive random mechanism (15, 85, 86, 116). Limited proteolytic degradation (80, 1/.5), as well as linking the E. coli enzyme to BrCNactivated Sepharose ( I 17), affects mainly the polynucleotide binding domains of subsite 11, and results in the loss of complete processiveness and the decrease of affinity for polynucleotides. The active center is preserved, however, indicating that subsite I is in some way hidden. These findings are in agreement with the a3 subunit structure, which envisages the enzyme to have a triangle profile with a central hole in which the active center might be located (59, 60, 117). A.

POLY M E RIZ AT I ON

1. Initiritioti of de Novo Synthesis

PNPase catalyzes de novo synthesis of polynucleotides. The mechanism of formation of the first internucleotide bond is still unclear and probably involves the reaction between two NDP molecules, out of which one serves as the accepting 5’ terminus. Analysis of the newly synthe113. 114. 115. 116. 117.

Kaufmann, G., and Littauer, U. 2. (1969). FEES L d r . 4, 79. Chou, J . Y., and Singer, M. F. (1970). JBC 245, 1005. Guissani, A. (1977). EJE 79, 233. Godefroy, Th., Cohn, M., and Grunberg-Manago, M. (1970). EJB 12, 236. Vang, N . H . , Drocourt, J. L., and Thang, M . N. (1979). BBRC 90, 606.

532

U. Z. LITTAUER AND H. SOREQ

sized polynucleotides reveals that they contain, at the 5’ terminus, a monophosphate group rather than the expected pyrophosphate group (1 18). It is possible that a 5’-pyrophosphate terminus is initially formed, followed by removal of the @phosphate (or NDP) from the 5’ terminus at a later stage in the reaction. A novel mechanism suggests the transfer of the p-phosphate of ADP such that the AMP product formed can be positioned on PNPase as the 5’-monophosphate terminus of the nascent poly(A) chain. This transfer could depend on the deoxyadenylate kinase activity that is associated with PNPase from M. luteus (1 19). It should be noted that ApA and pApA do not undergo phosphorolysis, and accumulate as resistant end products of poly(A) phosphorolysis (1 14, 120-122). This would imply that the initiation of de nova polymerization involves an initial irreversible step. 2 . Elongation The processive elongation of polynucleotides by PNPase proceeds at a linear rate and then reaches a plateau. The polymers formed are of high molecular weight and are homogeneous in size; no intermediate oligonucleotides are formed (84). On the basis of kinetic analysis, the presence of a transient intermediate polynucleotide of high molecular weight, which is subsequently degraded to an equilibrium mixture of short oligonucleotides, has been proposed for primer-dependent PNPase (123). The complex enzyme-polynucleotide does not dissociate during the elongation process, even when unfavorable substrates such as GDP are polymerized at high temperature and in the presence of MnZ+ (67). PNPase utilizes the Sp diastereomer (exoisomer) of NDPS as a substrate, whereas the Rp isomer is a competitive inhibitor. During polymerization an inversion occurs in the configuration of the phosphorous bond into the Rp type (endoisomer), as was shown by high performance liquid chromatography of uridine 2’ ,3’-cyclic phosphorothioate, enzymatically obtained from copolymers of UDP with adenosine 5’-0-(1thiodiphosphate) (124). 118. 119. 120. 121. 122. 825. 123. 124.

Harvey, R. A., and Grunberg-Manago, M. (1966). BBRC 23, 448. Craine, J. E., and Klee, C. B. (1976). Nucleic Acids Res. 3, 2923. Singer, M. F. (1958). JBC 232, 211. Madison, J. T., Everett, G. A., and Kung, H.-K. (1967). JBC 242, 1318. Madison, J. T., Holley, R. W., Poucher, J. S., and Connett, P. H. (1967). BBA 145, Cantor, C. R. (1968). Biopo/ymers 6, 369. Burgers, P. M. J . , and Eckstein, F. (1979). Biochemistry 18, 450.

17. POLYNUCLEOTIDE PHOSPHORYLASE

533

3. Equilibriiim PNPase directs either phosphorolysis of polynucleotides or polymerization of NDPs, depending on the reaction conditions and on the concentration of these two components in the reaction mixture. The mechanisms by which the enzyme drives these two reactions have been studied extensively with PNPase from M . futeus (82, 86) and from E . coli (85, 116), and detailed models have been proposed to explain the interrelations between various kinetic parameters that affect the dynamic equilibrium reached by the enzyme. The affinity of M . lirtrirs PNPase for either inorganic phosphate or oligonucleotide substrate is unaffected by the presence of either, and the initial rate of phosphorolysis depends linearly on the concentration of both. Oligonucleotides in which the 3'-OH group is blocked with a phosphate group are competitive inhibitors with respect to unblocked oligonucleotides, and noncompetitive with respect to inorganic phosphate. In contrast, the kinetics of phosphorolysis of polynucleotides shows that dADP is a competitive inhibitor with respect to both Pi and polynucleotide (86). Copolymerization of various NDPs occurs with M . lufeirs PNPase in a random fashion, indicating no special preference for any of the four common NDP substrates (f2.5). 4. Modified Sirbstrutes

Modifications of the NDP substrates serve to characterize the catalytic processes driven by the enzyme, the specificity of substrate recognition, and the properties of the active sites. Thus, blocking of the NDP at the 3' position yields a monovalent substrate, of which only one residue may be added to an oligonucleotide primer (126-128). It was shown that deoxynucleoside diphosphates are added to the 3' terminus of an oligonucleotide to a limited extent. The reason the polymerization of dADP cannot proceed readily seems to be due to the low affinity of the catalytic center (subsite I) to the DNA-like internucleotide linkage. When deoxyadenyl residues are added to the growing end of the chain, its a f h ity to the enzyme is lowered and the rate for further elongation is hence greatly reduced (I 13). Deoxyribonucleotides also act as inhibitors of PNPase. dADP inhibits competitively both the polymerization of ADP and the phosphorolysis of polynucleotides (86, 99), indicating that the 125. Seliger, H., and Knable, T. (1978). Nucleir Acids R r s . , Spec. Public. 4, S167. 126. Kaufmann, G., and Littauer, U. 2. (1970). EJB 12, 85. 127. Kaufmann, G., Fridkin, M., Zutra, A., and Littauer, U. 2. (1971). EJB 24, 4. 128. Bennett, G . N., Mackey, J. K . , Wiebers, J. L., and Gilham, P. T. (1973).Biochernistry 12, 3956.

534

U . Z. LITTAUER AND H.SOREQ

oligonucleotide primer covers the NDP binding subsite. However, when Mg2+is replaced by Mn2+,dADP is capable of copolymerizing with ADP (129). Other analogues, such as the periodate oxidation product of ADP, will block polymerization altogether (97) (see previous sections). PNPase displays a rather low specificity with regard to side chains on the purine or pyrimidine moieties (see previous sections), whereas it shows high specificity with respect to the number of phosphate groups on the nucleoside and the nature of the sugar moiety of the NDP substrate [cf. Ref. S ) ] . The polymerization parameters for various modified bases also serve to detect functional differences between PNPase from various strains of bacteria (130).

B. NUCLEOSIDE DIPHOSPHATE-P, EXCHANGE Two mechanisms were suggested for NDP-Pi exchange reaction: (1) The observed exchange reflects a reversible formation of a covalent, nucleoside monophosphate-enzyme complex, or (2) the apparent exchange is a result of combined polymerization and phosphorolysis reactions, occurring under approximate equilibrium conditions (8,39).The kinetic parameters of the exchange reaction appear to be similar to those of the polymerization reaction: It is preceded by a lag phase, activated by primers (8, 39), and occurs, to a limited extent, with deoxy NDPs, but only in the presence of oligonucleotide primers or NDPs (99, 113, 131). It was suggested that the use of dADP might facilitate isolation of the putative NMP-enzyme intermediate (113). However, no evidence for its formation could be obtained (131). Further support that the NDP-Pi exchange is the result of combined polymerization and phosphorolysis reactions is suggested from the arsenolysis of NDPs. Replacement of Pi by arsenate in the exchange reaction results in the arsenolysis of NDPs to nucleoside monophosphates. In the presence of primer-dependent PNPase from M. Iiiteus, arsenolysis of ADP, like its polymerization, is activated by oligonucleotides that have unesterified 3'-hydroxyl groups (116, 132). The kinetics of this reaction are consistent with the formation of a ternary complex between enzyme, oligonucleotide, and NDP. The formation of a new phosphodiester bond between the NDP and oligonucleotide and its subsequent arsenolysis is proposed for this reaction (132). A similar exchange reaction is catalyzed by a yeast ADP-sulfurylase, which does not show specificity for the sugar moiety, the nature of the NDP substrate, or 129. 130. 131. 132.

Chou, J. Y., and Singer, M. F. (1971). JBC 246, 7505. Swierkowski, M., and Shugar, D. (1969). Actu Biocliim. Polon. 16, 263. Chou, J. Y . , and Singer, M. F. (1971).JBC 246, 7486. Singer, M . F. (1963).JBC 238, 336.

17. POLYNUCLEOTIDE PHOSPHORYLASE

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the type of anhydride bond, and does not phosphorolyze polyribonucleotides. The mechanism by which the yeast enzyme catalyzes the exchange reaction appears to be a displacement of phosphate from NDP by different anions through formation of an intermediate AMP-enzyme complex (133). The kinetic parameters of the PNPase-directed exchange reaction were monitored by the appearance of an isotopic (I8O)shift in 31PNMR profile. Pi ('"04) yielded, during the exchange reaction, an L Y - P ( ' ~ O ~and ~~O a) /3-P(lH04),proving that bond cleavage occurs between the a-P and a-/3 bridge oxygen (134).

C. PHOSPHOROLYSIS In the presence of inorganic orthophosphate, PNPase acts as an exonuclease, releasing NDPs sequentially from the 3'-OH end of the polynucleotide substrate (135). PNPase readily phosphorolyzes single-stranded polynucleotides, but acts more slowly on multistranded structures (136, 137), or on polynucleotides with an extensive secondary structure, such as tRNA, rRNA (83, 137-/39), or mRNA [except the poly(A) tail, which is degraded rapidly; see Ref. 1/40)].The rate of phosphorolysis of RNA chains can be increased by raising the temperature of the reaction mixture (137, 140). The presence of a phosphate group at the 5' end does not prevent phosphorolysis. However, polyribonucleotides with a 3'-terminal phosphate group are not phosphorylyzed by the enzyme (8, 141). Dinucleotides, dinucleoside monophosphates, and, in some cases, trinucleotides are not substrates for phosphorolysis and these compounds accumulate as resistant end products (1 14, 120-122). PNPase phosphorolyzes short oligonucleotides (n 5 12) by a nonprocessive mechanism (82, 85, 112, 115, 142). In contrast, the enzyme tends to phosphorolyze long ( n I 133. Grunberg-Manago,M., Del Campillo-Campbell, A . , Dondon, L., and Michelson, A. M . (1966). BBA 123, 1. 134. Cohn, M . , and Hu, A. (1978). P N A S 75, 200. 135. Hilmoe, R . J . (1959). Ann. N . Y. Arnd. Sci. 81, 660. 136. Ochoa, S. (1957). ABB 69, 119. 137. Grunberg-Manago, M . (1959). JMB 1, 240. 138. Littauer, U. 2.. and Daniel, V. (1962). In "Acides Ribonucleiques et Polyphosphates," Colloq. Intern. du C.N.R.S., Strasbourg, p. 277. C.N.R.S., Paris. 139. Kimhi, Y. (1966). Doctoral Thesis, The Weizmann Institute of Science, Rehovot, Israel. 140. Soreq, H . , Nudel, U., Salomon, R., Revel, M . , and Littauer, U. 2. (1974). I M B 88, 233. 141. Singer, M. F., Heppel, L. A., Hilmoe, R. J., Ochoa, S . , and Mii, S. (1959). Ccm. Cancer Cotif. 3, 41. 142. Kaufmann, G . , Grosfeld, H . , and Littauer, U . 2. (1973). FEES Lerr. 31, 47.

536

U. Z. LITTAUER AND H. SOREQ

20) polynucleotides by a processive mechanism (i.e., the enzyme phosphorolyzes a single chain almost to completion before dissociating to initiate the phosphorolysis of another chain) (82, 83, 9G). The length of substrate at which the transition occurs between the two mechanisms depends on the sequence and the structure of the oligonucleotide (115, 142). Arsenate ions can replace inorganic phosphate in the degradation of polynucleotides by PNPase. The arsenolysis of polynucleotides liberates 5’-phosphorylarsenate nucleotides, which spontaneously hydrolyze to nucleoside monophosphates, arsenate, and H+ ions (143). Aminoacylated . tRNA chains can be phosphorolyzed by PNPase (126, 144). This phosphorolysis occurs by a similar mechanism to that observed with synthetic polynucleotides, as was shown for arsenolysis of valyl-tRNA, which yielded a valyl-adenosine monophosphate product (126). It has been shown that at 37” PNPase phosphorolyzes only part of tRNA molecules present in the reaction mixture, whereas the remaining chains appear to be completely intact (15). To phosphorolyze all the tRNA chains by the enzyme, the temperature of the reaction mixture has to be elevated over 45”, to permit a configurational change of the tRNA (83, 139). Under these conditions, PNPase phosphorolyzes tRNA in a processive mechanism and, similar to the degradation of synthetic polynucleotides, only NDPs and long substrate chains are present in the reaction mixture until the completion of the phosphorolysis reaction (83, 90). The configurational requirements that permit phosphorolysis of tRNA by PNPase are not related to the integrity of the anticodon loop, as tRNAP,h,e,reconstituted from split half molecules still retains the ability to undergo the change in conformation that permits phosphorolysis to occur (145). The transition between the two configurations of tRNA appears to involve large entropic changes, as shown for total unfractionatedE. coli tRNA (84) as well as for purified specific tRNA species (146n). The transition between the two configurational states appears to be initiated at a single “nucleation” center on the tRNA molecule (147, 148). The existence of multiple subsites for the interaction of PNPase with polynucleotides has been indicated from the various K, values that the enzyme displays with synthetic oligonucleotides of different lengths (1 14). This model was further substantiated by the comparative analysis of the 143. Singer, M. F., and O’Brien, B. M. (1963). JBC 238, 328. 144. Yot, P., Gueguen, P:,and Chapeville, F. (1968). FEBS Lett. 1, 156. 145. Beltchev, B., and Thang, M. N. (1970). FEBS Lett. 11, 55. 146. Beltchev, B . , Thang, M. N., and Portier, C. (1971). EJB 19, 194. 146a. Thang, M . N., Buckingham, R. H . , and Dondon, L. (1975). EJB 54, 93. 147. Danchin, A. (1972). FEES Lett. 19, 293. 148. Danchin, A,, and Thang, M . N. (1972). FEBS Lett. 19, 297.

17. POLYNUCLEOTIDE PHOSPHORYLASE

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kinetic parameters for the PNPase-directed phosphorolysis of long polynucleotides, as compared with short oligonucleotide chains (cf. 15). It should be kept in mind that the enzyme molecule appears susceptible to conformational alterations, depending on the substrate present (86). Michaelis constants for short oligonucleotides are higher than those observed for long polynucleotide chains, the transition occurring at 40 2 n 2 10 (85). The detailed equilibrium constants for the phosphorolysis reaction catalyzed by PNPase under physiological conditions have been determined, and the rate of phosphorolysis was found to be sensitive to variations in free Mg2+but relatively insensitive to changes in pH (149).

D. “TRANSNUCLEOTIDATION” PNPase has been shown to catalyze the transfer of nucleoside phosphate moieties from a polynucleotide donor to a polynucleotide acceptor (141). The polynucleotide rearrangement arises from a combination of phosphorolytic and addition reactions of NDPs, catalyzed by trace amounts of inorganic phosphate contaminating the reaction mixture (150). Addition of a phosphate removal system consisting of calf spleen phosphorylase and nicotinamide riboside will block this ‘‘transnucleotidation” reaction. IV.

Attributed Physiological Functions

The apparent ubiquity of PNPase in microorganisms suggests an important role in cell physiology; however, an unequivocal demonstration of its biological function is still lacking. The function of the enzyme has been explored in E. coli mutant cells deficient in PNPase (151-IS), in toluenized cells (62, 156, 157), and in osmotically shocked cells (158). In spite of its widespread occurrence in bacteria, the enzyme is not indispensable to cell metabolism. Escherichia coli mutant cells with defective (152) or very low PNPase activity (155) show no difference in their growth rate at 37”, but grow somewhat more poorly at 45” than their revertants (159). These 149. 150. 151. 152. 153. 154. 155. 156. 157. 158. 159.

Liegel, J . , and Guynn, R. W. (1979). JBC 254, 1992. Sninsky, J . J . , Bennett, G. N., and Gilham, P. T. (1974). Nucleic Acids Res. 1, 1665. Reiner, A. M. (1%9). J B 97, 1431. Reiner, A. M. (1969). J B 97, 1437. Krishna, R. V., Rosen, L., and Apirion, D. (1973). Nature, New B i d . 242, 18. Kinscherf, T. G., Lee, Y.F., and Apirion, D. (1974). Nucleic Acids Res. 1, 1439. Portier, C. (1980). Molec. Gen. Genet. 178, 343. Levin, D. H., Thang, M. N., and Grunberg-Manago, M. (1963). BBA 76, 558. Deutscher, M. P. (1978). JBC 253, 5579. Raue, H. A . , and Cashel, M. (1974). BBA 340, 40. Krishna, R. V., and Apirion, D. (1973). IS 113, 1235.

538

U. Z. LITTAUER AND H. SOREQ

nonlethal mutations affect the structural gene for the a-chains of E. coli PNPase and map close to the argC locus (151, 155). It was suggested that PNPase participates in RNA metabolism (160, 158), and that in contrast to nucleases that liberate nucleoside monophosphates from RNA, PNPase conserves phosphate bond energy by releasing NDPs (160, 161). The liberated NDPs can later be reutilized for RNA synthesis or reduced to dNDP and incorporated after phosphorylation into DNA (160). The participation of PNPase in rRNA and mRNA metabolism has, therefore, been sought. It appears, however, that PNPase is not directly involved in the depolymerization of RNA in E. coli cells (162, 153). Examination of several PNPase deficient E . coli mutants suggests a possible role for PNPase as a salvage enzyme involved in rRNA or mRNA degradation in stressed cells starved for carbon at 49" (69, 163-166). In addition, analysis of PNPase mutants implies that the enzyme may participate in lcic mRNA degradation in heat-shocked cells (167). PNPase may be involved in the inactivation of extraneous eukaryotic mRNA. The expression of the catabolic dehydroquinase gene ( g a - 2 ) from Neurospora C Y Q S S ~is increased as much as 100-fold when cloned in E. cofi strains deficient in PNPase. These results suggest that there are inherent structural differences between prokaryotic and eukaryotic mRNAs (168). It has been suggested that PNPase may have a role in stabilization of mRNA chains by modifying their 3' ends. Comparison of the thermolabile PNPase mutant PR27 to its isogenic counterpart PR100 shows that at 37" or 45" the synthesis of /3-galactosidase proceeds at about the same rate. However, at 49" the functional haif-life of /3-galactosidase is shorter in the PNPase mutant cells (153,159). Several experiments suggest that PNPase could play a role in polyadenylation of mRNA (157, 169, 170). Possibly the poly(A) tail on E . coli mRNA would have a stabilizing function, as has 160. Sekiguchi, M., and Cohen, S. S . (1963).JBC 238, 349. 161. Tumerman, L., and Ric, S . (1977). "Applications of Calorimetry in Life Sciences," p. 97. Walter de Gruyter, Berlin and New York. 162. Chaney, S. G . , and Boyer, P. D. (1972);.J M B 64, 581. 163. Kinscherf, T. G . , and Apirion, D. (1975). Molec. G m . Genet. 139, 357. 164. Kaplan, R., and Apirion, D. (1975). JBC 250, 1854. 165. Kaplan, R., and Apirion, D. (1975). JBC 250, 3174. 166. Cohen, L., and Kaplan, R. (1977). J B 129, 651. 167. Har-El, R., Silberstein, A . , Kuhn, J., and Tal, M. (1979). Molec. Gen. Genet. 173, 135. 168. Hautala, J. A., Bassett, C. L., Giles. N. H., and Kushner, S . R. (1979). P N A S 76, 5774. 169. Wunderli, W., Hutter, R., Staehelin, M., and Wehrli, W. (1975). U B 58, 87. 170. Ramanarayanan, M., and Srinivasan, P. R. (1976).JBC 251, 6274.

17. POLYNUCLEOTIDE PHOSPHORYLASE

539

been suggested for some eukaryotic mRNA species (171,172).In crude extracts of T2L phage-infected E. coli cells, poly(A) synthesis from ATP arises from the combined action of PNPase and ATPase (169). Poyriboadenylate polymerase isolated from E. coii PR7 PNPase mutant will use either ATP or ADP as a substrate, although in this case ATPase appears as an integral part of the enzyme (170).Poly(A) synthesis has also been examined in toluenized E. coli cells. Mutant cells PR7 and PR13, deficient in PNPase, were unable to synthesize poly(A) (157),which is in contrast to the experiments with crude extracts (170).It should be noticed, however, that none of the above mutants are completely devoid of PNPase, as assayed by their phosphorolytic activity (15.2).The possibility that PNPase exists as a multienzyme complex with ATPase is suggested by analysis of the antigenic composition of the plasma membrane of S . pyrogenes (26). V.

Research Applications

A.

POLYNUCLEOTIDE SYNTHESIS

PNPase has been found to be a useful tool for the synthesis of polynucleotides with varied composition, both in the presence and in the absence of primer oligonucleotides. 1. Homopolymers

A large variety of homopolyribonucleotides have been prepared with the aid of PNPase. Because of the tendency of poly(G) to form multistranded helices, the polymerization of GDP proceeds to a very limited degree (3, 5, 106,108). These difficulties may be overcome with PNPase from E. coli by raising the temperature to 60" and by the replacement of Mg" by Mn2+(67,173).Poly(G) can also be synthesized at higher temperatures (70") with the aid of B. stearothe~mopliilus PNPase (17)or with PNPase from Thermirs thrrmophillrrs, in the presence of Mg2+(174).Various preparations of polyinosinic acid synthesized by PNPase differed in their secondary structure andor tertiary conformation. These differences resulted in varied reactivity with anti-poly(1)-antiserum, as well as in dif171. Nudel, U . , Soreq, H . , Littauer, U. Z . , Marbaix, G., Huez, G . , Leclercq, M., Hubert, E . , and Chantrenne, H. (1976). EJE 64, 115. 172. Littauer, U. Z . , and Soreq, H. (1982). Progr. Nirclric Acid Rrs. 27, 53. 173. Thang, M. N . , and Grunberg-Manago, M. (1968). "Methods in Enzymology," Vol. 12B, p. 522. 174. Kikuchi, Y., Hirai, K., Hishinuma, F., and Sakaguchi, K . (1977). BEA 476, 287.

540

U . Z. LITTAUER AND H. SOREQ

ferent abilities to induce the production of interferon in virus-infected cells (175). The basis for the differences with anti-poly(1) antiserum could be due to variability in the amount of hypoxanthine that is accessible to the antibody (176). PNPase also polymerizes modified NDPs, although at relatively slow rates. The range of NDP modification can be further extended by the use of Mn2+ as a cofactor, or with a matrix-bound enzyme (177). Thus, PNPase catalyzes the synthesis of polypseudouridylic acid (178-18O), poly-Zthiocytidylic acid (181, 182), poly-Cthiouridylic acid (183, l M ) , polyribothymidylic acid (185), poly-5-methyluridylic acid (186 ), poly-5ethyluridylic acid (187), polyfluorouridylic acid (188), poly-8-chloroadenylic acid (189), poly-8-oxyadenylic acid (I90), polyfluoroadenylic acid (191), poly(0 6-methyl or ethy1)guanylic acid, (192), and many other homopolymers. Fluorescent nucleotide analogues (lin-benzo-ADP and lin-IDP) have been prepared for use as dimensional probes of PNPase binding sites (193, 194). In contrast, some modifications render the modified nucleotide unsuitable for polymerization by PNPase. These modified NDPs, such as 5-acetyluracl NDP, may act as weak inhibitors of the enzyme (195). Chemical modifications of specific groups in NDPs were an aid in asses175. Stollar, B. D., DeClercq, E., Drocourt, J.-L., and Thang, M. N. (1978). HE 82, 339. 176. Inouye, H., Fuchs, S. , Sela, M., and Littauer, U. Z. (1971). BEA 240, 594. 177. Brentnall, H. J., and Hutchinson, D. W. (1972). Tetrahedron Lett. 25, 2595. 178. Sasse, L., Rabinowitz, M., and Goldberg, I. H. (1963). BBA 72, 353. 179. Pochon, F., Michelson, A. M., Grunberg-Manago, M., Cohn, W. E., and Dondon, L. (1964). BBA 80, 441. 180. Goldberg, I. H. (1968). “Methods in Enzymology,” Vol. 12B, p. 519. 181. Scheit, K. H., and Faerber, P. (1971). EJE 24,385. 182. Faerber, P., Scheit, K. H., and Sommer, H. (1972). EJE 27, 109. 183. Eckstein, F., and Scheit, K. H. (1971). I n “Procedures in Nucleic Acid Research” (G. L. Cantoni and D. B. Davies, eds.), Vol. 2, p. 665. Harper and Row, New York. 184. Fiser, I., Scheit, K. H., and Kuechler, E. (1977). EJB 74, 447. 185. Griffin, B. E., Todd, A., and Rich, A. (1958). PNAS 44, 1123. 186. Grunberg-Manago, M., and Michelson, A. M. (1964). EBA 80, 431. 187. Biala, E., Jones, A. S., and Walker, R. T. (1980). Tetrahedron 36, 155. 188. Grunberg-Manago, M., and Michelson, A. M. (1964). EBA 87, 593. 189. Tavale, S. S., and Sobell, H. M. (1970). JMB 48, 109. 190. Folayan, J. O., and Hutchinson, D. W. (1977). BEA 474, 329. 191. Broom, A. D., Amamath, V., Vince, R., and Brownell, J. (1979). EBA 563, 508. 192. Mehta, J. R., and Ludlum, D. B. (1976). Biochemistry 15, 4329. 193. Leonard, N. J., Scopes, D. I. C., VanDerLijn, P., and Barrio, J. R. (1978). Biochemistry 17, 3677. 194. Leonard, N. J., and Keyser, G. E. (1979). PNAS 76, 4262. 195. Jones. A. S., Stephenson, G. P., and Walker, R. T. (1979). Tetrahedron 35, 1125.

17. POLYN UCLEOTIDE PHOSPHORYLASE

54 1

sing the influence of these groups on the physical and chemical properties of polynucleotides (196). Thus, the role of the 2’-hydroxyl group in RNA conformation has been studied with the aid of 2‘-modified polynucleotides (197-200). Polymerized uridine-5‘-diphosphorothioateexhibits a certain extent of protection against nucleolytic degradation (20f). Poly-S4methyl-4-thiouridylic acid displays a specific emission spectra at 520 nm (202). 2-Azaadenosine and 2-azainosine diphosphates (203), as well as 2-methyl- and 2-ethylthioadenosine diphosphates (204) can be polymerized to their respective homopolymers, and the modifications do not prevent the formation of double-stranded complexes by the modified polymers. Poly(2’-deoxy-2’-fluorodenylicacid) and poly(2’-chloro-2’deoxyinosinic acid) have rather similar properties to those of poly(A), but differ from poly(dA) (205, 206). In addition, poly-5-methoxyuridylic acid stimulated the binding of Phe-tRNA to 70 S ribosomes, although it was inactive in directing poly(Phe) synthesis (207). In contrast, poly(Zfluoroadeny1ic acid) codes for the synthesis of polylysine (191). 2. Heteropolymers Polymerization of a mixture of NDPs that contain different bases results in random copolymers. Purine NDPs in which the purine and ribose are in the syn-conformation (189) are poor substrates for PNPase. However, as is the case for GDP, they may be incorporated into copolymers to a varying degree with the normal NDPs. Thus, ribopolynucleotides that contain 8-substituted purine nucleotides, such as 8-bromoadenosine, 8-oxyadenosine, 8-bromoguanosine, 8-oxyguanosine, and 8-dimethylaminoguanosine (208), as well as 1-methyl-6-thioguanosine(209) are synthesized by copolymerization of the modified NDPs with ADP or 196. Michelson, A . M., Massoulie, J . , and Guschlbauer, W. (1967). Progr. Nucleic Acid Res. 6, 83.

197. Szer, W., and Shugar, D. (1966). J M B 17, 174. 198. Zmudzka, B . , Janion, C., and Shugar, D. (1969). BBRC 37, 895. 199. Zmudzka, B . , and Shugar, D. (1970). FEBS L e f t . 8, 52. 200. Torrence, P. F., Bobst, A . M . , Waters, J . A , , and Witkop, B. (1973). Biochemistry 11, 3962. 201. Eckstein, F., and Gindl, H. (1969). FEBS Lett. 2, 262. 202. Scheit, K . H . (1970). BBA 209, 445. 203. Fukui, T., Kakiuchi, N . , and Ikehara, M. (1978). BBA 520, 441. 204. Fukui, T., and Ikehara, M. (1979). BBA 562, 527. 205. Ikehara, M . . Fukui, T., and Kakiuchi, N . (1978). Nucleic Acids Res. 5, 1877. 206. Kakiuchi, N . , Fukui, T., and Ikehara, M. (1979). Nucleic Acids Res. 6, 2627. 207. Hillen, W., and Gassen, H. G. (1979). BBA 562, 207. 208. Ikehara, M . , Tazawa, I., and Fukui, T. (1969). Biochemistry 8, 736. 209. Amarnath, V., and Broom, A. D. (1977). BBA 479, 16.

542

U. Z. LITTAUER AND H. SOREQ

GDP. Copolymers that contain other base analogs, such as xanthosine, N’-methyluridine, N‘-acetylcytidine, and many others, have also been prepared, and serve to examine the role of rare and of “nonsense” bases in directing in virro protein synthesis (210, 21 I). 5’-Mercaptouridine 5’-diphosphate has been copolymerized with UDP and the resulting copolymer, after formation of double-stranded complex with poly(A), served as a potent inhibitor for DNA-dependent RNA polymerase (RNAdependent DNA nucleotidyltransferase, EC 2.7.7.7) (212). In contrast, polynucleotides that contain Oz- and 04-alkyluridine (213) or 2-thiocytidine (214) serve as templates for RNA polymerase activity. Heteropolymers that contain 2‘-O-methyladenylic acid and 2’-O-methylcytidylic acid have also been prepared (215, 2161, and dihydrouridine was more efficiently incorporated into heteroribopolymers than l-(fl-D-ribofuranosy1)-(a + P)5,6-methyleneuracyI (217). The stereochemistry of PNPasedirected internucleotide bond formation has been probed by polymerization of the exoisomer of adenosine Y-04 1-thiodiphosphate), which undergoes inversion of its configuration into the endoisomer when copolymerized with UDP by PNPase (124). The fluorescent analog of adenosine, 1-N6-etheno-2-azaadenosine,has been incorporated into heteropolymers with ADP, UDP, or IDP and provides means for probing the structure of these polymers (218). The effect of spin-labeled copolymers on the reaction catalyzed by avian myoblastosis virus RNA-dependent DNA polymerase was studied by PNPase-directed copolymerization of 4-thiouridine and uridine, and it was shown that increasing amounts of potentially reactive thiol groups (or spin labels) enhance the inhibitory properties of the copolymers as compared to poly(U) (219). Simplified methods for the large-scale preparation of homooligonucleotides (220) and of heterooligonucleotides that contain modified nucleosides (221) have also been reported. A description of the various approaches utilized to 210. Michelson, A. M., and Grunberg-Manago, M. (1964). BBA 91, 92. 211. Michelson, A. M., and Pochon, F. (1966). BBA 114, 469. 212. Ho,Y.-K., Aradi, J., and Bardos, T. J . (1980). Nitcleic Acids Res. 8, 3175. 213. Singer, B . , Fraenkel-Conrat, H., and Kusmierek, J. T. (1978). PNAS 75, 1722. 214. Kroger, M . , and Singer, B. (1979). Biochemistry 18, 91. 215. Rottman, F., and Johnson, K. L. (1969). Biochemistry 8, 4354. 216. Simuth, J . , Strehlke, P., Niedballa, U., Vorbruggen, H., and Scheit, K. H. (1971). BBA 228, 654. 217. Torrence, P. F., and Witkop, B. (1972). Biochemistry 11, 1737. 218. Yip, K. F., and Tsou, K. C. (1979). Biopo/ymers 18, 1389. 219. Warwick, P.E., Hakam, A., Bobst, E. V., and Bobst, A. M. (1980). PNAS 77,4574. 220. Shum, B. W.-K., and Crothers, D. M. (1978). Nucleic Acids Res. 5, 2297. 221. Schetters, H., Gassen, H. G . , and Matthaei, H. (1972). BBA 272, 549.

17. POLYNUCLEOTIDE PHOSPHORYLASE

543

synthesize various building blocks for polynucleotide synthesis has been reviewed (222).

B . SYNTHESIS OF OLIGONUCLEOTIDES WITH A DEFINED SEQUENCE Under high salt concentrations, PNPase adds only a few nucleotide residues to the 3‘ end of a dinucleotide primer. This property of the enzyme served for the first preparations of oligonucleotides of defined sequence (223). The addition of one or two guanyl residues to oligonucleotide primers is achieved by incubation with PNPase from Thermu.~ thermophilrrs at 37”. At this relatively low temperature, poly(G) formation is inhibited (224). Monovalent addition of GMP residue to guanosine-free oligonucleotides, obtained by T1 ribonuclease digestion of RNA, can also be carried out by the simultaneous action of PNPase and T1 ribonuclease (224-226). Similarly, copolymers with a terminal pyrimidine residue are obtained by polymerization of a mixture of purine and pyrimidine NDPs with PNPase in the presence of pancreatic RNase (227). Two functional regions can be defined in the NDP monomers that serve as substrates for the polymerization reaction catalyzed by PNPase: The P-phosphate residue, which is eventually released as inorganic phosphate, and the free 3‘-hydroxyl group of the incoming NDP, which becomes the new accepting terminus (126). Certain modifications of the sugar moiety of the NDP substrate may convert it to a “monofunctional” substrate for PNPase. Such NDP derivatives, blocked in their 3’-hydroxyl function (probably due to steric hindrance), do not sustain de novo polymerization but are able to transfer one nucleotidyl residue to an oligonucleotide initiator, thus serving as chain terminators. The blocking group can be subsequently chemically removed from the oligonucleotide product, permitting a succession of single addition reactions to be carried out. This procedure has been utilized for the stepwise synthesis of polyribonucleotides of defined sequence (126, 228). 222. Seliger, H . , Haas, B . , Holupirek, M., Knaeble, T.,Todling, G . , and Philipp, M . (1980). N d r i c . A t k f s l i c ? . ~ .. Syrnp. S r r . N O . 7. 191 . 223. Thach, R. E. (1966). Zn “Procedures in Nucleic Acid Research” (G. L. Cantoni and D. R. Davies, eds.), p. 520. Harper and Row, New York. 224. Kikuchi, Y . , Hirai, K . , and Sakaguchi, K. (1979). J B 86, 1427. 225. Szeto, K. S . , and Soll, D. (1974). Nucleic Acids Res. 1, 171. 226. Kikuchi, Y., and Sakaguchi, K. (1978). Nircleic-Acids Res. 5, 591. 227. Saunders, C . A., Sogin, S. J . , and Halvorson, H. 0. (1979). A B 95, 171. 228. Kaufmann, G . , Zutra, A., and Littauer, U. Z. (1971). Isr. J . Chem. 9, 44BC.

544

U. Z. LITTAUER AND H. SOREQ

NDPs containing a variety of blocking groups have been employed for the monovalent addition of a single nucleoside residue to a given oligonucleotide primer (229).These include the corresponding 2’(3’)-O-isovaleryl (127, 230), and 2’(3‘)-O-a-methoxyethyl(128, 150, 231 -234) diphosphates that were added to trinucleotide primers with a free 3’-OH group. After removal of the protecting groups by treatment with weak alkali (isovaleryl) or acid (methoxyethyl), the products can serve as acceptors for a second single-addition reaction. Oligonucleotides of defined sequence of four to seven residues have been synthesized by these methods (127, 228, 231). NDPs that contain 2‘(3’)-dihydrocinnamoyl (235) and the photolabile 2’-0-(0 -nitrobenzyl) groups have been utilized for the monoaddition to tri- and tetranucleotide primers (236, 237). The monoaddition reaction is accompanied by a limited rearrangement of the initiator oligonucleotide ( l 2 7 ) , which can be circumvented by coupling the reaction with an enzyme system that utilizes inorganic phosphate either present or formed in the reaction mixture (150, 238). Combination of these and other reactions, such as the use of T4 RNA ligase to ligate the synthesized oligonucleotides, permits the synthesis of oligoribonucleotides of defined sequence of appreciable length (226,239,240).NDPs in which the C-2’-C-3’ bond has been cleaved (ox-red nucleosides) by periodate oxidation followed by borohydride reduction may serve as monovalent terminators of PNPase-catalyzed polymerization, and can also be used for radioactive labeling of the 3‘ termini of polyribonucleotides (232). 229. Kossel, H., and Seliger, H. (1975). In “Recent Advances in Polynucleotide Synthesis” (W. Herz, H. Grisebach, and G. W. Kirby, eds.), p. 467. Springer-Verlag, Berlin and New York. 230. Walker, G. C., and Uhlenbeck, 0. C. (1975). Biorhernistry 14, 817. 231. Mackey, J . K., and Gilham, P. T. (1971). Natrire (London) 233, 551. 232. Hawley, D. M., Sninsky, J. J., Bennett, G. N., and Gilham, P. T. (1978). Biochemisrp 17, 2082. 233. The Nucleic Acid Synthesis Group, Shanghai (1979). Aria Biorhirn. Biophys. Sin. 11, 290. 234. Sninsky, J. J., Hawley, D. M., and Bennett, G. N. (1975). FP 34, 702. 235. Kikuchi, Y., Hirai, K., and Sakaguchi, K. (1975). J B 77, 469. 236. Ikehara, M., Tanaka, S., Fukui, T., and Ohtsuka, E. (1976). Nucleir Acids Res. 3, 3203. 237. Ohtsuka, E., Tanaka, S., Hayashi, M., and Ikehara, M. (1979). BBA 565, 192. 238. Kikuchi, Y., Someno, K., and Sakaguchi, K. (1977). Agr. Biol. Chem. 41, 1531. 239. Kaufmann, G . , and Littauer, U. Z. (1974). PNAS 71, 3741. 240. Gumport, R. I. and Uhlenbeck, 0. C. In “Gene Amplification and Analysis” (J. G. Chirikjian and T. S. Papas, eds.), Vol. 11, in press. Elsevier North Holland, New York.

17. POLYNUCLEOTIDE PHOSPHORYLASE

c.

POLYMERIZATION

OF

545

DEOXYRIBONUCLEOTIDES

PNPase is unable to phosphoroloyze DNA (8). However, the enzyme can direct the reversible addition of a single deoxynucleotidyl residue to ribooligonucleotide primers. Further addition of deoxynucleotide residues to the resulting product is very difficult (99, 113, 131, 241, 242). PNPase does not readily catalyze the de nuvo synthesis of (dA), chains, probably because it is a poor substrate for chain initiation (113, 131,243). However, in the presence of Mn2+, E. culi PNPase catalyzes the transfer of deoxyribonucleotide residues from dNDPs to the 3’ OH end of an oligodeoxyribonucleotide primer having a minimal length of three nucleoside residues. This allows the synthesis, by repeated addition of single residues, of oligodeoxyribonucleotidesof defined sequence, although the overall yield is rather poor. The kinetics of the addition reactions differ for various deoxyribonucleoside 5’-diphosphates and for different primers (244-248). The limited addition reaction displayed with deoxyribonucleoside diphosphates contrasts with the extended polymerization that has been observed for a number of dNDP derivatives that contain substituents at the C-2’ position (198-200, 217, 242, 243, 249-254). PNPase also adds to oligodeoxynucleotide primers modified deoxynucleoside diphosphates, such as 5-methyldeoxycytidine, N4-hydroxydeoxycytidine, and deoxyuridine. Some modifications, such as 5-mercurideoxyuridine, prevent the addition of the modified nucleoside base to deoxyribooligonucleotide primer by PNPase (255).

241. Feix, G . (1972). BBRC 46, 2141. 242. Batey, I . L., and Gilham, P. T. (1974). Biochemistry 13, 5395. 243. Rottman, F., and Heinlein, K. (1968). Biochemistry 7, 2634. 244. Gillam, S., Rottman, F., Jahnke, P., and Smith, M . (1977). P N A S 74, 96. 245. Gillam, S . , Jahnke, P., and Smith, M. (1978). JBC 253, 2532. 246. Gillam, S . , and Smith, M . (1980). “Methods in Enzymology,” Vol. 65, p. 687. 247. Wu, R., Bahl, C. P., and Narang, S. A. (1978). Progr. Nucleic Acid Res. 21, 101. 248. Trip, E. M., and Smith, M. (1978). Nucleic Acids R e s . 5, 1529. 249. Janik, B . , Kotick, M. P., Kreiser, T. H., Reverman, L. F., Sommer, R. G . , and Wilson, D. P. (1972). BBRC 46, 1153. 250. Hobbs, J., Sternbach, H . , and Eckstein, F. (1971). FEBS Lett. 15, 345. 251. Hobbs, J . , Sternbach, H . , and Eckstein, F. (1972). BBRC 46, 1509. 252. Hobbs, J . , Sternbach, H . , Sprinzl, M . , and Eckstein, F. (1972). Biochemistry 11, 4336. 253. Khurshid, M., Khan, A , , and Rottman, F. M. (1972). FEBS Lett. 28, 25. 254. Tazawa, I . , Tazawa, S . , Alderfer, J. L., and Ts’o, P. 0. P. (1972). Biochemistry 11, 493 1. 255. Trip, E. M . , and Smith, M. (1978). Nircleic Acids R e s . 5, 1539.

546 D.

U . Z. LITTAUER AND H. SOREQ

CONJUGATION T O INSOLUBLE MATRIX

PNPase from both E. coli (42,117,256-259)and M . Iuteus (256)has been bound to a variety of insoluble matrices, such as cellulose nitrate filters (260),cellulose beads (261),mercerized cellulose (256),Sepharose 4B (42, 47, 228, 256), hydrazide agarose (257), diazotized p-aminobenzenesulfonylethyl (ABSE) agarose, ABSE-Sephadex G-200, and ABSE-cellulose (258, 259). Cellulose-bound PNPase can polymerize NDPs under pH conditions at which phosphorolysis is negligible (256). The insoluble PNPase has therefore been used to improve the yield of the polymerization reactions, especially those that involve atypical bases and are difficult to carry out, such as poly 8-chloroadenylic acid (177) and poly(1) chains (257). Cellulose- and Sepharose-bound PNPase phosphorolyze polynucleotide chains at a slower rate than that of the soluble enzyme (42, 228, 260), and display K , values for long polynucleotides that are higher by two orders of magnitude than those measured for the soluble enzyme (117). The phosphorolysis of long RNA molecules by Sepharose-bound PNPase involves three active subunits, as has been titrated by the removal of poly(A) tails from globin mRNA (42). Unlike the soluble enzyme, bound PNPase phosphorolyzes polynucleotides by a nonprocessive mechanism, although the kinetic parameters of the phosphorlysis of short oligonucleotides are unaltered ( 1 17). The insoluble PNPase has several advantages over the soluble enzyme, both for analytical and preparative purposes. The same enzyme preparation can be recycled multiple times (42,258,259),and the separation of the reaction products from the enzyme is greatly simplified. Thus, insoluble PNPase has been used for the enzymatic synthesis of polynucleotides ( 1 17, 258, 259) as well as for controlled phosphorolysis of mRNA (42), of viral RNAs (262, 2631, and of whole TMV viral particles, in which the 256. Smith, J . C . , Stratford, I. J . , Hutchinson, D. W., and Brentnall, H. J. (1973). FEBS Lett. 30, 246.

257. Bachner, L., De Clercq, E., and Thang, M. N . (1975). BBRC 63, 476. Liu, N.-J., and Lin, Y. (1979). Acttr Biochim. Biophys. Sin. 11, 87. 258. Yang, K.-Y., 259. Yang, K.-Y., Liu, N.-J., and Lin, Y. (1979). Acra Biochim. Biophys. Sin. 11, 104. 260. Thang, M. N . , Graffe, M., and Grunberg-Manago, M. (1968). BBRC 31, 1 . 261. Hoffman, C. H . , Harris, E., Chodroff, S., Michelson, S., Rothrock, J. W., Peterson, E., and Reuter, W. (1970). BBRC 41, 710. 262. Salomon, R . , Sela, I . , Soreq, H., Giveon, D., and Littauer, U. Z. (1976). Virology 71,74. 263. Salomon, R . , Bar-Joseph, M., Soreq, H . , Gozes, I., and Littauer, U. Z. (1978). Virology 90, 288.

17. POLYNUCLEOTIDE PHOSPHORYLASE

547

3‘-terminal nucleotides are vulnerable to the nucleolytic attack by PNPase even in the presence of the viral protein coat (264). E.

RADIOLABELED NUCLEOTIDES AND FINGERPRINTING OF OLIGONUCLEOTIDES S Y N T H E S I S OF

PNPase has been used to synthesize radiolabeled polyribonucleotides from NDP monomers (265,266). It has also been used for sequence analysis of short oligoribonucleotides. These are phosphorolyzed by PNPase starting from the 3’ end in a stepwise fashion, and by a nonprocessive mechanism (82, 85, 112, 1/.5), to yield a mixture of NDPs and a limit oligonucleotide that cannot further be degraded by the enzyme. One may use labeled oligonucleotides or include [32P]orthophosphatein the reaction mixture. By following the order in which the released P-labeled NDPs appear during the phosphorolysis of a given oligonucleotide, it is possible to determine the nucleotide sequence from the 3‘ end up to 2-3 residues from the 5’ terminus (142). This scheme served to develop a method for sequence analysis of short oligonucleotides. PNPase-directed labeling of nucleolytic cleavage-oligonucleotides, aids in the fingerprint analysis of RNA sequences. RNA fragments derived by T1 RNase are dephosphorylated with bacterial alkaline phosphatase to yield oligonucleotides with free 3‘-hydroxyl groups, which may in turn serve as primers for polymerization by PNPase. In the presence of as a substrate and T1 ribonuclease, only a single a-labeled [CX-~~PIGDP GMP is added to the pancreatic RNase-derived fragments (225). Several procedures that use PNPase have been developed for the labeling of NDPs and NTPs at their @-position. One of these utilizes the exchange reaction catalyzed by PNPase between [32P]inorganicphosphate and the @-phosphatemoiety of a given NDP. The [@-32P]NDP obtained can then be phosphorylated to generate the [P-32P]NTPderivative (267-270). Another technique that yields [@-32P]NTP with a very high specific activity, exploits the phosphorolysis properties of PNPase. According to this 264. Littauer, U . Z . , Soreq, H., and Cornelis, P. (1980). In “Enzyme Regulation and Mechanism of Action” (P. Mildner and B. Ries, eds.), FEBS, Vol. 60,p. 233. Pergamon, New York. 265. Leder, P., Singer, M. F., and Brimacombe, R. L. C. (1%5). Biochemistry 4, 1561. 266. Singer, M. F., Hilrnoe, R . J . , and Heppel, L. A. (1960). JBC 235, 751. 267. Littauer, U. Z . , Kimhi, Y., and Avron, M. (1964). AB 9, 85. 268. Gilboa, E., Soreq, H . , and Aviv, H. (1977). EJB 77, 393. 269. Vennstrom, B . , Pettersson, U . , and Philipson, L. (1978). Nircleic Acids Res. 5, 205. 270. Eliasson, R., and Reichard, P. (1978). JBC 253, 7469.

548

U. 2.LITTAUER AND H. SOREQ

procedure, a polyribonucleotide of choice is phosphorolyzed by PNPase in the presence of carrier-free 32P-inorganicphosphate. The resulting [p32P]NDPproduct is then phosphorylated by pyruvate kinase, which drives the reaction to completion (271, 272). [p-32P]Purinetriphosphates prepared by this method serve as useful precursors in studying the initiation of eukaryotic mRNA (268) and their 5'-terminal caps (273), as well as in studying the initiator RNA of short nascent DNA chains (Okazaki pieces) (270).

F. SYNCHRONOUS PHOSPHOROLYSIS AS A N ANALYTICAL TOOL 1. Removal of Poly(A) Tracts from mRNA

The 3'-exonucleolytic activity of PNPase has been used for &heanalysis of the size and composition of the 3'-terminal sequence of RNA molecules (140, 171, 172, 264, 274-278). The analysis is based on the property of the enzyme to phosphorolyze long polynucleotides by a processive mechanism. The use of molar excess of PNPase over the substrate establishes a synchronous mode of phosphorolysis, in which NDP molecules are sequentially released from the 3' terminus of the RNA chains. In order to follow the course of phosphorolysis, [32Plorthophosphateis included in the reaction mixture and the released &labeled NDPs are analyzed by DEAE-cellulose paper chromatography (140, 142) or by PEI-cellulose ascending thin-layer chromatography (27.5, 279). The size of the shortened RNA molecules is then determined by gel electrophoresis on polyacrylamide-agarose composite gels (140, 263) or in gels under denaturing conditions (276, 278). In some cases 32P-labeledor 13H]uridinelabeled RNA was included in the phosphorolysis reaction and the released NDPs are labeled accordingly (276, 278, 280). 271. Leung, K.-L., and Yamazaki, H. (1977). Can. J . Eiochem. 55,223. 272. Kaufmann, G., Choder, M . , and Groner, Y. (1980). AE 109, 198. 273. Groner, Y., Gilboa, E., and Aviv, H. (1978). Biochemistry 17, 977. 274. Littauer, U. Z., Salomon, R., Soreq, H., Fleischer, G . , and Sela, I. (1975). I n "Organization and Expression of the Viral Genome. Molecular Interaction in Genetic Translation" (F.Chapeville and M. GrunbergManago, eds.), Vol. 39, p. 133. Roc. 10th FEBS Meeting, Paris. 275. Vournakis, J. N., Efstratiadis, A., and Kafatos, F. C. (1975). PNAS 72, 2959. 276. Grosfeld, H . , Soreq, H . , and Littauer, U. Z. (1977). Nucleic Acids Res. 4, 2109. 277. Kaempfer, R . , Hollender, R., Soreq, H., and Nudel, U. (1979). EJB 94, 591. 278. Soreq, H., Sagar, A. D., and Sehgal, P. B. (1981). PNAS 78, 1741. 279. Deshpande, A. K., Chatterjee, B., and Roy, A. K . (1979). JEC 254, 8937. 280. Sehgal, P. B., Soreq, H., and Tamm, I. (1978). PNAS 75, 5030.

17. POLYNUCLEOTIDE PHOSPHORYLASE

549

At 0" the poly(A) tails of mRNA molecules are readily phosphorolyzed while the rest of the RNA chains remain intact (140, 171). The rate of poly(A) phosphorolysis varies with the ionic strength, and ranges between 7.5 nucleotides per chain per minute at a 1.0 M NaCl concentration and 75.0 nucleotides per chain per minute at an ionic strength of 0.15 M (140, 264). The calculated rate of phosphorolysis is based on the assumption that all RNA chains are bound to enzyme molecules and phosphorolyzed synchronously at the same rate. Analysis of a heterogeneous RNA population therefore yields an average rate measurement. This implies that the measured rate of phosphorolysis may be underestimated in cases where not all the chains possess poly(A) tails (264). The length of the phosphorolyzed poly(A) tail has been estimated by comparative gel electrophoresis of native and deadenylated mRNA and by determination of the number of moles of ADP that are liberated per mole of RNA (140, 276). Using these methods, it has been established that the average size of the poly(A) tail for different preparations of rabbit globin mRNA is between 120-150 residues (140, 277). Similar experiments with the rat liver mRNA for a-2p-globulin revealed a variety of lengths for its poly(A) tails, ranging between 40 to 175 residues (279). 2. Phosphorolysis qf 3 ' Seyirences from R N A As mentioned above, at 0" only poly(A) tails are phosphorolyzed, possibly due to the difference in secondary structure between homopolynucleotides and heteropolynucleotides. Raising the temperature of incubation to 37" allows the phosphorolysis of heterogeneous sequences from RNA populations. Even at this elevated temperature and at low salt concentration, the rate of phosphorolysis varies greatly among the RNA species tested. Thus, after the removal of the poly(A) region from globin mRNA at O", the rest of the RNA chains are phosphorolyzed synchronously at 37" at an average rate of 9 nucleotides per chain per minute (227). A similar rate of phosphorolysis is observed with the larger (1.4 X lo6MW) RNA from Carnation mottle virus (263). TMV RNA, in contrast, is phosphorolyzed at a much slower rate of 3.5 nucleotides per chain per minute (274). This may result from the more compact conformation of the tRNAlike structure at the 3' end of the TMV RNA, that is specifically aminoacylated with histidine (262). In vivo protection of RNA sequences at their 3' end may also be indicated from the fact that rho protein, an RNA synthesis termination factor fromE. coli, binds tightly to poly(C) or poly(U) and prevents their degradation by PNPase (281). 281. Galluppi, G . R.,and Richardson, J . P. (1980). J M B 138, 513.

550

G.

U. Z. LITTAUER A N D H. SOREQ PROBE FOR T H E REGULATORY FUNCTION OF T H E 3'-OH REGIONOF RNA

1. Regulatory Role of Poly(A) fvom Vurious mRNA Species

Synchronous phosphorolysis of RNA by PNPase has been used to examine the role of the 3' nontranslatable regions of RNA chains (cf. 172). Highly purifiedE. coli polynucleotide phosphorylase (42)was used to phosphorolyze the poly(A) tracts of rabbit globin mRNA under conditions in which poly(A) is removed but the rest of the molecule remains intact (140, 264). The deadenylated globin mRNA is translated in vitro as efficiently as native mRNA for a short while. Upon longer periods of incubation, the rate of protein synthesis decreased more rapidly with the deadenylated mRNA than with the native mRNA (140), suggesting that the presence of the poly(A) sequence may stabilize the functional activity of mRNA molecules in vitro. This stabilization is not limited to mRNA molecules. Addition of poly(A) segments to E. coli 5 S RNA, which is also carried out with the aid of PNPase, increased the stability of the 5 S RNA against endonucleolytic attacks (282). However, the interaction between globin mRNA and the initiation factor that binds methionyl tRNAfMe'remained unchanged when phosphorolysis was used to remove the poly(A) tail or even the 90 nucleotides adjacent to it in the untranslated 3' sequence (277). In contrast, deadenylation of ovalbumin mRNA by PNPase was reported to reduce the initiation process (283). It is not clear, however, whether o r not other regions of the mRNA were altered as well during the deadenylation procedure, reported for ovalbumin mRNA. The removal of the poly(A) region clearly decreases the functional and physical stability of globin mRNA in microinjected Xenopus oocytes. Following an equilibration period, the native poly(A)-containing globin mRNA remains fully active for at least 72 hr (284), and so does globin mRNA species from which their poly(A) tails are shortened down to 32 adenylate residues (171). Poly(A)-free globin mRNA and a globin mRNA population that contains an average length of 16 adenylate residues per chain showed a rate of decay with a of about 6 hr. Thus, the poly(A) region must contain a minimal number of about 30 adenylate residues to ensure its protective function ( 1 71). The stabilizing role of poly(A) on mRNA is not a general phenomenon (172). Thus, the physical and functional stability of human interferon 282. 283. 284. trenne,

Hieter, P. A . , LeGendre, S. M . , and Levy, C. C. (1976). JBC 251, 3287. Doel, M . T., and Carey, N . H. (1976). Cell 8, 51. Marbaix, G., Huez, G., Burny, A., Cleuter, Y., Hubert, E . , Leclercq, M., ChanH., Sores, H., Nudel, U . , and Littauer, U. Z. (1975). P N A S 72, 3065.

17. POLYNUCLEOTlDE PHOSPHORYLASE

55 1

mRNA species in microinjected Xenoprrs oocytes is not affected by the removal of poly(A) tails from their 3' termini with PNPase (278, 280). Since poly(A) tails exist on most species of mRNA, it appears that the biological role of poly(A) other than as a stabilizing element remains to be revealed.

2. Role (d3' Termini in tRNA and in rRNA Replacement of the 3'-terminal adenosine moiety of tRNA with 2'- and 3'-deoxyadenosine afforded tRNA species useful in defining the nature of the partial reactions which comprise protein biosynthesis. Thus, incubation of an enzymatically abbreviated tRNA (tRNA-C-COH) with 2'deoxy-3'-O-~-phenylalanyladenosine and PNPase yielded tRNA terminating with the corresponding aminoacylated deoxynucleoside. The yield of this product is increased by including 20% methanol in the reaction mixture (285). Processive removal of 160 nucleotides from the 3' end of E. coli 16 S rRNA was found to have little if any effect on the ability of the phosphorolyzed rRNA to be reconstituted into 30 S ribosomal subunits, which contain all of the native ribosomal proteins and bind formylmethionyltRNA with equal efficiency t o native 30 S subunits, but have low capacity to direct protein synthesis (286). 3. Role of 3' Sequences in Viral R N A PNPase has been extensively employed to reveal the role of 3' sequences in numerous polyadenylated as well as poly(A)-deficient viral RNA species. The poly(A) tail of mRNA from Sendai virus (287) and from measles virus (288) have been shown to be nonexposed to exonucleolytic attack by PNPase. Deadenylation of poliovirus RNA abolishes its infectivity, as a result of the inability of the RNA to serve as a template for the viral replicating enzyme (289). Several viral RNA and viral mRNA species contain mixed nucleotide sequences at the 3' end, rather than a poly(A) tail. The 3' end of TMV RNA, which is devoid of a poly(A) tail, is essential for its infectivity. It has been observed that synchronous phosphorolysis of about 5 nucleoside residues per chain completely abolished the infectivity of the phosphorolyzed TMV RNA (262). Even within the viral particles, the same 285. Chinault. A. C . , Kozarich, J. W., Hecht, S. M . , Schmidt, F. J . , and Bock, R. M. (1977). Biochernb/ry 16, 756. 286. Zagorska, L., Szkopinska, A., Klita, S., and Szafranski, P. (1980). BBRC 95, 1152. 287. Marx, P. A., Jr., F'ridgen, C., and Kingsbury, D. W. (1975). J . Gen. Virol. 27, 247. 288. Hall, T. C. (1979). fntern. R e v . Cytol. 60, 1. 289. Dasgupta, A , , Zabel, P., and Baltimore, D. (1980). Cell 19, 423.

552

U. Z. LITTAUER AND H . SOREQ

3'-terminal nucleotides of TMV RNA appear to be vulnerable to exonucleolytic attack, and their removal by immobilized PNPase destroys their infectivity (264 ). A plant virus of a different architectural design is the Carnation mottle virus (CarMV), consisting of round particles with no vulnerable termini. When translated in cell-free extracts, CarMV RNA operates as a polycystronic message, which induces the synthesis of three distinct polypeptides of molecular weights of 77,000, 38,000, and 30,000. The 38,000 polypeptide is the subunit of the viral coat protein. In contradistinction with TMV RNA, the infectivity and the translational activity of CarMV RNA chains is gradually reduced following the removal of 3'-end sequences with the aid of PNPase. The rate of decrease of the infectivity is faster than the ability to sustain in v i m translation of the viral coat protein. Moreover, the reduction in the rate of synthesis of the 77,000 product is even faster than loss of infectivity. These observations imply that this unidentified large polypeptide, but not the viral coat protein, may be essential for infection and that the translation of CarMV RNA into this protein is highly dependent upon the intactness of the vulnerable 3' end (263). 4. Role of 3'-Noncoding Sequences in mRNA

The coding regions in all known cases of mRNAs, whether polyadenylated or not, are followed by 3'-nontranslated heteropolymeric sequences, which differ in length for individual mRNA species. It appears that phosphorolysis of the entire 3'-noncoding region, including the AAUAAA hexanucleotide transcript, does not abolish the translational efficiency of rabbit globin mRNA (277,264).Similar conclusions were drawn when the entire 3'-noncoding sequence was deleted from globin mRNA by doublestranded nuclease, following its hybridization to a cloned cDNA probe (290). Moreover, the 3'-noncoding region does not participate in the formation of the initiation complex, since the interaction between globin mRNA and the initiation factor that binds methionyl tRNAfMetremained unchanged when the entire 3'-noncoding region was removed by phosphorolysis with PNPase (277). The phosphorolytic removal of the poly(A) tails and the entire 3'noncoding regions from interferon mRNA species with the aid of PNPase does not significantly alter the translational efficiency or stability of these molecules when microinjected into Xenopus oocytes (278).Therefore, the AAUAAA hexanucleotide, which is included in the deleted region, and 290. Kronenberg, H. M . , Roberts, B. E., and Efstratiadis, A. (1979). Nucleic Acids Res. 6 , 153.

553

17. POLYNUCLEOTIDE PHOSPHORYLASE

the whole of the 3'-noncoding region, do not appear to contribute to the regulation of interferon mRNA stability in the Xenoprrs system.

H. PNPAsE-DIRECTED LABELING OF POLY N uc L EOTI DES

THE

3'-OH ENDOF

Primer-dependent PNPase has been used to add poly(C) sequences to the 3' terminus of RNA from potato spindle tuber viroid, and the resulting RNA served as template for QP replicase. The poly(G) sequence at the 5' end of the product provides a potential means to separate template from product and to study the properties of both RNA chains (291 ). Polyadenylation of viral RNA species makes them substrates for reverse transcription. The resulting labeled cDNA can then be sequenced, as was carried out with RNAs purified from preparations of vesicular stomatitis virus (292). The ability of PNPase to add poly(A) tails to the 3'-OH end of RNAs (tRNA, 5 S RNA or poly(A)-deficient mRNA) was also utilized for gene mapping. The poly(A) tailing is accomplished by use of a 2- to 3-fold excess of PNPase over RNA and 20-200 I.LMADP. Under these conditions 10-2096 of the RNA molecules acquire a poly(A) tail of about 60-400 residues long. The poly(A)-containing RNA is separated from the nonreacted RNA by oligo(dT)-cellulose chromatography. The in v i m polyadenylated RNA is then hybridized with a linear duplex DNA to which poly(dT) tails, or poly(dBrU) tails have been added with terminal deoxynucleotidyltransferase. The poly(dT) pairs with the poly(A) on the RNA and is readily recognized in the electron microscope (293).

291. Owens, R. A., and Diener, T. 0. (1977). Virology 79, 109. 292. Rowlands, D. J. (1979). PNAS 76, 4793. 293. Engel, J. D., and Davidson, N. (1978). Biochemistry 17, 3883.

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Section IV

RNA Modzficatim

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RNA Methylation DIETER SOLL

LARRY K. KLINE

I. Introduction . . . . . . . . . . . . . . 11. Structures and General Assay Procedure . 111. Specific Methyltransferase Enzymes . . . A. 5-Methylcytidine . . . . . . . . . . B. I-Methyladenosine . . . . . . . . . C. 1-Methylguanosine . . . . . . . . . D. N2-Methylguanosine . . . . . . . . . E. 7-Methylguanosine . . . . . . . . . F. Ribothymidine . . . . . . . . . . .

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

. . . .

. . . .

. . . .

. . . .

. . . .

. . . .

. . . .

. . . .

. . . .

. . . .

. . . .

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

. . G. 5-Methylaminomethyl-2-Thiouridine. . . IV. Conclusion . . . . . . . . . . . . . . . . 1.

. . . . . . . .

. . . .

. . . .

. . . .

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

557 558 559 559 561 561 562 563 564 565 566

Introduction

The methylation of RNA gives rise to a variety of methylated nucleotides, and methylated derivatives of all four major base components of RNA are known. The RNA methyltransferases catalyze the posttranscriptional modification of RNA, and methylated nucleosides are present in rRNA and mRNA as well as in tRNA. The methylation of RNA has been reviewed previously in this series ( I ) . The majority of methylated nucleosides are found in tRNA and have been discussed in the context of tRNA modification (2, 3). A description 1. K e n , S. J., and Borek, E. (1973). “The Enzymes,” 3rd ed.,Vol.IX, Part B, p. 167. Recognition,” (H.

2. Agris, P. F., and SOH, D. S. (1977). I n “Nucleic Acid--Protein Vogel, ed.) p. 321. Academic Press, New York.

3. Nishimura, S . In “Transfer RNA: Structure, Properties and Recognition,” (P. R. Schimmel, D. SOU, and J. N. Abelson, eds.), p. 59. Cold Spring Harbor Laboratory, Cold Spring Harbor, New York. 557 THE ENZYMES,VOL. XV Copyright @ 1982 by Academic Press, Inc. All rights of reproduction in any form reserved. ISBN &12-122715-4

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DIETER SOLL AND LARRY K. KLINE

of bacterial tRNA methyltransferases has also appeared ( 4 ) and contains references to earlier work. An excellent review dealing with the synthesis and possible biological functions of tRNA methylation contains extensive literature references and should also be consulted ( 5 ) . This discussion focuses on the properties of the enzymes responsible for the methylation of tRNA, particularly on enzymes that have been highly purified. The more general area of tRNA modification is considered in a separate chapter in this volume (6), as is the capping of mRNA(7).

II. Structures and General Assay Procedure

The structures of the methylated nucleosides known to occur in RNA are given in Fig. 1. In almost all cases an enzyme that catalyzes the methylation of RNA is assayed according to the following general reaction: RNA

+ [3H-or 14C-methyl]S-adenosylmethionine+

[ITor - 3H-methyl]RNA + S-adenosylhomocysteine

The radioactive RNA reaction product is then isolated, usually by acid precipitation, and the incorporation of methyl groups into RNA is determined, Alternatively, the RNA is isolated by phenol extraction andor DEAE-cellulose chromatography, hydrolyzed, and the resulting radioactive nucleotides are separated and identified by chromatographic procedures. S- Adenosylmethionine serves as the methyl donor in the vast majority of RNA methylations. However, the discovery that the methyl group of ribothymidine in some bacterial tRNA species is derived from a folic acid derivative (8, 9) necessarily requires alternate assay procedures, as discussed in Section II1,F. The substrate RNA is usually derived from a different source than the methyltransferase, since homologous RNA would already be completely methylated. In addition, “methyl-deficient” tRNA, isolated from mutants 4. Greenberg, R., and Dudock, B . S . (1979). “Methods in Enzymology,” Vol. 59, p. 190. 5. Nau, F. (1976). Biochimie 58, 629. 6. Kline, L. K., and Soil, D. S.,Chapter 19, this volume. 7. Shuman, S . , and Hurwitz, J. (1981). “The Enzymes,”Chapter 9, this volume. 8. Delk, A. S . , and Rabinowitz, J. C. (1975). PNAS 72, 528. 9. Schmidt, W., Arnold, H. H . , and Kersten, H. (1975). Nirckic Acids R e s . 2, 1043.

559

18. RNA METHYLATION 0

OH OH

(a) HYCH'

3

HOHYC 0

OH OH

(f)

HOHzC

OH OH

OH OH

FIG. 1. Methylated nucleosides present in RNA: (a) Ribothymidine (rT); (b) 3-methylcytidine (m3C); (c) 5-methylcytidine ( m T ) ; (d) I-methyladenosine (m'A); (e) 1-methylguanosine (m'G); (0 N6-methyladenosine (maA); (g) 7-methylguanosine (m7G);(h) N*-methylguanosine (m2G); (i) NZ,N*-dimethylguanosine (mlG).

of E. coli, has been useful in the detection and isolation of tRNA methyltransferase enzymes (see e.g., 10-12). 111.

Specific Methyltransferase Enzymes

A.

5-METHYLCYTIDINE

The methyltransferase responsible for the formation of 5-methylcytidine (Fig. 1,c) in tRNA has been detected in rat liver (13, 14) 10. Srinivasan, P. R., and Borek, E. (1964). Uioclzernisfry 3, 616. 11. Marinus, M. G., Morris, N . R., So11, D., and Kwong, T. C. (1975). J . Bucterid. 122, 257. 12. Aschhoff, H.J., Elton, H., Arnold, H. H., Mahal, G., Kersten, W., and Kersten, H. (1976). Nucleic Acid3 Res 3, 3109. 13. Rodeh, R., Feldman, M., and Littauer, V. Z. (1967). Uiorhemistr,y 6, 451. 14. Gambaryan, A. S., Venkstern, T. V., and Baev, A. A., (1976). Mol. B i d . ( R e s . ) 10, 846.

5 60

DIETER SOLL AND LARRY K. KLINE

and yellow lupine seed ( 1 3 , and has been partially purified from HeLa cells (16). The HeLa cell enzyme was estimated to be greater than 1000-fold purified and was free of other tRNA methyltransferase activities. The enzyme was isolated from a postribosomal supernatant fraction of the cells, using DEAE-cellulose and CM-Sephadex chromatography. The enzyme is stable at 0" for 5 to 6 months, and for over 1 year at -70". The methyltransferase from HeLa cells has a pH optimum of 7.25 and the molecular weight was estimated to be 72,000 by sucrose gradient centrifugation. Sodium dodecyl sulfate-polyacrylamide gel electrophoresis of the most purified enzyme fractions revealed a number of protein components, indicating the lack of enzyme homogeneity. The methylase enzyme was inhibited 61 and 97% by 100 mM NaCl and 100 mM (NH&S04, respectively. No enzyme activity was detected in the presence of 5 mM magnesium chloride. The assay procedure measured the incorporation of methyl groups from S-aden~syl-[methyl-~H]methionine into a variety of RNA substrates. In all cases the product of the methylation reaction was exclusively msC, as determined by nucleoside analysis of the labeled RNA. The HeLa enzyme was active in methylating both natural and synthetic polyribonucleotides. Esclzerichin coli tRNAPheis the best substrate, but other E . coli and yeast tRNAs also serve as substrates. The HeLa methyltransferase also methylated rRNA, tobacco mosaic virus RNA, and brome mosaic virus RNA. Several synthetic RNAs were also substrates, although poly(C) and DNA were not methylated. Evidence suggesting that the HeLa enzyme preparation was, in fact, a tRNA methyltransferase was indicated by the fact that the enzyme methylated cytidine residues in E. coli tRNAPhe,which are located in the same region in which m5C is found in eukaryotic tRNAs. However, enzyme digestion of the methylated plant viral RNAs did not suggest sequence specificity for the methylation of these substrates. (Both plant viral RNAs have 3' ends that can be aminoacylated by aminoacyl-tRNA synthetases.) An interesting aspect of these studies was that the HeLa enzyme did not methylate a second cytidine residue in E. coli tRNAPhethat occurs in a position that is normally found methylated in yeast tRNAPhe.This observation suggests that either another msC tRNA methyltransferase is present in yeast, or that some structural feature of the E . coli tRNAPhesubstrate is not recognized by the enzyme. The fact that the HeLa msC methyltransferase activity for each type of RNA substrate cosedimented in sucrose gradients 15. Wierzbicka, H . , Jakubowski, H. and Pawelkiewicz, J. (1975). Nucleic Acids Res. 2, 101.

16. Keith, J. M., Winters, E. M . , and Moss, B. (1980). JBC 255, 4636.

18. RNA METHYLATION

561

suggests, but does not prove, the presence of a single enzyme. An understanding of the nature of the substrate specificity of the HeLa enzyme clearly requires further study.

B.

1-METHYLADENOSINE

Methyltransferase enzymes that catalyze the formation of I-methyladenosine (Fig. 1,d) in tRNA have been partially purified from HeLa cells (17) and wheat germ (18). The rat liver enzyme has been purified to homogeneity ( 1 9 ) and the properties of this enzyme are noted below. The rat liver enzyme was purified by ammonium sulfate fractionation followed by column chromatography on DEAE-Sephadex and phosphocellulose. The procedure yielded an 8000-fold purification and the enzyme preparation gave a single band on polyacrylamide gel electrophoresis. The molecular weight of the tRNA adenine- l-methyltransferase was determined to be about 95,000, as judged by gel filtration on Bio-Gel P150. The enzyme catalyzed the formation of l-methyladenine at the invariant adenine residue in the TG$C loop of the E. coli tRNA2G'" substrate. The S-adenosylmethionine methyl donor was found to have a K m of 3 x lo-' M. The enzyme is stable in 40% glycerol at -70" for at least 1 year. The rat liver enzyme requires the addition of cations for activity; 20 to 40 mM putrescine is most effective. In the presence of optimum concentrations of putrescine or spermidine, 1 mM magnesium ion was found to be inhibitory. The methyltransferase enzyme from rat liver catalyzed the formation of m'A in a variety of purified E. coli tRNA species. The K m values for E. coli tRNAfMe'and tRNAB'" were found to be 12 x lo-$ M and 33 x lo-$ M respectively. The data collected in the study suggests that the conformation of the tRNA substrates may play an important role in the enzyme recognition process, in contrast to sequence specificity being the major factor (20). C.

I-METHYLCUANOSINE

The methyltransferase that catalyzes the formation of I-methylguanosine (Fig. 1,e) in tRNA has been purified 6800-fold from rat liver 17. Agris, P. F., Spremulli, L. L., and Brown, G. M. (1974). A B B 162, 38. 18. Schnabel, J . J . , and Chang, S. H., unpublished results. 19. Glick, J. M., and Leboy, P. S . (1977). JBC 252, 4790. 20. Kuchino, Y., and Nishimura, S. (1974). Biochemistry 13, 3683.

562

DIETER SOLL AND LARRY K. KLINE

(21). UsingE. coli tRNA as a substrate, the m'G enzyme was purified by a

series of steps including DEAE-Sephadex, Sephacryl S200, and phosphocellulose column chromatography. The enzyme did not contain ribonucleases and other methyltransferase activities. The molecular weight of the enzyme is about 83,000 as determined by chromatography on Bio-Gel P150. The purified enzyme catalyzed the incorporation of methyl groups from S-adenosylmethionine into an E. coli tRNAfMetsubstrate, other purified E. coli tRNA species being inactive as substrates. This result was not unexpected in view of the location of 1-methylguanine in eukaryotic tRNAs, as well as the prediction of site specificity ( 5 . 2 1 ) . The properties of the m'G enzyme are similar to other methyltransferases in cation requirements; polyamines are stimulatory. The guanine- 1-methyltransferase activity, like the rat liver adenine- 1-methyltransferase (19>, is extremely sensitive to inhibition by S-adenosylhomocysteine; the K f values for the mlG and m'A enzymes are 0.11 and 0.85 p M , respectively.

D . Nz-METHY LG u A N OSI N E Substantial purifications of tRNA methyltransferase enzymes involved in the formation of N2-methylguanosine (Fig. l,h), located at position 10 from the 5' terminus of tRNAs, have been described in chicken embryo (22) and rat liver (21) systems. The chicken embryo enzyme was purified approximately 1000-fold in a two-step procedure using phosphocellulose and S-adenosylhomocysteineSepharose column chromatography. The molecular weight of the enzyme was determined by Sephadex G-200 chromatography to be 77,000. The purity of the m2G methyltransferase enzyme could not be determined; four protein bands were observed in sodium dodecyl sulfate-polyacrylamide gels. The enzymes catalyzed the transfer of methyl groups from S-adenosylmethionine into several E. coli tRNA substrates, including tRNAArg,tRNALe", tRNAfMet,tRNAVa'and tRNAPhe.tRNAPhewas the best methyl acceptor, with a K, of 3 x lo-' M . The m2G nucleotide is located in position 10 of the E. co/i tRNAPhe. The use of S-adenosylhomocysteie affinity chromatography for the purification of methyltransferases as previously noted (22) is an important experimental approach and may be applicable to other methyltransferase enzymes. The m2G tRNA methyltransferase enzyme from rat liver has been purified 6200-fold, although the preparation is not homogeneous (21 ). The 21. Glick, J. M., Averyhart, V. M., and Leboy, P. S. (1978). BBA 518, 158. 22. IZZO, P., and Gantt, R. (1977). Biochemisfrv 16, 3576.

18. RNA METHYLATION

563

fractionation scheme involved DEAE-Sephadex and phosphocellulose chromatography. The molecular weight of the enzyme, determined by gel filtration (Bio-Gel P150), was 69,000. Like the chicken embryo enzyme (22),the rat liver preparation used E. cofi tRNAphe,tRNAVa',and tRNAArg as substrates for the formation of m2Gin position 10 in the tRNAs. The K,,, values for S-adenosylmethionine in the rat liver ( 2 1 ) and chicken embryo (22) systems were similar: 2 and 1.38 p M , respectively. It is noteworthy that the purified rat liver enzyme has maximal activity at concentrations of spermidine (0.1 mM) or putrescine (5 mM) that differ considerably from the results obtained with less purified enzyme preparations ( 2 1 , 2 3 ) .These observations again call attention to the necessity for caution in the interpretation of data obtained with impure enzymes.

E. 7-METHYLGUANOSINE The nucleoside 7-methylguanosine (Fig. 1 ,g) is usually located at position 55 from the 5' end of tRNA (5).B. sirbtilis tRNAfSfet differs from the E. coli tRNAmet in that a guanine residue instead of 7-methylguanine is present in the extra arm of the tRNA sequence. This similarity in structure has allowed the use ofB. subtilis tRNAfMet as a substrate in the purification of the m7G methyltransferase from E. coli (12). The enzyme has been purified some 1000-fold using tRNA-Sepharose affinity chromatography. The elution profile of m7G methyltransferase activity indicates that the enzyme can use both B.subtilis tRNAfMet and undermethylated E. coli bulk tRNA as substrates. (The latter substrate was isolated from an E. coli K12 met- rel- strain). Sephadex G-200 exclusion chromatography resolved the enzyme activity in two peaks having molecular weights 100,000 and 300,000. These studies also detected a m7G methyltransferase activity that could use undermethylated bulk E. coli tRNA, but not the B. sirbtilis tRNAfMet, as a substrate. The results were interpreted as indicating the presence of at least two m7G tRNA methyltransferases present in E. coli, only one of which recognizes the B. sirbtilis tRNAfMetsubstrate. The fact that m7G is present in B. sirbtilis tRNAPhe(24), but not present in B. sirbtilis tRNAf"et also indicates that some structural feature of the tRNAfMe' prevents methyltransferase recognition in B. subtilis. The precise factors that allow only one of the E. coli m7G methyltransferase activities previously described to recognize the B. subtilis tRNAmet substrate are unknown. 23. Leboy, P. S . , and Glick, J. M. (1976). BBA 435, 30. 24. Amoid, H., and Keith, G . (1977). Nircleic Acids Res. 4, 2821.

564

DIETER SOLL AND LARRY K. KLINE

F. RIBOTHYMIDINE Ribothymidine (Fig. 1,a) occurs in the vast majority of tRNAs in the GT+C loop (3, 5, 6, 25). The rT-forming enzyme has been purified to homogeneity from E. coli using polyethyleneimine precipitation, phosphocellulose, and Blue Sepharose affinity elution chromatography (26). The enzyme consists of a single polypeptide chain of molecular weight 40,000. Its pH optimum is 8.4. The K,,,values for its substrates, S-adenosyl-L-methionine and (wheat germ) tRNAGIY,are 12.5 and 1.1 ELM, respectively. The E. coli enzyme catalyzes the transfer of methyl groups from S-adenosylmethionine into tRNA species that contain uridine in the normal rT location. Wheat germ tRNAGIYspecies in particular serve as excellent substrates for the E. coli enzyme (26). In all cases, the methylation gives rise to an rT residue in the GTJIC loop of tRNA. Since this residue is present in crude wheat germ tRNA and known to occur in wheat germ tRNAPhe(27), it is clear that the rT methyltransferase present in wheat germ does not recognize some structural feature of the tRNAGIY species, in contrast to the E. cali rT methyltransferase. These considerations illustrate the difficulties in determining the structural factors of the tRNA that are recognized by the methyltransferase enzymes. [See, for example, a discussion in Refs. (26, 28).] The methyl group of ribothymidine, as well as all other methyl groups present in the methylated nucleosides of tRNA, was long presumed to be derived from S-adenosylmethionine. This generalization is no longer valid. The initial observations that the methyl groups of rT that occur in B. subtilis andS.fureculis tRNAs are derived from a folate derivative (8, 9,29, 30) have led to detailed studies on the mechanism of rT formation in the tRNA of these organisms (31, 32). The enzyme that catalyzes the formation of rT in S. fuecalis has been purified to homogeneity (31) and catalyzes the formation of rT in tRNA 25. Sprinzl, M., Grueter, F., Spelzhaus, A., and Gauss, D . H. (1981).Nucleic Acids Res. 9, rl.

26. Marcu, K. B., Mignery, R. E., and Dudock, B. S. (1977). Biochemistry 16, 797. 27. Dudock, B . S . , Katz, G . , Taylor, E. K . , and Holley, R. W. (1969). PNAS 62, 941. 28. Marcu, K . , Marcu, D., and Dudock, B. (1978). Nucleic Acids Res. 5, 1075. 29. Romeo, J. M., Delk, A. S. , and Rabinowitz, J. C. (1974). BBRC 61, 1256. 30. Kersten, H . , Sandig, L., and Arnold, H. H. (1975). FEBS Leu. 55, 57. 31. Delk, A. S . , Nagle, D . P., Jr., and Rabinowitz, J . C. (1979). I n “Chemistry and Biology of Pteridines” (R.L. Kisliuk and G. M. Brown, eds.), p. 389. Elsevier-North Holland, New York. 32. Delk, A. S., Nagle, D. P., Jr., and Rabinowitz, J. C. (1980). JBC 255, 4387.

565

18. RNA METHYLATION

(31. 3 2 ) according to the following reaction: tRNA(UW)

+ CH,=THF + FADH,

-B

tRNA(TJIC) + THF

+

FAD

As indicated in the reaction above, the enzyme uses the methylene group derived from 5,lO-methylenetetrahydrofolate(CH,=THF) and reduced flavin adenine dinucleotide (FADH2) as the reducing agent. The enzyme has been given the systematic name 5,IO-methylenetetrahydrpfolate :tRNA (uracil-5-)-methyltransferase(FADH,-oxidizing), and the trivial name folate-dependent ribothymidylsynthase (32). The enzyme from S . fuecalis was purified to homogeneity by procedures involving DEAE-cellulose, phosphocellulose, and tRNA-Sepharose column chromatography. The assay procedure is based on the release of tritium from the [5-3H]uridine-labeled tRNA substrate prepared from S. ,fuecnlis grown under folate-free conditions (33). The native enzyme has a molecular weight of 115,000 (determined by Sephadex G-150 column chromatography) and exhibits a single protein band of molecular weight 58,000 in sodium dodecyl sulfate-polyacrylamide gels. The enzyme is specific in the formation of rT in the TJIC loop of tRNA and requires both folate and FADHz. The K , values of the bulk tRNA and folate substrates are 2.5 p M and 1 mM, respectively (31). This folate-dependent formation of rT occurs in S. fuecufis, B. subtilis, and other gram-positive organisms. However, the methyl donor for rT synthesis in rRNA may be different. It was shown in M.fysodeikricus that S-adenosylmethionine is the precursor for rT found in 23 S rRNA ( 9 ) . These studies have clearly illustrated an alternative biosynthetic pathway for the origin of the methyl groups of rT in tRNA. Whether other methylated components of RNA arise in a similar manner is an open question. It is noteworthy that the S.faecalis tRNA is also a substrate for the S- adenosylmethionine-dependent E. coli methyltransferase (34).It will be interesting to learn if other tRNA substrates, such as the wheat germ tRNAGiYspecies, are also recognized by the S. fuecaiis enzyme.

G . 5-METHYLAMINOMETHYL-2-THIOURIDINE The methyltransferase that catalyzes the addition of a methyl group to form 5-methylaminomethyl-Zthiouridinein tRNA has been purified from 33. Delk, A. S . , Nagle, D. P.,Jr., and Rabinowitz, J. C. (1979). BBRC 86, 244. 34. Delk, A. S . , Romeo, J . M . , Nagle, D. P.,Jr., and Rabinowitz, J. C. (1976). JBC 251, 7649.

566

DIETER SOLL A N D LARRY K. KLINE

E. coli (35). The properties of this enzyme are discussed in another chapter in this volume (6).

IV.

Conclusion

Although the methyltransferases were discovered almost 20 years ago (36), progress in the purification of the enzymes has been slow. It is evident that problems in the selection of suitable substrates and the instability of the enzymes have been the major factors. However, the past several years have been productive in this area. The utilization of tRNAaffinity chromatography as well as the availability of purified tRNA species of known sequence have been key elements in this progress. The discovery of the folate-dependent methylation of tRNA was an unexpected and novel finding. It appears that our knowledge of the substrate specificities and recognition parameters of the methyltransferase enzymes will soon be greatly enhanced. The isolation of RNA methylase mutants (37-40) should help our understanding of the biological regulation and function of this important class of RNA modification enzymes.

35. 36. 37. 38. 39.

Taya, Y., and Nishimura, S. (1973). BBRC 51, 1062. Fleissner, E., and Borek, E. (1963). Biochemistry 2, 1093. Phillips, J. H . , and Kjellin-StrHby, K. (1978). J M B 26, 509. Bjork, G. R., and Isaksson, L. A. (1970). J M B 51, 83. Marinus, M. G., Morris, N. R., Soll, D., and Kwong, T. C . (1975). J . Bncteriol. 122,

251.

40. Bjork, G . R., and Kjellin-Striby, K. (1978). J . Borteriol. 133, 508.

Nucleotzde Modajication in RNA LARRY K. KLINE

DIETER SOLL

I. Introduction . . . . . . . . . . . . . . 11. Modification of Uridine . . . . . . . . . A. Structures of Uridine Derivatives . . . B. Specific Uridine Modifying Enzymes . 111. Modification of Cytidine . . . . . . . . . IV. Modification of Adenosine . . . . . . . . A. Structures of Adenosine Derivatives . . B. Specific Adenosine Modifying Enzymes V. Modification of Guanosine . . . . . . . . A. Structures of Guanosine Derivatives . . B. Specific Guanosine Modifying Enzymes VI. Conclusion . . . . . . . . . . . . . .

1.

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

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

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

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

567 568 568 570 574 575 575 575 578 578 578 582

Introduction

The isolation and characterization of the enzymes responsible for the modification of RNA bases poses a number of difficult yet interesting and challenging problems. Although over 50 modified nucleotides have been identified in RNA, most of which occur in tRNA, the enzymology of RNA base modification is in its infancy. Genetic and biochemical evidence indicate that all modified bases in RNA are formed by an enzymatic modification of the polynucleotide transcript. A major problem in the study of the modification enzymes is 567 THE ENZYMES. VOL.X V Copyright @ 1982 by Academic Press, Inc. All rights of reproduction in any form reserved. ISBN 812-1227 15-4

568

LARRY KLINE AND DIETER SOLL

one of obtaining a suitably unmodified or undermodified RNA that can be used as substrate to aid detection or purification of the modifying enzymes. In addition, the temporal order in which the various base modifications occur within a single RNA molecule is unknown. In spite of these problems some progress in our understanding of the reactions leading to the formation of the modified bases in RNA has been made. In this chapter we attempt to cover the literature through early 1980. Our purpose is to focus on the isolation and properties of the enzymes responsible for RNA base modification, with the exception of methylation which is covered in a separate chapter in this volume (f ), as is the capping of mRNA (2). The reader is also referred to the discussion of tRNA processing in this volume ( 3 ) ,since the temporal sequence of tRNA base modification will no doubt be related to the maturation of the RNA transcript. We include a listing of all modified bases found in RNA (with the exception of bases formed by the addition of only methyl groups). This listing illustrates the variety of modified bases present in RNA and serves as a reminder that our knowledge of the biosynthesis of these compounds is far from complete. Previous reviews dealing with the more comprehensive areas of structure, function, and biosynthesis of modified nucleotides have been published (4-6). II. Modification of Uridine

A. STRUCTURES OF URIDINE DERIVATIVES Figure 1 lists the known modified uridine nucleotides found in RNA. They occur exclusively in tRNA ( 6 ) ,with the exception of pseudouridine, $, which has also been found in rRNA (7-f0),5 and 5.8 S RNAs from 1. So11, D. S . , and Kline, L. K., Chapter 18, this volume. 2. Shuman, S.,and Hurwitz, J. (1981). Chapter 9, this volume. 3. Kole, R., and Altman, S . , Chapter 14, this volume. 4 . Agris, P. F., and Soll, D. S . (1977).fn “Nucleic Acid-Protein Recognition,” (H. Vogel, ed.), p. 321. Academic Press, New York. 5 . McCloskey, J. A., and Nishimura, S. (1977). Accounts Clzern. Res. 10, 403. 6. Nishimura, S. (1980). I n “Transfer RNA: Structure, Properties and Recognition” (P. Schimmel, D. So11, and J. Abelson, eds.) p. 59. Cold Spring Harbor Laboratory, Cold Spring Harbor, New York. 7. Dubin, D. T., and Gunlap, A. (1967). BBA 134, 106. 8. Fellner, P. C. (1969). EJB 11, 12. 9. Hall, R. (1971). “The Modified Nucleoside in Nucleic Acids.” Columbia Univ. Press, New York. 10. Amaldi, F., and Attardi, G. (1968). JMB 33, 737.

569

19. NUCLEOTIDE MODIFICATION IN RNA

2.,Hl:oH

B

0

OH OH

S

d

HOHYC

OH OH

OH OH

(h)

H

0 N

8 $

,

~

~

y

~

~

~

~

,

~

~

~

~

S'lN

OH OH

OH OH

OH OH

FIG. 1. Modified uridine derivatives: (a) Pseudouridine (JI); (b) dihydrouridine (D); (c) 5-methoxyuridine (mo5U); (d) uridin-5-oxyacetic acid (V); (e) uridin-5-oxyacetic acid methyl ester (mV); (f) 5-(methoxycarbonylmethyl)uridine(mcm5U); (9) S-(carboxymethylaminomethy1)uridine (cmnmg); (h) 4-thiouridine (s4U); (i) 5-methyl-2-thiouridine (m5sZU); (j) 5-(methoxycarbonylmethyl)-2-thiouridine(rncm5szU);(k) 5-methylaminomethyl-2-thiouridine (mnm5s2U);(1) 5-carboxymethylaminomethyl-2-thiouridine(cmnm5s2U);(m) 3 4 3 amino-3-carboxypropy1)uridine(acp3U).

570

LARRY K L l N E A N D DIETER SOLL

yeast (11, 12), and animal cell nuclear RNA (13). The great variety of enzymatic modification that occurs on the RNA uracil residues is evident from the complexity of structures obtained. Oxidation of the heterocyclic base (c-e) as well as reduction (b) is observed. The substitution of sulfur for the oxygen atoms at either the 2 or 4 ring position also occurs (h-1). It is noteworthy that at least some of the modified uracil nucleotides must arise by the action of more than one enzyme, since more than one ring position is modified (i-I). It is likely that modifications containing more than one functional group are also formed in a series of enzymatic steps (d-g, j-I). The enzymes responsible for the formation of the modified bases must possess an extremely high degree of specificity, since the ring position and modifying group are specific to each nucleotide, and the exact position of the nucleotide in the RNA chain is a determinant. This latter factor is an important consideration in the selection of possible substrates, as noted in the following sections.

B.

SPECIFIC U R I D I N E M O D I F Y I N G

ENZYMES

I. Psertdoiiridine Almost all tRNAs contain $ (Fig. l,a), the exceptions being the methionine initiator tRNAs from yeast and Neurosporu. The $ residues occur most often in the anticodon loop and stem and in the “TJIC” loop (6). The formation of JI in tRNA is catalyzed by at least two enzymes, pseudouridylate synthetase I (PSI), which is responsible for the formation of $ in the anticodon region, and pseudouridylate synthetase I1 (PSII), which is presumably involved in the formation of other $ residues in tRNA. Pseudouridylate synthetase I has been purified 1000-fold from Salmonella typhimirriirm (14). The enzyme is the product of thehisT gene i n s . typhimirrium ( 1 5 ) , and studies of this enzyme have been facilitated by the use of hisT mutants, which produce tRNA species that lack the $ modification in the anticodon region but contain the $ modification in the T$C 1 1 . Miyazaki, M. (1974). J . Biochem. (Tokyo) 75, 1407. 12. P.ubin, G . M. (1973). JBC 248, 3860. 13. Shibata, H., Ro-Choi, T. S., Reddy, R., Choi, Y . C . , Henning, D., and Busch, H. (1975)..me 250,3909. 14. Arena, F., Ciliberto, G . , Ciampi, S . , and Cortese, R. (1978). Nitcleic Acids Res. 5, 4523. 15. Cortese, R . , Kammen, H. O . , Spengler, S. J . , and Ames, B . N . (1974).JBC 249, 1103 (1974).

57 1

19. NUCLEOTIDE MODIFICATION IN RNA

loop (16). The strategy for obtaining a tRNA substrate for assay of PSI has been to isolate tRNA from [5-3H]uridine-labeledhisT S . ryphimurium cultures (14, 15. 17). The activity of the PSI enzyme can then be followed by a tritium release assay (15, 17) according to the following scheme: tRNA([5-3H]uridine)

+ 3H (U- 4)

tRNA

The reaction mixture is treated with Norit A charcoal which absorbs the tRNA but not the released 3H, and is then filtered. The released 3H appears in the filtrate and is a measure of J, formation and hence PSI activity. Using the tritium release assay, the PSI enzyme from S . typhimurium has been purified by a sequence of steps utilizing streptomycin sulfate precipitation, DEAE-cellulose, and Sephadex G- 100 chromatography. The PSI enzyme was obtained 90% pure in 10-15% yield, as judged by SDS-polyacrylamide gel electrophoresis and assuming the major protein band was PSI (14). The PSI enzyme eluted from Sephadex G-100, indicating a molecular weight of about 50,000. A second PSI activity eluted in the 10,000- 15,000 dalton region. The two activities were not interconvertible and the relative amount of activity "varied significantly from preparation to preparation." The 50,000 dalton PSI activity (1000-fold purification) could be converted to a dimer in the presence of tRNA, although there was no evidence of a tRNA-enzyme complex, as judged by centrifugation in glycerol gradients in the presence of labeled tRNA. Both pseudouridylated and unpseudouridylated tRNA induced this phenomenon. The 50,000 dalton PSI enzyme was relatively stable at 4" (-50% activity after 40 days storage) and full activity could be restored by preincubation with mercaptoethanol. The importance of SH groups for enzyme activity was indicated by enzyme inhibition in the presence of iodoacetamide. Using 3H-labeled tRNATYrisolated from hisT mutants, the mutantderived tRNATYrwas converted to wild-type tRNATYrby PSI in vitro, as judged by chromatography on RPC-5 (14). Similar results with partially purified enzyme have been obtained using E. coli tRNATYrprecursors (18), and with E. coli extracts on mutant tRNALe"(19). The general picture of pseudouridine formation that emerges from the 16. Singer, C . E . , Smith, G . R., Cortese, R . , and Arnes, B . N. (1972). Natrrre N e w B i d . 238, 72. 17. Mullenbach, G . T., Kammen, H . O., and Penhoet, E. E. (1976). JBC 251,4570. 18. Ciampi. M . S . , Arena, F., and Cortese, R. (1977). FEES Lett. 77, 75. 19. Allaudeen, H. S., Yang, S. K., and So11, D. (1972). FEBS Lett. 28, 205.

572

LARRY KLINE AND DIETER SOLL

studies previously noted, is that in tRNA, the J, residues located in the anticodon loop are formed by the action of PSI, probably at the tRNA precursor stage. The J, residues located in the T W G loop are formed by the action of PSII, an enzyme activity as yet uncharacterized. The reason for the apparent instability of PSII (18) is unclear. It is clear that the enzymology of J, formation in RNA requires further work. Pseudouridine formation in tRNA has been detected in extracts of a variety of eukaryotic cell lines (20), but the purification and properties of these activities have not yet been published. 2. 4-Thiouridine

The presence of sulfur-containing uridine residues in tRNA (Fig. 1, h-1) is well known (4, 3,although the enzymes responsible for the formation of these nucleosides are, for the most part, uncharacterized. One of the systems first investigated was the formation of 4-thiouridine (Fig. 1 , h) in E. coli (21). A sulfurtransferase system was partially purified from E. coli extracts and contained two activities (Factor A and Factor C) required for the transfer of sulfur from [35S]cysteineinto tRNA. The tRNA substrates for the reaction were isolated either from E. coli B or from an E. coli strain that contained tRNA with a lowered sulfur content (produced by sulfur depletion in a cysteine-requiring strain). The sulfurtransferase assay involved the incubation of tRNA substrate, [35Slcysteine,ATP, and Factors A and C in buffer that contained Mgz+. Sulfur incorporation was determined following extraction of the incubation mixture with phenol, discharge of aminoacylated tRNA at alkaline pH, and subsequent alkaline hydrolysis of the RNA product. The resulting nucleotide mixture was then separated on DEAE-cellulose and the radioactive 4-thiouridine was identified. Transfer of 35S label from [35S]cysteineinto tRNA to form 4-thiouridine was dependent upon both Factors A and C, both factors being heat-labile. The function of either Factor A or C alone was not clear. The sulfur donor in this sytem is apparently cysteine (P-mercaptopyruvate being inactive) in contrast to other systems noted in Section II,B,3 (22, 23). The sulfurtransferase system from E. coli catalyzes the incorporation of 35S from cysteine exclusively into 4-thiouridine when E. coli tRNA is the substrate. Substitution of yeast tRNA as a substrate resulted in not only the formation of 4-thiouridine but other unidentified sulfur-labeled nucleotides as well (21). The use of purified tRNA species in this system should help clarify the 20. 21. 22. 23.

Muhlenbach, G. T., Kammen, H. O., and Penhoet, E. E. (1976). JBC 251, 4570. Abrell, J . W., Kaufman, E. E., and Lipsett, M. N. (1971). JBC 246, 294. Wong, T., Weiss, S. B . , Eliceiri, G., and Bryant, J. (1970). Biochemistry 9, 2376. Wong, T., Harris, M. A., and Jankowicz, C. (1974). BiuchPmistry 13, 2805.

19. NUCLEOTIDE MODIFICATION IN RNA

573

products and/or site(s) of thiolation. A "rapid" assay procedure for the E. coli sulfurtransferase enzymes has also been published (24). 3. Thiolclted Pyrimidines

The incorporation of 35Sinto tRNA in v i m has also been investigated in two other systems. An enzyme preparation from Bacillus subtilis (23) was found to catalyze the incorporation of 3sS from either [35S]cysteineor [3sSlP-mercaptopyruvateinto tRNA; the K, for P-mercaptopyruvate was 200-300 times smaller than that for cysteine. These observations are not in conflict with those of the E. coli system, however (21), since the product of the sulfurtransferase preparation from B. subtilis was not 4-thiouridine, but other thiolated nucleotides. The identity of the nucleotide products is unknown. The assay system utilized yeast tRNA as a substrate and required ATP. Ribosomal RNA did not serve as a substrate for the incorporation of sulfur, although denatured salmon and calf thymus DNA were suitable acceptors. The thiolated nucleotides formed from the DNA substrates were not examined further. A sulfurtransferase preparation has also been isolated from rat brain tissue (23). The properties of the enzyme fraction are similar to the B. subtilis preparation (22) in that the reaction involves the transfer of 35S from P-mercaptopyruvate into tRNA in the presence of ATP and a divalent metal ion (Mg2+or Mn2+). Like the B. subtilis system, the incorporated 35S was not found in 4-thiouridine, but in other unidentified products. The results of these studies on the formation of thiolated nucleotides in tRNA clearly indicate that additional work is required, particularly with regard to the identification of the nucleotide products formed. The utilization of purified tRNA substrates should simplify the identification of the thiolated products and facilitate the purification of the enzyme(s) responsible for their synthesis. The sequence and structural parameters of the tRNA substrates recognized by the sulfurtransferase enzymes may then be clarified. 4. Other Modified Uridine Nucleosides 0. 3-(3-Amino-3-cnrbo.rypropyl)irriidine.The in vitro formation of 3(3-amino-3-carboxypropy1)uridine(acp3U) (Fig. 1, rn) has been observed in E. coli extracts (25). Using E. coli methyl-deficient tRNAPheas a substrate, the incorporation of radioactivity from either S-adenosyl-L[~arboxyl-'~C]methionineor S-adenosyl-~-[2-~H]methionine was observed. Reisolation of the labeled RNA followed by hydrolysis and

24. Kayne, M . S. , and LaBone, T. (1979). Anal. Biocliem. 98, 146. 25. Nishimura, S . , Taya, Y., Kuchino, Y., and Ohashi, Z. (1974). BBRC 57, 702.

574

LARRY KLINE AND DIETER SOLL

two-dimensional TLC indicated the formation of the acp3U nucleotide. The enzyme(s) that catalyzes the formation of acp3U has not been purified, although the results of the previously described experiments indicate that the 3-amino-3-carboxypropyl group is derived from the methionine of S-adenosylmethionine and is transferred intact to a specific uridine residue in the tRNA substrate. b. 5-Methylaminomethyl-2-thiouridine. The tRNA methylase that catalyzes the formation of the terminal methyl group in 5-methylaminomethyl-Zthiouridine (mnm5s2U) (Fig. 1, k) has been purified to near homogeneity (26). This provides one of the few examples in which a tRNA-modifying enzyme has been completely purified. The purification of the methylase from E. coli involved DEAE-cellulose chromatography followed by affinity chromatography that utilized Sepharose-bound tRNAGLU (E. coli tRNAG1"contains the mnm5s2Unucleotide in the first position of the anticodon). The enzyme fraction eluted from the affinity column was nearly homogeneous, as judged by SDS-gel electrophoresis. The enzyme catalyzed the transfer of methyl groups from [l4CC] methyl-labeled S-adenosylmethionine into a methyl-deficient tRNAG'". The labeled methyl group of the tRNA product was located only in the mnm5szUnucleotide, as determined by hydrolysis of the tRNA and two-dimensional TLC of the resulting nucleotides. The actual location of the rnnm5s2Uresidue in the tRNA, as determined by RNase TIdigestion, was in the predicted site location in the tRNA fragment that contained the anticodon region. This study provides an important illustration of the utilization of purified tRNA species as substrates and the resulting specificity of nucleotide modification. The results of the study also indicate that affinity chromatography using tRNA-Sepharose columns may be applicable to the purification of other RNA modification enzymes.

111.

Modificution of Cytidine

Figure 2 lists the modified cytidine derivatives identified in RNA. The cytidine derivatives that contain only methyl group modifications are discussed elsewhere in this volume ( I ) . We are not aware of published reports on the enzymology of the modifications shown in Fig. 2, although it seems possible that 2-thiocytidine (Fig. 2, b) could be one of the unidentified thiolated nucleotides (22, 23) discussed previously in the sulfurtransferase systems. 26. Taya, Y.,and Nishimura, S . (1973). BBRC 51, 1062.

19. NUCLEOTIDE MODIFICATION IN RNA

575

8

HNCCHi

irH2

OH OH

(a)

(b)

FIG. 2. Modified cytidine derivatives: (a) N4-acetylcytidine (ac4C); (b) 2-thiocytidine (ST).

IV.

Modification of Adenosine

A.

STRUCTURES OF ADENOSINE DERIVATIVES

The modified adenosine nucleosides known to occur in RNA are shown in Fig. 3. The variety and number of adenosine modifications suggest that several enzymatic steps may be involved in the formation of a given modified adenosine nucleotide. The modified adenosine residues occur most often in the first position or on the 3' end of the anticodon of tRNA (4,5).

B.

SPECIFIC ADENOSINE MODIFYING ENZYMES

1. N'2sopenrenyladenosine The initial in live studies on the biosynthesis of N'4sopentenyladenosine (PA) (Fig. 3,c) indicated that the isoprene side chain of i6A was derived from mevalonic acid (27, 28). Subsequent in virro experiments in yeast and rat liver ( 2 9 ) , as well as E. coli (30), demonstrated that the isopentenyl side chain donor was isopentenylpyrophosphate. The enzyme responsible for the attachment of the 5-carbon side chain to an adenosine residue in tRNA has been partially purified from yeast (31) and E. coli ( 3 2 , 3 3 ) .The assay procedures used in these systems involved the incubation of radioactively labeled A'-isopentenylpyrophosphate (formed in a preliminary incubation with A3-isopentenylpyrophosphate 27. 28. 29. 30. 31. 32. 33.

Peterkofsky, A. (1968). Biochemi.stry 7 , 472. Fittler, F., Kline, L. K., and Hall, R. H. (1968). Biochemistry 7, 940. Fittler, F., Kline, L. K., and Hall, R. H. (1968). BBRC 31, 571. Bartz, J. K., Kline, L. K., and So11, D. (1970). BBRC 40, 1481. Kline, L. K., Fittler, F., and Hall, R. H. (1969). Biochemisfry 8, 4361. Bartz, J., and So11, D. (1972). Biochimie 54, 31. Rosenbaum, N., and Gefter, M. (1972). JBC 247, 5675.

576

LARRY KLINE AND DIETER SOLL CH, NHCH~CH=C’ ‘CHI

N-N

52

HOHtC

OH OH

OH OH

HOH’c& , , OH OH

FIG. 3. Modified adenosine derivatives: (a) 1-Methylinosine (m’1); (b) inosine (I); (c) Ne-isopentenyladenosine (PA); (d) 2-methylthio-Ne-isopentenyladenosine(msZi6A); (e) N-[(9-~-~-ribofuranosylpurin-6-yl)carbamoyl]threonine (PA); (f) N-[9-P-D-ribofuranosylpurin-6-yl)N-methylcarbamoyl]threonine(&A); (9) N-[N-[(9-/3-~-ribofuranosylpurin-6yl)carbamoyl]threony1] 2-amido-2-hydroxymethylpropane-l-3-diol.

and an isomerase isolated from pig liver) with the tRNA substrate and enzyme. Reisolation of the radioactive RNA product by either DEAEcellulose chromatography (31) or phenol extraction (32) and subsequent hydrolysis indicated the formation of PA. The tRNA substrate used in these assays was either a permanganate-treated tRNA [the permanganate removes some of the isoprene side chains present in mature tRNA (31)] or, in the case of the E. coli enzyme, a tRNA substrate isolated from mycoplasma (30, 32) or undermodified E. coli su,’ tRNATY’(33). Mycoplasma tRNA is known not to contain i6A and therefore serves as a substrate for the E. coli enzyme. The best substrate for the E. coli enzyme was a purified rat liver tRNAP, which does not contain PA; however, it has the same primary sequence in the anticodon loop and stem as rat liver tRNAY, which is known to contain i6A next to the anticodon (32). TheE. coli enzyme has been purified approximately 350-to 550-fold (32, 33) and has a molecular weight of 50,000-60,000 as determined by chromatography on Sephadex GlOO (32) or glycerol gradient centrifugation (30). The E. coli isopentenyltransferase enzyme uses only tRNA substrates; E. coli ribosomal RNA, f2 RNA, and polyadenylic acid do not serve as acceptors of the isopentenyl group. The modification enzyme

19. NUCLEOTIDE MODIFICATION IN RNA

577

therefore appears to be specific for tRNA. Both the yeast (31)and E. coli (32,33) enzymes have a pH optimum of about 8 and require a divalent cation for activity. The enzymes require sulhydryl groups for activity as evidenced by their sensitivity to hydroxymercuribenzoate. Studies with the E. coli enzyme indicate that the tRNA isopentenylation reaction is not reversible (32). 2. 2-Methylthio-N6-Isopentenyludenosine The sequence of steps leading to the formation of 2-methyl-

thio-N6-isopentenyladinosine(ms2i6A) (Fig. 3, d) in E. coli has been determined (34-36). Although the enzymes have not been purified, the studies illustrate that at least three separate intermediates of a modified nucleoside in tRNA have been identified. Escherichia coli SUJ umber suppressor tyrosine tRNA contains ms2i6Anext to the anticodon and structural intermediates of this nucleoside have been detected in tRNATYr species (34). Likewise tRNA isolated from a relaxed E. coli cys- merstrain (grown in the absence of these amino acids) contains msZi6A,i6A and presumably s T A (36).The data indicate that the steps leading to ms2i6A formation are isopentenylation, thiolation, and methylation, respectively. The final methylation was demonstrated in vitro utilizing S- adenosylmethionine as the methyl donor (35).

3. N-[ (9-/3-~-Ribofuranosylpurin-6-yl)carbumoyl]threonine The formation of N-[(9-~-~-ribofuranosyIpurin-6-yl)carbamoyl]threonine (PA) (Fig. 3, e) has been studied in vivo (37,38) and the results indicated that the side chain of t6A is derived from threonine. In vitro experiments in E . coli (39,40) led to .a partial purification of the PAforming enzyme. The assay consisted of measuring the incorporation of L-[ ''C]thre~nine into tRNA utilizing t6A-deficient tRNA as a substrate (40).The tRNA substrate deficient in t6A was isolated from a threoninestarved culture of an E. coli strain, which is a threonine auxotroph and also a relaxed control mutant. The substrate was periodate-treated before use to destroy the terminal 3'-ribose and thus prevent the threonineaminoacylation reaction. The assay required ATP and Mg2+, and was dependent upon the presence of bicarbonate. 34. 35. 36. 691. 37. 38. 39. 40.

Gefter, M. L., and Russell, R. L. (1969). JMB 39, 145. Gefter, M. L. (1969). BBRC 36, 435. Agris, P., Armstrong, D. J . , Schafer, K. P., and SOU, D. (1975). Nucl. Acids Res. 2, Powers, D. M., and Peterkofsky, A. (1972). BBRC 46, 831. Chheda, G., Hong, C . , Piskorz, C., and Harmon, G. (1972). Biorhem. J . 127, 515. Korner, A , , and SOU, D. (1974). FEES Lett. 39, 301. Elkins, B. N . , and Keller, E. (1974). Biochemistry 13, 4622.

578

LARRY KLINE AND DIETER SOLL

The PA-forming enzyme was isolated from sonicated E. coli cells after ammonium sulfate precipitation and chromatography on Bio-Gel A. The formation of t6A in this sytem had a pH optimum of 7.7-8.2. The enzyme activity eluted from a Bio-Gel P column corresponding to a molecular weight of 50,000-60,0OO,although a loss of 75-90% of the enzyme activity was observed. The enzyme catalyzed the bicarbonate-dependent incorporation of threonine into the tRNA substrate. Hydrolysis of the tRNA product and subsequent electrophoretic separation of the labeled products indicated that both labeled bicarbonate and labeled threonine were incorporated into PA. Other PA-like nucleosides have been detected in tRNA [see Ref. (do)]. These nucleosides contain glycine or serine instead of threonine. An interesting observation from the studies (40) is that [14C]glycinecan also be incorporated into tRNA, and that unlabeled glycine inhibits the incorporation of ~-['~C]threonine into the tRNA substrate. These observations suggest that a single enzyme may be responsible for the formation of the t6A-like nucleosides in tRNA. V.

Modification of Guanosine

A.

STRUCTURES OF GUANOSINE DERIVATIVES

Figure 4 lists modified guanine structures found in RNA. The modified guanine structures are the most complex nucleotides from a structural point of view. Indeed the modified nucleoside Q (Fig. 4,d) is, in fact, not a modified guanine at all, but contains a 7-deazapurine nucleus in place of the purine ring system.

B. SPECIFIC GUANOSINE MODIFYING ENZYMES 1. Base Y

The modified base Y, or wyeosine, (Fig. 4,a) and related structures (Fig. 4,b,c) are nucleotides found adjacent to the 3' end of the anticodon in the tRNAPhespecies present in eukaryotic cells (4-6). Studies on the biosynthesis of Y have been restricted to in vivo experiments (41, 42). Utilizing guanine-requiring mutants of yeast, the results indicate that l4C-1abeled guanine is incorporated into base Y in yeast tRNA. These results were drawn from experiments in which the specific 41. Li, H. J . , Nakanishi, K . , Grunberger, D., and Weinstein, I. B. (1973). BBRC 55, 818. 42. Muench, H. J . , and Thiebe, R. (1975). FEES Lett. 51, 257.

19. NUCLEOTIDE MODIFICATION IN RNA

579

FIG.4. Modified guanosine and related derivatives: (a) Base Y (yw); (b) base peroxy Y (oyw); (c) base Yt (W); (d) R = H Q (gueuosine or Quo),; R = P-D-mannosyl (manQ); R = P-D-galactosyl (galQ).

activities of the 14C-labeled GMP and Y were determined following the hydrolysis of the in vivo [‘4C]guanine-labeledtRNA. In a similar fashion it was shown that the third ring of the Y structure is formed in yeast (42) from the 3-amino-3-carboxypropyl of methionine, whereas in mammalian cells lysine is involved in its formation (43). 2. Base Q The biosynthesis of Q, or queuosine, (Fig. 4,d) is one of the beststudied as well as most unusual examples of RNA base modification. The initial observations (44, 45) that radioactively labeled guanine is incorporated into rabbit reticuloyte tRNA in the absence of RNA synthesis led to the discovery of a novel “RNA guanylation” or “guanine insertion” reaction (46-48). Subsequent work with purified enzymes isolated from rabbit erythrocytes (49, 50), E. coli (51, 52), and rat liver (53) indicated that queuine, the base of Q (or a precursor of base Q), is inserted into tRNA in place of guanine in a transglycosylation reaction. This reaction is unique 43. Pergolizzi, R. G., Engelhardt, D. C., and Grunberger, D. (1979). Nucleic Acids Res. 6, 2209. 44. Hankins, W. D., and Farkas, W. R. (1970). BBA 213, 77. 45. Farkas, W. R . , Hankins, W. D., and Singh, R. (1973). BBA 294, 94. 46. Farkas, W. R., and Singh, R. (1973). JBC 248, 7780. 47. Dubrul, E. F., and Farkas, W. R . (1976). BBA 442, 379. 48. Okada, N . , Yasuda, T., and Nishimura, S. (1977). Nucleic Acids Res. 4, 4063. 49. Howes, N . K . , and Farkas, W. R . (1978). JBC 253, 9082. 50. Katze, J . R.,and Farkas, W. R . (1979). PNAS 76, 3271. 51. Okada, N., and Nishimura, S. (1979). JBC 254, 3061. 52. Okada, N., Noguchi, S., Kasai, H., Shindo-Okada, N., Ohgi, T., Goto, T., and Nishimura, S. (1979). JBC 254, 3067. 53. Shindo-Okada, N., Okada, N., Oghi, T., Goto, T., and Nishimura, S. (1980). Biochemistry 19, 395.

580

LARRY KLINE AND DIETER SOLL

in tRNA modification in that it involves insertion of a modified base into tRNA by cleavage of the N-glycosidic bond; the phosphodiester bond of the polynucleotide chain remains intact. However, the cleavage of an N-glycosidic bond is also involved in the formation of pseudouridine (151, although only the “simple” rotation of the uracil base is involved in the formation of the C-C bond between the C-5 of the pyrimidine ring and the C-1of the ribose. To avoid ambiguity, we refer to the enzyme responsible for the formation of base Q in tRNA as “tRNA:guanine transglycosylase” (TRGT) in congruence with other publications (53, 54). The TRGT enzyme preparations from rabbit erthrocytes (49) and E. coli (51) have been purified to homogeneity. The rat liver enzyme has also been extensively purified (53). The properties of these enzymes are listed in Table I. The specific activities of the TRGT enzyme preparations were omitted because assay conditions are not directly comparable. The assay method used in the enzyme purification procedures involved the incubation of 3H- or 14C-labeledguanine, unfractionated yeast tRNA, buffer, and enzyme. Following incubation and acid precipitation of the tRNA, the incorporation of labeled guanine into the tRNA substrates was measured. The methodology utilized in the rabbit erythrocyte purification included ammonium sulfate precipitation of the erythrocyte hemolysate followed by sequential column chromatography on Sephacryl S200, DE-32 cellulose, Blue Sepharose, and phosphocellulose. The 2600-fold purification resulted in a 5% enzyme yield (49). The E. coli enzyme was purified 5000-fold using ammonium sulfate precipitation followed by DEAEcellulose, DEAE-Sephadex A50, phosphocellulose, and Sephadex G200 column chromatographic procedures (51). The yield of the E. coli enzyme was about 30%. The partially purified rat liver enzyme was isolated by DEAE-cellulose and phosphocellulose chromatography. The 250-fold purification of the rat liver enzyme was accomplished with about a 55% yield. The modified nucleoside queuosine occurs in tRNA species in which queuine is exchanged for guanine in the first anticodon position, namely, tRNATYr,tRNAHiS,tRNAASp,and tRNAAsn. The mechanism by which the nucleoside Q is formed in the tRNA species in the rabbit erythrocyte and rat liver systems appears to be the exchange of queuine for guanine in the tRNA substrate. This conclusion is obtained by inspection of the K, values in Table I. This mechanism would predict that free queuine should be present in extracts of these cell sys54. Crain, P. F., Sethi, S. K . , Katze, J. R . , and McCloskey, J. A. (1980). JBC 255, 8405.

19. NUCLEOTIDE MODIFICATION IN RNA

58 1

TABLE I PROPERTIES OF tRNA: G U A N I NTRANSGLYCOSYLASES E

Property Molecular weight of native enzyme (method) Subunit structure (MW of subunits) K,, Guanine ( M ) K,, Queuine ( M ) K,, 7-(Aminomethyl)7-deazaguanine(M) K , (tRNA substrate) pH optimum Enzyme storage

Rabbit erythrocyten 104,000

(Sephacryl S200)

Rat liverb

80,000 58,000d (Sephadex (3200) (Sephadex G200)

Yes (60,000 + 43,000) 1.5 x 10-7 4.5 x 10-se

E. colic

No 8.3 x 10-7 2.9 x 10-7 2.1. x lo-'

3.3 x 10-9 (Yeast tRNA*SP) 7.4 7.3 10% Glycerol at 5093 Glycerol at -20" -80" (25% IOSS of activity per month)

5.3 x 10-8

Not a substrate 1.4 x

7.0 50% Glycerol at -20" (no activity loss in 3 months)

From references 45, 46. 50. From reference 49. From references 47 and 58. A MW of 46,000 was found using SDS-polyacrylamide gel electrophoresis. Value is inhibition constant of queuine in guanine exchange reaction.

tems. Queuine has, in fact, been isolated from bovine amniotic fluid (50, 5 4 ) . Based on our present data, the biosynthesis of Q in mammalian systems therefore appears to be the incorporation of queuine into tRNA catalyzed by the TRGT enzyme. The biosynthetic steps leading to the formation of free queuine are unknown. The biosynthetic pathway leading to the formation of nucleoside Q in tRNAs in E. coli differs from that in animal systems. As noted in Table I, queuine is not a substrate for the E. coli TRGT enzyme. A precursor of queuine, 7-(aminomethyl)-7-deazaguanine,appears to be utilized by the E. coli enzyme. Support for this mechanism is obtained from the observation that 7-(aminomethyl)-7-deazaguanine has been isolated from acid extracts of E. coli (52).The biosynthetic steps leading to the formation of Q in tRNA following the incorporation of 7-(aminomethyl)-7-deazaguanine are unknown, as are the reactions leading to the attachment of the sugar moieties of Q (Fig. 4,d). It is worthy of note that the E. coli enzyme has been used as a reagent to

582

LARRY KLINE AND DIETER SOLL

compare the levels of Q-containing tRNAs in normal and tumor cells (55). Queuine, the precursor of Q in tRNA in animal cells, has also been suggested to be a possible essential dietary factor (54). Further studies on the biosynthesis of queuine and its precursors are required to evaluate fully the different mechanisms that apparently exist for the formation of Q-containing tRNAs in animal and bacterial cells. It will also be interesting to determine whether other modified nucleosides present in RNA are formed by similar transglycosylase reactions. VI.

Conclusion

The enzymology of RNA nucleotide modification continues to provide a great deal of interest and challenge to the biochemical researcher. The lack of availability of tRNA substrates that can be used to detect and purify the modification enzymes continues to be a major problem. The isolation of mutants in the tRNA modification enzymes (15, 56-59) offers some promise in this regard, as does the isolation of tRNA precursors. Our knowledge of the details of the enzymology of RNA modification is extremely limited, relative to the number and variety of modified nucleosides that have been identified. In addition, the presence of modified nucleosides in the tRNAs of mitochondria and chloroplasts should also be noted. Whether the enzymes responsible for organelle RNA modification are unique or are the same as their cytoplasmic counterparts is an open question.

55. Okada, N . , Shindo-Okada, N . , Sato, S . , Itoh, Y. H . , Oda, K . , and Nishimura, S. (1978). P N A S 75, 4247. 56. Bruni, C. B., Colantuoni, V., Sbordone, L., Cortese, R., and Blasi, F. (1977). 1. Bucieriol. 130, 4.

57. Eisenberg, S. P., Yarus, M., and Soll, L. (1979). J M B 135, 1 1 1. 58. Laten, H . , Gorman, J . , and Bock, R. M. (1978). Nucleic Acids Res. 4, 4329. 59. Janner, F.. Vogeli, G., and Fluri, R. (1980). J M B 139, 207.

A u thor Index Numbers in parentheses are reference numbers and indicate that an author’s work is referred to although the name is not cited in the text.

AB, G., 416, 423(555) Abelson, J. N., 32, 470, 487, 489, 496(16) Abraham, A.. 223 Abraham, G., 246, 262(13) Ahrams, A,, 520 Abrams, R., 185. 218, 230(20), 235(1) Abramson, R., 131(93), 132 Abrell, J. W.. 572, 573(21) Abrosimova-Amelyanchik, N. M., 449, 452(70), 465, 467(70) Acharya, A. S . , 335, 336(109), 337(110) Achberger, E., 79, 80(132) Ackerrnann, S.. 244 Acs, G., 261. 423, 497 Adams, A,. 203 Adams, B., 259 Adams. J. M., 513 Adams, P., 395 Adelstein. S. J.. 342 Adhya, S., 63(28), 64,80(28), 83(28), 86(28), 100, 101, 102167) Adler. J., 83 Adman, R., 83 Adolf, G., 240, 241(76) Agarwal, K . L., 28, 34, 35(21) Agris, P. F.. 557. 561, 568, 572(4), 575(4), 577, 578(4) Ahmed. A . K.. 391(406), 392. 413(405) Aiba, H . , 81, 82(142) Akaboshi, E., 475. 476(20) Aktipis, S.. 343. 344(168) Alazard, R. J., 8 Albanesi, D., 413, 414(520) 583

Alberts, B. M., 65, 176, 177(87), 178(100, 101) Albrecht, H. P., 381 Alderfer, J. L., 545 Aldrich, C., 48 Alexander, M., 185 Alford, B. L., 53, 54(75), 57(75) Allaudeen, H. S., 571 Allen, F. W., 428 Allen, L. C., 432 Allende, J. E., 354 Allewell, N. M., 325, 364, 365 Allerhand, A., 330, 335(75), 374, 37375) Almendinger, R., 238 Altman, S., 213,470,471,472(6).473(2,3,9, 10). 474(2, 9-13), 475(2. 13, 15. 16). 476(13, 20). 479(13, IS), 481. 568 Alzner-Deward, B , , 57 Arnaldi, F., 568 Arnarnath, V., 540, 541(191) Arnemiya. K.. 67 Arnes, B. N.. 570, 571(15), 580(15), 582(15) Amrnon, H. L., 345 Anderson, D. G., 450 Anderson, C. W., 494 Anderson, J. A,, 112 Anderson, S., 179 Anderson, S . F., 214 Andenon. W. F., 34, 35(20), 42(20) Ando, T.. 284, 285(27), 442, 446(40) Andoh, T., 502 Andreatta, R., 335, 347(106), 350( 106), 35 I( 106)

584 Andrews, N . , 101 Andrews, S. J., 135 Andria, G., 359, 421(263) Anfinsen, C. B., 338, 356, 358, 385, 391, 393, 394, 395(361), 396(251), 397, 425 Angel, L., 465, 466(133), 467(133), 468(133) Anraku, N., 4, 22 Anraku, Y.,8, 9(30), 10(30), 12, 13(30, 4 9 , 502, 503 Anthony, D. D., 191, 196(50), 197(50). 207(50) Antoniades, D., 220, 221(9), 222(9), 238(9) Antonoglou, O., 220, 221(9), 222(9), 238(9) Antonov, I. V., 329, 33372, 73), 364(73), 376(72), 377(73), 381, 382(72). 406(73), 415(73) Apirion, D., 476,483(24), 489,511,512,525, 537, 538(69, 153, 159) Arai, K.-I., 160, 161(22), 162(22,23, 36-38), 1651341, 170(22,43), 171(22,23,43), 278 Arai, N., 170, 171 Arata, Y.,444, 445(50), 457, 463(50, 92) Arena, F., 570, 571(14), 572(18) Arendes, J., 240 Arian, S., 343 Ariga, H., 181, 182(153) Arima, T., 465 Armstrong, 577 Arnold, H. H., 558, 559, 563(12), 564(9), 565(9) Arnold, J., 80 Arus, C., 323 Aschhoff, H. J., 559, 563(12) Atchinson, R., 471 Atkins, J. F., 494 Atsuya, I., 131 Attardi, G., 568 August, J., 68 August, J . T., 268,269(15), 270(15), 271(14), 272(14), 273(14), 275, 276(15), 279(50) August, T., 227 AuguSt-Tocco, G., 433 Aujean, 0..286 Auld, D. S., 131, 132, 320, 323(23), 423(23) Augenstein, L.-G., 343 Averyhart, V. M., 562, 563(21) Aviv, H., 547, 548(268) Avramova, Z., 237, 239(63) Avramova, Z. V., 429 Avron, M.,547

AUTHOR INDEX Axelrod, B.,331 Axelrod, V. D., 202, 465 Babich, A,, 248, 262(16), 496 Babinet, C., 525 Bacheler, L., 179 Bachmann, B. J., 23, 489 Bachner, L., 546 Backrnan, K., 20 Baeova, M., 465 Baer, M., 471, 472(9a) Baer, M. F., 473 Baev, A. A., 559 Baez, J. A., 34, 35(22), 32(22), 38, 40(22, 33). 56(33) Baglioni, C., 259, 284, 285(22, 25). 286, 287(22), 290, 293, 298, 299(56), 300(56), 301(46), 302(46), 303(46), 304(46, 66). 306(66), 310(22, 60, 61). 311 Bahl, C. P., 545 Bailey, J., 90, 100 Baldi, M. I., 147 Baldwin, R. L., 341, 352, 354, 355, 385, 386(353), 390(154, 399), 391(243-246). 397 Baldy, M. W.,22 Ball, A., 284, 285(20), 286(20), 296(20) Ball, L. A . , 282, 284, 285(24), 287, 289(34, 39). 291(24), 292(24), 294(34. 39). 295, 296(24), 297(24), 300(39), 301(39), 302(39, 40, 5 8 ) , 303(40), 304(39, 40), 306(39, 58). 307(58, 73), 310(58, 73) Baltimore, D., 88, 1037). 259, 551 Bandyopadhyay, P., 93(37), 94, 104(37) Banerjee, A., 246, 258(1), 260, 262(13) Banejee, D. K., 311 Bannejee, A. K., 268, 273 Barany, G., 360 Barbehenn, E. K., 521, 522(44), 523(44), 524(44) Bardbn, A., 418 Bardos, T. J . , 542 Barbosa, E., 248, 254(23) Baril, B., 182 Baril, E. F., 182 Bar-Joseph, M., 546, 548(263), 549(263), 552(263) Barnard, E. A., 320, 375(17), 398, 407(442), 411(441), 432 Baroudy, B., 263

AUTHOR INDEX Barrell, B. G., 165 Barrett, C.. 99 Barrio, J. R . , 38, 40(36), 53(36), 540 Barrio, M. G.. 38, 40(36). 53(36) Barry, E. J., 326 Barry, J., 176, 177(87) Bar-Shavit, R.. 179 Bartholeyns, J . , 327, 328. 414, 418, 425, 433(58) Bartkowiak, S., 184. 199(13) Bartz, J . K., 575, 576(30, 32) 577(32) Baskin, F., 241 Bassett, C. L.. 538 Basu, S . . 100, 101(67), 102(67) Batey. I. L., 545 Battaner, E., 498 Baudhuin, P., 418, 425, 433 Bauer, S., 323, 426(30) Bauer. W., 139 Bautz, E., 88, 90(9). 93, 96, 98(9), 101(9), 103(35, 44), 105(35, 44, 70). 106 Bautz, E. K. F., 6, 62, 71(8), 74(8). 75(8), 77(8), 111(17), 112, 113(17), 123 Bautz, F., 88, 90(9), 98(9), IOl(9) Bayer, A. A., 202 Bayley, H.. 345, 388(188) Baynes, J. W., 328 Beacham, J., 443 Beard, P., 18 Beaudreau, G., 268 Beaven. G. B., 340, 341(147), 384(147) Beck, E., 168 Becker, A., 4, 12(13), 15(13), 17(13). 27(12, 13), 51 Becker, R. R., 318, 407, 408(488), 410(488) Becker, W. M., 110, 137(6) Beckmann, J. S., 32 Bedows. E., 17, 18(75), 32, 42(3) Beers, R. F.. Jr., 414, 518, 519, 5% 529, 530(100, 101) Behrendt, E., 76, 77(112), 82(112), 84(112) Beier, H.. 95, 96(41), 97(41), 98, 99(51), 100(5 I ) Beintema, J. J., 319, 325, 330, 335(80), 339(80), 340. 345, 346(80), 358, 363, 367(80), 369(42). 370(80), 371(80), 372(80), 372(80), 376(42), 377(42, 80), 378(80), 386(80), 389(80), 395(80), 398(462-4651, 399(9, 10, 139, 443, 444, 448, 449, 454, 455). 400(7, 9, 10, 139,

443, 444, 445-451, 454, 455, 456, 458460, 461a, 462-465). 403(10, 445, 456, 472),404(9, 10. 139, 190,454,455,465), 4039. 10, 443-448, 465). 406(42), 407(450), 409(10, 139, 446, 449, 450, 451, 456, 463), 410(486), 415, 421(443, 445, 446, 454), 429(258) Belagaje, R., 28 Beletskaya. 0. P., 467 Bell, G. I., 119, 126(51), 128, 135, 136(112) Bell, L., 231, 232(42) Bellemore, G., 515 Bellet, A. J . D., 180 Bellmann, B., 432 Bello, J . , 324, 325(38, 391, 33 I , 334(38, 39) Beltchev, B., 536 Bendori, R . , 308 Benedetti, P., 147 Bhicourt, C., 200 Benisek, W. F., 331 Benjamini, M., 343 Benkovic, S. J., 38, 39(37) Bennett, G. N., 533, 537, 544(128, 150) Bennetzen, J., 139, 143(150), 144(150) Benoist, C., 150 Benson, R. W., 131 Benvin, S . , 290, 293, 299(56), 300(56), 301(46), 302(46), 303(46), 304(46, 66), 306(66) Benz, E., 156 Benz, E. W., Jr., 156, 157(11), 158(11), 159(11), 160(11), 161(15), 162(39), 16315. 39). 166(11, IS, 39), 167(39) Benz, F. W., 338, 369, 370, 386(312), 387(313), 389(312, 313), 395(312), 396(312, 427). 431, 432(647) Berg, D., 74, 75(92) Berg, P.. 62, 83, 86, 180, 185, 188(29), 211(29), 212(29), 223 Berger, A., 349, 350, 351, 357 Berger, H., 25 Berger. S. L., 427 Berges. J.. 524 Bergman, J . , 259 Berissi, H . , 301, 302(65), 304(65), 305(65), 306(65), 30/(65) Bernardi, A., 203 Berns, A. J. M., 417 Bernstein, H., 22 Berry, S., 260

586 Bertazzoni, U., I8 Bertsch, L., 107 Bertsch, L. L., 170 Best, A. N., 185, 186(36), 187(36), 193(36), 194(36), 196, 197(361, 210(67) Beychok, S., 81, 130, 384, 421 Beyer, R., 227, 232(31) Bezborodova, S . I . , 449, 452(70), 465, 467(70, 141) Biaka, E., 540 Bibilashvili, R. Sh., 528 Biddlecome, S . , 455 Bigelow, C. C., 395, 396(425) Bikoff, E. K., 471, 482(7), 483(7) Billeter, M. A., 273 Billiau, A., 312 Billups, C., 382, 384 Biondi, L., 329 Birenbaum, M . , 502, 511(17) Birge, C. H., 487, 496(17), 498 Birkenmeier, C. S., 427 Birnie, G., 244 Birsdall. N. J., 292,301(49), 302(49), 306(49) Bishaeye, S., 490 Bishay, E. S.. 417, 418(570) Bishop, D. H. L., 273 Bishop, J. M., 184 Bishop, J. O., 498 Bishop, R., 214 Bishop, R. J., 48 Bison, O., I48 Bittner, M., 176 Bjork, G. R., 566 Blackburn, B. J., 278 Blackburn, G. M., 380, 381(329) Blackburn, P., 326, 328, 334, 3 3 3 4 3 , 406(45), 418(575), 419, 420(574, 575). 421(45, 47), 422(45, 47). 423, 424, 425, 426, 433(57) Blair, D. G. R., 114 Blake, R. D., 16 Blasi, F., 582 Blatti, S. P., 111, 113(9), 116(9), 120(9), I23(9) Blobel, G., 361, 397(273), 424 Bloemendal, H., 417,418,419,423(558), 424 Bloemhoff, W., 348(217), 350, 356 Blomberg, F., 375 Blum, A. D., 355, 391(243) Blumenthal, T., 66,7357). 268,269,270( 13,

AUTHOR INDEX 21, 24). 271(23), 272(27. 28, 31), 273(24, 29, 30), 274, 275(18), 276(39), 277(13, 24, 27), 278(23, 30, 62) Bobst, A. M., 541, 542, 545(200) Bobst, E. V., 542 Boccu, E., 338, 388(127) Bock, R. M., 551, 581(58), 582 Bode, J., 380 Bode, V. C., 4 Boezi, J., 67,88,93(10), 98(10), 102(10), 231 Bogehagen, D. A., 119 Bogenhagen, D. F., 148 Bogorad, L., 117, I19(46), 123(46), 124(46), 151, 152 Boguski, M., 219 Boguslawski, G., 227 Bolen, D. W., 386, 387(373) Bolle, A., 21, 22, 176 Bollum, F., 221, 222(12). 223(12), 224(12), 225(12), 226( 12). 227( 12), 229( 12), 230( 12), 23 1(12), 233( 121, 236( 12), 237(12), 240(55), 243(12, 18) Bollum, F. J., 16, 182 Bolscher, B. G. J. M., 330,335(80), 339(80), 346(80), 367(80), 370(80), 371(80), 373(80), 377180). 378(80), 386(80), 389(80), 395(80) Bon, S., 528, 533(99), 534(99), 545(99) Boni, R., 352 Bonner, J., 68 Bonnet, J., 185, 186(38), 187(38), 188(38), 189(38), 190(38), 194(38), 197(38), 200(38), 202(38), 207(38) Bont, W. S., 423, 424 Boone, R., 246, 261(10), 262(10) Boothroyd, J., 100 Borchardt, R., 263 Borek, E., 557, 559, 566 Borer, P. N., 52 Borin, B., 388 Bonn, G., 335, 346, 347, 350(104), 351(104), 356 Borisov, V. V., 329, 335(73), 364(73), 377(73), 406(73), 415(73) Borisova, S . N., 323, 329, 335(73), 364(73), 406(73), 415(73) Borsch, C. J . H.. 51 Borsett, L. M., 135 Bosch, L., 271, 273(29) Both, G., 258

AUTHOR INDEX Bothner-By, A. A., 350, 355(221) Bothwell, A. L. M., 473, 474(13), 475(13), 476(13), 479(13) Botstein. D.. 9 Bouche, J.-P., 156, 158 Bouhnik, J., 184, 2l4( 18) Bouloy, M., 259, 260(54), 265(56) Bovey, F. A., 330, 335(78), 346(78), 349, 367(78), 368(78. 214), 370(78, 214). 373(78), 374(214). 390 Bowles, M. G., 47 Bowman, C. M.. 308 Bowman, E. J., 473, 474(1 I , 12). 475( 16) Boyce, R. P . . 4 Boy de la Tour, E., 21, 176 Boyer, H. M., 47 Boyer, H. W., 16. 20(67), 21(67), 28 Boyer, P. D., 538 Bradbury, J. H.. 320, 324(18, 19). 327, 366, 367(298, 299). 369(18), 370(298, 299), 372, 375(18, 19), 376(18), 423(19) Bram, R. J., 486, 491(11), 493(11), 494, 4931 I ) Brandts, J. F., 382, 390 Branno, M., 414 Braun, R., 111(18), 112, 113(18), 116(18), 120, 123(18, 5 5 ) Brawerman, G., 227, 231, 232(46), 236, 238(59) Breant, B., 137 Brennan, C. A , , 46 Brennan, M., 390 Brenneman, F. N., 525, 529, 539(106) Brentnall, H. J., 540, 546(177) Bresler, A,, 62 Breter, H., 227, 232(31) Brewer, E. N., 423 Briat, J. F., 152 Brimacombe, R., 506 Brimacornbe, R. L.. 547 Brishammar, S., 520 Brodeur, R.. 242 Brodner, 0. G., 133 Brody, S., 418 Broeze, R. J., 288 Bromley, P. A., 496, 514 Broom, A. D., 540, 541(191) Brosins, J., 506 Brown, D. D., 148, 149 Brown, E. L., 28

587 Brown, G. E., 304, 306. 307(86) Brown. G. M., 561 Brown, J . E., 356 Brown, J. R., 338 Brown, L. R., 320. 324(18, 19). 369(18), 37318, 19), 376(18), 423(19) Brown, R. E., 282, 283, 284, 285(12, 17), 286(17), 287(17), 291(7, 1 3 , 292(12, 13, 15). 293(52, 53). 293131, 296(52, 53), 297(53), 298(53), 301(49), 302(49), 30352, 53, 54). 306(49, 52, 53), 307(52, 53), 312(52, 53) Brown, R. K., 363 Brown, R. S . . 464 Brown, S . , 269, 270(24), 272, 273(24), 277(24), 278(62) Brown, W. E., 320, 323(24) Browne, D. T., 421 Brownlee, G. G., 228, 463, 464 Brownell, J., 540, 541(191) Bruce, A. G., 41, 57(42), 58(42) Bruce, S. A., 6, 9(26), 47 Brugge, J., 180 Bruice, T. C., 341 Brunel, C., 498, 499(69) Bruni, C. B., 582 Brunovski, I . , 89 Brutlag, D., 17 Bryant, F. R., 38, 39(37) Bryant, J., 572, 573(22), 574(22) BUC,H., 106 Buchanan, J. M., 525 Buchi, H., 28 Buchowicz, J., 136 Buchwalder, A., 150 Buhler, J . M., 119, 126, 128, 134(79), 135, 136(13), 137 Bujard, H., 63(30), 64, 83(30), 86(30) Bull, P., 133, 134(99) Bullock, M. L., 8, 23, 24(108), 25(108) Burgero, P. M. J., 198 Burgers, P. M. J., 532, 542(124) Burgess, A. B., 120 Burgess, A. W., 344, 345(184), 385, 387, 388(184), 395, 396(381, 422), 406 Burgess, R. R., 62, 63(7), 64,65(7), 66(43), 67(7, 56),68(7), 70(8), 73, 74(8), 75(7, 8, 561, 76(7), 77(7, 8, 56, 931, 81(7, 93), 82(7), 94, 111, 114, 115(35), 116(8), 117(8), 120, 121(61), 122, 124(61), 130,

AUTHOR INDEX 132(86), 134, 138, 139(133), 141(32, 133), 142(133), 143(133, 159). 144(133) Burghouts, J. Th. M., 424 Burke, R. L., 176, 177(87), 178(101) Burma, D. P., 62,490, 503,504(24), 505(24), 506 Burny, A . , 550 Burstein, E. A , , 467 Burstein, Y.,338, 339(130), 340, 344(130) Burtis, K., 76 Burton, L. E., 419 Burton, Z., 74, 77(93). 81(93), 94 Busch, H., 570 Biisen, W., 499 Bush, J., 62, 66(12), 70(12), 72(12), 73(12), 77(12), 105 Buss, W. C., 114 Bustin, M., 426 Busto, P., 200 Butel, J., 180 Butler, E., 88, 91(12), 93(12, 2 3 , 95(12, 25). 96(25), 100, 101(12), 102(12, 25) Butterworth, P. H. W., 140 Buzash-Pollert, E., 495 Byrnes, R. J., 417, 418 Bystrov, V. F., 329, 33371). 369(71), 379(71) Cabrer, B., 282, 284, 285(18), 304(18), 306, 307(86) Caldarea, C., 242 Caldi, D. G., 357, 432(255) Camble, R., 443 Cameron, J. R., 8 Cameron, V., 37, 38(32), 42(32), 51, 55(32, 68) Cammack, K. A . , 505 Campagnori, F., 18 Campbell, I. D., 372, 373(316a) Campbell, J . L., 494 Campbell, L., 25 Campbell, M. K., 445, 449, 450(73), 452 Campino, C., 131 Canaani, D., 259 Canellakis, E. S., 184, 185(5), 188(26) Cannistrado, V. J.. 494 Cano, A , , 147 Cantatore, P., 239 Canter, V . M., 433 Cantoni, G. L., 203

Cantor, C. R., 320, 322(16a), 532 Canuel, L. L., 330, 335(78), 346(78), 349, 367(78), 368(78, 214), 370(78, 214), 373(78), 374(214) Capasso, S., 413 Capesius, I., 520 Capon, D., 162, 165(40), 166(40) Carbon, J. A., 202, 213 Carey, N. H., 550 Carlisle, C. H., 337, 365, 373(119) Carlson, W. D., 364 Carmichael, G. G., 268, 270( 12), 272, 275(18), 276 Carrara, G., 147 Carre, D. S., 185, 186(33), 187(33, 49), 191, 192(53), 193(53), 194(33, 49). 195(53), 196(49), 197(33), 200(33), 201, 202, 2 I O(49) Carriquiry, E.. 200 Carroll, W. R., 425 Carsana, A., 415, 429(540) Carson, F. M., 51 Carter, A., 92, 98(29), 99, lOO(29, 58), 105(29) Carter, J. R., 338 Cartese, R., 215 Carty, R. P., 325, 331, 332(86), 375(41), 423 Carusi, E. A., 180 Caruthers, M. H., 28 Cashel, M., 537, 538(158) Casoli, C., 107 Cass, K. H., 132 Cassani, G., 16 Casti, A., 242 Castles, J. J., 502 Castroviejo, M., 200 Cathala, G., 498, 499(68) Cech, C., 69 Celotti, L.. 338, 388(127) Celantano, J., 57 Center, M. S., 27 Cernosek, S. F., 443 Cerny, G., 503 Cerutti, P., 202 Cha, C.-Y., 339 Chaiken, I. M . , 345, 347, 351, 355(208), 365 Chakrabarty, A. K., 364 Chakraborty, P. R., 90,93(24,37), 94,98(24, 361, 101(24), 103, 104(37, 71) Chakraburtty, K., 503, 504(24), 505(24)

AUTHOR INDEX Challberg, M. D., 180, 181, 182(150) Chamberlin, M. J.. 62, 63(25), 64, 65(11), 66(11, 12),67(11,59),69(14,66),70(12), 71(1 I , 66,77), 73(1 I , 12, 65, 66, 77), 74, 75(66, 74. 92), 77(12), 78(11), 79(14), 80(11, 41), 82(66), 83(14, 25), 85(14), 86(14, 15,79,90), 88,89,90,91(8), 92(8, 26, 281, 93(25), 94(8, 32), 95(25), 96(25), 98(8, 261, 99(8, 30, 49, 50, 51). lOO(30, 49, 50, 51), lOl(49, 50), 102(25, 32), 103(26, 28, 32), 104(26, 28, 301, 105(26, 50), 108(76), 113, 223 Chambon, P., 110, 111(27), 112, 113(12,27), I17(27), 119(45), 120(45), 122(45), 123(27, 45). 124(45). 126(45), 127(45), 137(131), 138, 139(132), 142(140), 143(140), 144(148), 146, 147, 148, 150 Chambrach, A . . 521, 522(44). 523(44), 524(44) Champoux, J . J 176 Chan, J. S., 397 Chandler, D. W.. 139, 140(143), 141(143) Chaney, S. G.. 538 Chang, S., 213 Chang. S. H., IS. 16(62), 28(621, 561 Chantrenne, H., 243, 397. 539, 548(171), 549( I7 I ), 550( 17 I ) Chapeville, F.. 185, 186(33), 187(33. 49), 191, 192(53), 193(53), 194(33. 49), 195(53), l96(49), 197(33), 200(33), 201, 210(49), 536 Chapman, B. E., 366 Chaplinski, T., 425 Chatterjee, B . , 548, 549(279) Chattoraj. D., 179 Chavez, L. G., Jr., 362, 363, 389, 393(388), 394(388), 395, 396(388) Chelbi-Alix. M. K., 521. 527(51) Chen, A. K., 383 Chen, D., 520 Chen, M. C., 386 Chen, P.. 32. 34(8). 48(8), 49(8) Chen. S., 263 Chen. Y.. 34 Cheng, H. N . , 390 Chen-Kiang, S., 248, 262 15) Chernajousky, Y.. 287, 298. 302, 304(59), 30359. 751, 307(59), 308(59) Chertov, 0.. 74 Chetvin, I. I., 329, 335(71), 369(71), 379(71)

.

589 Chesterton, C. J., 119. 123, 140 Chestier, A . . 139, 143(144) Chestuktin. A. V., 9 Chevalley, R., 21, 176 Chheda, G . , 577 Chiancone, E., 412 Chien, J. R., 12. 13(45) Chinault, A. C., 551 Chirgwin. J . M., 361, 427 Choate, W. L., 425 Choder, M., 548 Chodroff, S., 546 Choi, Y . C., 570 Chou, J.. 180 Chou, J. Y., 526, 530(82). 531(86), 532(114), 533032, 86), 534, 53562, 114), 536(82. I14), 537(86), 545(131), 547(82) Chou, P. Y., 406 Chow, L. T . , 181 Christman. J . K., 423 Chuguev, 1. I., 202 Chung, C. W., 184 Churchick, J. E., 384 Ciampi. S.. 570. 571(14), 572(18) Ciarrocchi, G., 10 Ciliberto, G., 570, 571(14) Cina, J., 134 Cinader, B., 362 Clark, S., 89, 105, 106(17) Clayton, D., 179 Cleary, P., 80 Cleland, W. J., 234 Cleland, W. W., 198, 338 Clemens, M. J . , 282, 283, 291(7), 295, 304(14, 57) Cleuter, Y., 243, 550 Cleveland, D. W., 124 Cochet-Meilhac, M., 146 Cohen, J. S . , 330, 335(79), 339. 346(79), 347(203), 354, 355, 366, 367(297), 368(203), 369(238, 297). 370(79. 297). 372(135),. 373(79, 237, 239). 374(203a), 376, 377(305), 379(239, 305). 380(305), 381(305, 329), 382(305), 386, 388(239), 396(375), 418(79), 462 Cohen, L., 506, 512(42), 538 Cohen, L. A., 347, 355(208), 365 Cohen, M. S., 407 Cohen. S. S., 195, 502, 528, 538 Cohn, M., 531, 533(116), 534(116), 535

590 Cohn, W. E., 412, 540 Colantuoni, V., 582 Colby, C., 284, 285(20), 286(20), 296(20) Cole, C., 180 Cole, P., 275 Coleman, J., 90, 91, 96, 100, 103(21, 22, 27), 486 Coleman, M. S., 233, 240(55) Colonns, R.. 259 Condit, R. 98. 99(52), 104(52) Conley, M. P., 32, 34(8), 48(8), 49(8, 58) Conlon, S., 89, 90(19), 94(19), 95(19) Connett, P. H., 532, 535(122) Conrad, S. E., 494 Content, J., 286, 311(29), 312 Contreras, R., 150, 483 Cook, K. H., 391 Cooley, W., 50, 51(64, 65) Cooper, J., 260, 265(58) Cooper, M. R . , 390, 391(396) Corcoran, C., 339 Corden, J., 148, 150 Cornelis, P., 547, 548(264), 549(264), 550(264), 551(264). 552(264) Corte, G., 502, 511(12) Cortese, R., 475, 570, 571(14, IS), .572(18), , 580(15), 582(15) Corti, A , , 242 Cory, J . G., 191, 197(51), 207(5l) Cory, S . , 513 Coulter, D. E., 135 Coulter, M., 83 Coupar, B. E. H., 119, 123 Court, D., 63(29), 64, 80(29), 83(29), 86(29) Court, M., 98, 100(48), 101(48), 102(48), 105(48) Courtois, Y., 448, 457(69) Courvalin, J. G., 146 Coutsogeorgopoulus, C., 184 Cowgill, R. W., 341, 342, 384 Cowles, J. R., 519 Coy, G., 68 Cozzarelli, N . R . , 4, 5(14), 9, 16, 17(19), 20(67), 21(67), 28(19), 32, 33, 34(8), 35(21), 36(18), 37(5), 38(5, 16). 39(5), 40(5), 41(5, 3 3 , 42(5), 47(18), 48(8), 49(8), 50(63). 51(63) Cozzone, P. J., 429 Crain, P. F.. 447, 580, 581(54). 582(54) Craine, J. E., 521, 522(44), 523(44), 524(44). 532

AUTHOR INDEX Cramer, F., 184, 185. 186(34), 187(34), 189(34), 190(34, 541, 191(34), 192(54, 5 3 , 194(54), 196, 197(54), 200, 213, 215(3), 464 Cranston, J. W.. 32, 34(12), 36(12). 37(12). 40(12), 41(12), 42(12), 47(12) Crawford, L., 180 Crawford, N., 506 Creighton, T . E., 338, 385, 392(129), 397 Cremer, K., 525 Crerar, M. M., 135 Crescenzi, V., 346, 388 Crestfield, A . M., 324, 330, 405(35), 413, 42 I , 428 Criddle, R. S . , 150, 151(213) Crippa, M., 138 Croissant, O., 139, 143(142) Crompton, M. W., 366, 367(298), 370(298) Crook, E. M., 418, 425 Crothers, D. M., 542 Crouch, R. J., 489,490(24),496,498,499(70) Cuchillo, C. M., 323 Cudny, H., 184, 185(41). 186, 187(41), 189(41), 190(41), 192, 193(56), i94(56), 195(56), 199(13). 215. 480. 481(31), 482(3 I ) Culotti, J . , 26 Dadok, J., 350, 355(221) Dahlberg, J. E., ’273, 476, 494, 495(37) Dahmus, M. E., 136, 137(119b) D’Alessio, G., 385, 41 I , 412, 413(348, 517), 414(348. 500, 517, 520), 433 D’Alessio, J. M., 1 1 I(l5). 112, 1 l3(l3, 14, 15). 116(13, 14, 151, 117(13, 14, 1 3 , l19(13-15), 123(14), 128, 129(82) Daley, K., 84 Damodoran, N. P., 381 Danchin, A , , 536 Daniel, V . , 185, 195(30), 21 1(30), 212(30), 535 Danna, K. J., 179 Darlix, J. L., 487, 496, 514 Darnell, J. E., 227, 231, 235, 236(69), 238, 239, 248, 262(15, 16), 498 Darvey, I. G., 428 Das, M. K., 336 Dasgupta, A . , 551 Datta, A . K., 502 Dauguet, C., 139, 143(145) Daumigen, M., 360

AUTHOR INDEX David, E. S., 135 David, M., 52 Davidson, N . , 494. 553 Davies, G. E., 426 Davies, K. E., 128 Davies, M., 176 Davies, P. L., 464 Davis. N . L., 496 Davis. R. W., 8, 9, 15, 27(61) Davison, B., 65, 72 Dawid, I. B.. 150, 151(218) Dayhoff, M. O., 403 Deakyne, C. A,, 432 Dearborn, D. G.. 320. 324(20), 376(20, 34, 37) Debacker, M., 67 de Boer, H., 72 Debov, S. S., 520 De Clercq, E., 312, 540, 546 DeCoen. J. L., 397 decrombrugghe, B., 80 deDuve, C., 237 DeGorgi, C., 239 deHaseth, P., 56 deHaseth. P. L., 134. 276 Dekker, C. A,, 465, 466(128) Delaage, M., 341 de Lamirande, G.. 417 Del-Campillo-Campbell, A , . 535 Delk, A. S., 558, 564, 565(31, 32) Delorbe, W. J., 116, 117(42), 126, 139(73), 143(73) De Lorenzo, F., 397 Delvig, A. A., 520 deMartynoff, G., 423 Demma, G., 412 Dengler. B., 52 Denhardt, G. H., 21, 176 Dennis, E. A,. 462 Dennis, P. P., 494 Deonier, R. C., 494 DePamphilis, M. L., 179 Depew, R. E., 49, 50(63), 51(63) DePrisco, R., 411, 414(500) Derbyshire, R., 150 De Robertis. E. M., 215 deSante, D., 222, 229(15) Deshpande, A. K., 548, 549(279) Desiderio, S. V., 181. I82(150) Desmyter, J., 312 De Somer, P., 312

59 1 Desreux, V., 416 Desrosier, R., 231 Deugau, K. V., 14, 20(55), 21(55) Deutscher, M., 511 Deutscher, M. P., 183, 184, 185, 186(37), 187(37, 45, 46), 188(17, 37, 45, 46), 189(37), 190(37. 4 3 , 191(37), 192, 193(43, 44), 194(45, 61, 70), 19344, 48, 59). 196(45,61). 197(61,70), 198(43.60), 199(45, 60, 61), 200(61, 63, 64), 201(37, 60. 76), 202(46, 60, 63), 203, 204, 205(44), 206, 207(44, 45, 48, 60, 63, 64, 70,96), 208(61,70), 209(69), 210(45, 61, 63, 64, 691, 21 l(60, 69, 70), 212(44), 213(2, IOO), 214(17, 48), 215(99, I O I ) , 479480(30), 481(30, 31), 482(30, 31, 33), 537, 538(157), 539(157) Devos, R . , 243 De VrieLe. G.. 398, 400(458) DeWaard, A,, 176 Dezelee, S., 117, 120(47), 123(47), 126, 127(47), 138, 140 Di Bello, C . , 345, 347, 353208). 365 Dickerson. R. E., 20, 403 Dickson, B., 135 Dickson, E., 259,464,475,486,494(8), 496, 499 Dickson, R. C., 48, 49(58) Di Donato, A,. 412. 414 Dieckrnann. M., 185, 188(29), 21 1(29), 212(29) Diener, T. O., 553 Dietz, F. W., Jr., 526, 530(75) Diez, J., 236, 238(59) Dijkstra, J., 416. 423(555) Dijkstra, K., 366, 367(301, 302) Diopoh, J., 327 Dirheimer, G., 521, 527(51) Diringer. R.. 62 Divizia, M., 286, 311(29), 312 Dixon, G. H.. 464 Dixon, H. B. F., 436, 439 Dobberstein. B.. 361, 397(273) Dobkin, C., 275, 276(55), 279 Dobson, C . M., 372, 373(316a) Dodds, J. R., 139, 143(146) Doel, M. T., 550 Doetsch, P., 302, 303(69) Doi, R., 63, 65, 76(16), 78(16), 79(16), 132 Doi, T.. 44, 54(47), 55(47), 56 Dolganov, G. M., 9

592 Dolgikh, D. A., 365 Domdey, H., 464 Domingo, E., 279 Dondon, L., 526,527(79), 535,540 Donis-Keller, H., 58,464,497 Dorson, J., 221,222(12),223(12), 224(12),

225(12), 226(12), 227(12), 229(12, 1 3 , 230(12), 231(12), 233(12), 236(12), 237(12),243(12) Doscher, M. S ., 346,364 Dose, K . , 240 DoskoEil, J., 415,429(540) Dougherty, J. P.,287,288,289(38), 290(38), 292(38), 294(38), 299(47), 300(38, 47). 301(38, 47), 303(47),304(38), 305(38) Douglass, J., 277 Dowd, S. R.,443 Dower, W.J., 284.285(17),286(17), 287(17) Doyle, B . , 344 Drabarek, S . , 443 Dressler, D., 159,161(15), 162,165(15,4042). 166(15,40). 174 Dreyer, C.. 140,l4l(l56),490 Dreyer, W.J., 450 Drigin, Y.F., 36,37(25) Drocourt, J. L.. 521. 527(50,51). 531,540, 546( I 17) Dubert, J . M., 525 Dubin, D. T., 568 Dubois, M. F., 304 Dubrul, E. F., 579,581(47) Duceman, B. W., 136 Dudkin, S. M.,323, 329, 335(71, 72), 369(71), 376(72), 377(72), 379(71, 72), 381(72),382(72),429 Dudock, B. S . , 558, 564(8) Duerinck, F. R.,185. 188(32), 202(32) Duester, G. L., 111(115), 112. 113(13, 15). 116(13,15). 117(13, 15). 119(13,15) Duffy, J.. 76,78(120),80(120) Dugaiczyk, A., 28 Dugds, H.. 386,396(372) Duguet, M.,170 Duisterwinkel, F. J., 341,342(155),372(155) Dullin, P., 184 Dunn, B. M..347,351,355,365 Dunn, J., 62,71(8), 74(8), 75(8), 77(8), 88, 90(9), 93,98(9), 99,lOO(53). lOl(9)

Dunn. J. J., 10,485,486(1),487(15),489(1,

6,15),490(6,15). 491(5, 12),493(5, 15). 494181,4936).496

AUTHOR INDEX Dunne, F. T., 410 Dupuis, G., 443 Dynan, W.S., 114, 115(35), 141(32), 142,

l43(1 5 9 ~149 Eastlake, A., 393 Eaton, M.A. W., 521,527(49) Ebel, J.-P., 185, 186(38). 187(38), 188(38),

189(38), 190(38), 194(38), 197(38), 201(38), 202(38), 207(38), 210,21 I(%), 21398) Ebisuzaki, K . , 25 Eckstein, F., 198,430,431,(643,644). 461, 532,540,541,542(124), 545 Eddlemann, H., 48 Edelman, G. M.,12,18(49) Edenberg, H.,179 Edgar, R.S . , 21,176 Edmonds, M.,185,218,219(2),221,222(11), 223(2, I I ) , 224(l I ) , 225(2,1 I ) , 226(l I), 227, 228(2), 229(11), 230(11, 20). 231(11), 233(11), 234(11), 235(1), 236(11).237(11). 238(11),243(11) Edy, V. G., 490 Efstratiadis, A., 275,548,552 Eftink, M. R.,385,446 Egami, F., 435,436(2), 439,440(2), 441(2), 442(2), 444(2), 445(2), 447(I), 44"). 449(2), 450(2, 72), 452(4), 457,458(2), 462,464,465(2), 466(2, 127), 467(127), 468(127), 501 Egan, W., 339,372 Egly, J. M., 425 Egrie, J., 237,241(66) Ehrlich, S. D., 20,21(87) Eich, E . F . , 417,423 Eichler, D. C., 465, 466(133). 467(133), 468(133) Eikhom, T. S., 268,273 Eisenberg, S., 170 Eisenberg, S. P., 582 Ekstein, F., 39 Eliasson, E., 178 Eliasson, R..179,547,548(270) Eliceiri, G . , 572,573(22), 574(22) Elkins, B. N.,577,578(40) Elodi, P.. 332 Elson, D., 501, 503(2) Elson, M., 319,394(11),406(11) Elton, H., 559, 563(12) Emelyanenko, V. I . , 467

593

AUTHOR INDEX Emerson, T. R., 426 Emmens, M., 363, 398, 400(451), 403(472), 404,409(45 I ) Engel, J . D., 553 Engbaek, F., 65 Engelhardt, D. C., 579 Engelke, D. R., 148, 149(197) England, T. E., 32, 38, 40(34, 36), 41(40), 42(7, 34), 43(7), 53(34, 36), 57(40) Engle. J. L., 131(93), 132 Englehardt, R., 65 Englesberg, E., 80 Enomoto, T., 181, 182(151) Enrione, M. L., 184 Ensinger, M., 246, 248( 1 I ) < 249, 256 Eoyang, L., 268. 271(14), 272(14), 273(14) Epand, R. M., 355, 366(237), 367(237), 368(237), 369(237), 370(237), 373(237), 379(237), 386(237) Eperon, I. C., 214 Epinatjeff, C., 453, 457, 463(81) Eppstein, D. A., 284, 285(23), 302(23), 304(23), 306(23) Epstein, C. J., 391, 397 Epstein, R. H., 21, 22, 176 Erenrich, E. S., 430. 431(644) Erickson, J., 179 Erickson, P. M., 332, 333(93) Erickson, R. J., 528 Esteban, R. M., 282, 309 Evans, H. H., 112 Evans, J . A., 192, 193, 195(59), 198(60), 199(60), 200, 201(60, 76). 202(60). 207(60), 21 1(60), 214 Everett, G. A,, 532, 535(121) Eylar, E. H., 410 Fabisz-Kijowska, A,, 184 Faerber, P., 540 Fahrney, D., 332 Falaschi, A., 10, 83 Falchuk, K. H.. 131. 132(87) Falco, S., 88, 107( 14) Falcoff, E., 282, 286, 304 Falcoff, R., 282. 304 Falk, P., 386 Fancher, H., 223 Faras, A. J., 131, 132(91), 184, 192 Fareed, G. C., 7, 8(29), 9(29), 15, 16(29,63), 17(29, 63), 18(29), 19(29, 63), 21, 22, 2363). 27(29. 90). 179

Farina, B.. 398, 399(457), 400(457), 41 2(457), 42 I(457) Farkas, W. R., 579, 580(49), 581(45-47.49, 50)

Farrell, P. J., 284, 285(18), 287, 288(38). 289(38), 290(38), 292(38), 294(38). 300(38), 301(38), 304(38, 38). 305(38) Fasman, G. D., 406 Faust, M., 246 Favre, A,, 202 Federman, P., 282. 301, 302(65), 304(65), 305(65), 306(65), 307(65) Federoff, N. V., 268, 270(16) Fedorov. B. A., 365 Feeney, R. E., 320, 323(22), 324 Feeny, J . , 380 Feitelson, J., 343 Feix, G., 16, 227, 231(27), 233(27), 234(27), 235(27), 273, 274(37), 275, 278, 279, 545 Feldberg, R. S . , 18 Feldman, M., 559 Felicioli, R. A., 447 Fellner, P. C., 568 Felsenfeld, G., 415 Ferbus, D., 287, 292(41), 301(41); 302(41), 303(41, 68) Ferrari. S., 475, 499 Fey, G., 180 Fiddes, J. C., 165 Fields, R., 436 Fiers, W., 150, 228, 243 Fiers, W. C., 185, 188(32), 202(32) Fietta, A,, 147 Filipowicz, W., 246, 258(3) Filippi, B., 346 Filira, F.. 329 Findlay, D., 462 Findlay, J . B., 431 Fini, C., 414, 415 Finkelstein, A. V., 385, 386 Finkenstadt, W. R., 373 Finn, F. M., 335, 346, 347(105, 1061, 350(105, 106). 351(106, 201), 352, 355(221). 443 Finn, M., 57 Fire, A., 147, 263 Fischamn, B., 81 Fischer, S. G., 124 Fiser, I., 540 Fisher, R., 66, 75(57) Fitch, W. M., 403

594 Fitt, E. A., 522, 526(57), 527(57), 536(90) Fitt, P. S . , 520, 521, 522(45), 526(45, 57), 527(45,’57), 530(75, 81), 536(90) Fittler, F., 202, 575, 576(31), 577(31) Fiume, L., 146 Flammang, T., 65 Fleischer, G., 548, 549(274) Fleischman, R. A., 27, 28(129) Fleissner, E., 566 Fletcher, P. L. Jr., 465, 466(132), 467(132) Flint, S. J., 140 Flintoff, W., 83 Florentev, V. L., 528, 534(97) Floridi, A., 411, 412, 414(500, 509), 415 Flory, P., 179 Fluri, R., 582 Folayan, J. 0.. 540 Folsom, V., 502 Fontana, A., 350, 351(220), 385, 413(348), 4 14(348) Forlani, L., 412 Foss, K., 480, 482(32) Foster, L. B., 423 Foulds, J., 214 Fox, C. F., 145 Fox, J., 79, 80(129) Frabotta, M., I05 Fraden, A., 287, 308 Fraenkel-Conrat, H., 265, 542 Francke, B., 179 Frank, E., 84 Franze de Fernandez, M. T., 268, 269(15), 270(15), 271(14), 272(14), 273(14), 276( 15) Fraser, A. R., 56 Fraser, N., 464 Freedman, M. H., 355, 375 Freeman, K. B., 491 Frenkel, G. D., 23 Fresco, J. R., 16, 203 Fridkin, M., 533, 544(127) Friedman, H., 364 Friedman, M. E., 339 Friedman, R. M., 282, 288, 311 Fritz, P. J., 423 Froebe, C. L., 332, 333(93) Fromageot, P., 117, 119, 120(47), 123(47), 126, 127(47), 128, 134(79), 135(77), 136(113), 137, 140, 144 Fruchter, R. G., 330, 413

AUTHOR INDEX Fruscoloni, P., 147 Fuchs, S . , 397, 540 Fuhrmann, S. A., 32 Fujii, N., 359, 360(265) Fujii, T., 432 Fujii, Y., 464 Fujioka, H., 340 Fujiyama, K., 56 Fukada, K.,487, 496(16) Fukemi, Y., 276 Fukuda, F., 65 Fukuda, I., 465, 466(135) Fukuda, R., 76. 77(113), 79, 84, 132 Fukui, T., 541, 544 Fukumoto, R., 56 Fulling, R., 436, 444(7) Fung, D. S.,346, 364 Furia, A., 415, 429(540) Furnaux, H . , 108, 248, 249(22), 250(22), 251(22), 252(22), 255(22), 263(22) Furth, J., 244 Furth, J. J., 185 Furuichi, Y., 246, 248(5), 257(5), 258, 259, 261(5), 262(5), 263, 464 Fuse. A., 312 Futai, M., 502 Futterer, R., 181 Gaastra, W., 319, 358, 398(462 4 6 3 , 399(9, 4491, 400(7, 9, 447,449, 458, 461a, 462, 4651, 403 (472), 404(9, 4 6 9 , 405(9, 447, 4 6 3 , 406, 407, 408(440, 462), 409(449, 486). 410(486), 429(258) Gabel, D., 387, 396(381). 405 Gabriel, T. F.,13, 14(54), 16(54), 17(54), 28(54) Gaertner, E., 185, 186(34), 187(34), 189(34), 190(34), 191(34) Gage, L. P., 119, 147, 214 Gagnon, C., 417 Gajda, A. T., 521, 522(45), 526(45). 527 Gallerani, R., 268, 270(10), 272(10) Galli, R., 414 Gallo, R. C., 145, 147(173), 148(173) Galluppi, G. R., 549 Gallwitz, D., 243 Galzigna, L., 352 Galiazzo, G., 337, 344, 356(117) Gambaryan, A. S.,559 Game, J. C., 26

AUTHOR INDEX Ganesan, A. T . , 4, 6(15), 17(15), 18(15) Gangloff, J . , 210. 211(98). 215(98), 521. 527(51) Ganoza, M. C., 56 Ganther, H. E., 339 Gantt. R., 562, 563(22) Garber, R . L., 147. 214, 473, 474(12), 475(15), 479(15) Gardiner, K., 475, 476(18) Garel, J . R . , 327, 339, 341, 388, 390(154), 391(394) Garzillo, A . M., 413, 414(520) Gassen, H . G., 541, 542 Gasteland, R. F., 502, 505, 512, 513(23), 514 Gauss. D. H . , 205, 564 Gavilanes, J. G., 326. 335(45), 406(45), 420, 421(45), 422(45) Gawronski, T . H . , 352, 354, 355 Gayley, P. J., 292, 293(53), 296(53), 297(53), 298(53), 305(53), 306(53), 307(53), 3 I2(53) Geballe, A . P., 9, 34, 36(18), 38, 41(35), 47( 18) Gefter, M. L., 4, 12(13), 15(13), 17(13), 27(12, 13), 147, 148, 160, 170, 263, 471, 482(7), 483(7), 575, 576(33), 577(33) Gegenheimer, P., 476, 483(24) Geidarov, T . G., 323 Geider, K . , 86, 160, 168 Geiduschek, E. P., 22, 76, 78(120), 80(120), 88 Gellert, M . , 4, 5(9), 8, 12, 13(46), 139, 20), 17(20), 18(20, 46), 19(20), 23, 24(108, log), 25(108), 27(9, 20), 160 Gemershausen, J . , 249 Gennari, G., 344 Geoghegan, K. F., 324 Gerber, A . D., 390. 391(396) Gerken, T . A . , 320, 324(20), 376(20, 37) Germond, J . E., 140 Geroch, M., 203 Gershowitz, A.. 220,225(6), 228.230(6), 246, 248, 249(20), 251(20), 256(28), 261( lo), 262(10) Getz, M., 244 Ghelardini, P., 26 Ghiron, C. A , , 385, 446 Ghora, B. K.. 476, 483(24) Ghosh, N., 304, 305(76), 306(76)

595 Ghosh, R. K . , 210, 212, 21399, IOI), 479, 480(30), 481(30), 482(30, 33), 511 Ghosh, S . , 506 Giacherio, D., 180 Giacomoni, P., 103 Giege, R., 202 Gielow. W . , 80 Gilbert, C. S . , 282, 292, 293(52. 53), 296(52, 53), 297(53), 298(53), 299, 301(64), 302(58), 305(52, 53, 54), 306(52, 53, 58), 307(52, 53, 58), 310(58), 312(52, 53) Gilbert, L. I., 111(19), 112, 113(19), 117(19), l23( 19) Gilbert, S . , 72 Gilbert, W., 20. 57, 63(26), 64,76(26), 83(23, 26), 86(23, 26), 98, 100(47), 102(47), 103(47), 134, 464, 497 Gilboa. E., 282, 547, 548(268) Giles, N. H., 538 Gilham, P. T . , 37, 38(30), 39(30), 55(30), 533, 537, 544(128, 150), 545 Gillespie, D., 238, 241(67) Gillis. E., 243 Gillum. A., 179 Gillum, A . M., 463 Gilman, M., 71, 78 Gilman, M. Z., 62, 6311). 66(11), 67(11), 71(11), 73(11), 78(11), 80(11) Gilvarg, C., 222, 243(18) Gindl, H . , 541 Ginsburg, D., 485 Giordano, F., 413 Giormani, V., 329 Girard, M., 168, 168(45) Girgenti, A . J., 191, 197(51), 207(51) Giron, M. L., 2 2 3 2 9 , 226, 233(25) Gissinger, F., 1 1 1 , 113(12), 117, 119(45), 120(45), I22(45), I23(45), 124(45), 126(45), 127(45), 138, 139 Giveon, D., 546, 549(262), 551(262) Givol. D., 397 Gladstone, L., 61 Glassberg. J . , 161 Glazer, R.. 232 Glazier, K., 487, 496(17) Glick, D. M., 320, 375(17) Glick, J. M., 561, 562(19), 563(21) Glickman, R . , 231 Glitz, D. G., 319, 394(11), 406(11), 465, 466( 133). 467(133), 468(133)

596 Glukhov, B. N., 433 Gniazdowski, M., 139 Godefroy, T., 525,526,528,531(85), 533(85, 991, 534(99), 535(85), 537(85), 545(99), 547(85) Godefroy-Colburn, T., 519,520(15), 525(15), 526, 527(79), 528, 530(15), 531(15), 536(15), 537(15) Godson, B. N., 165 Goelz, S., 276 Gold, L., 51 Gold, M. H . , 325, 407 Goldberg, 1. H., 540 Goldberger, R. F., 338, 391, 397 Goldfeder, A . , 150, 151(219) Goldthwait, D. A., 132, 185, 191, 196(50), 197(50), 207(50) Golgher, R. R., 292, 293(52), 296(52), 304, 305(52, 54), 306(52), 307(52), 308(79), 312(52) Golomb, M., 92, 98, 99(30, 49, 50, 51). 100(30,49, 50, 511, lOl(49, 50), 104(30), 105(50)

Gomez-Guillen, M., 213 Gonzalez, R. G., 278 Gonzalez, N.. 66 Goodman, D.,249 Goodman, H . M., 16, 20(67), 21(67), 28, 47, 184, 214, 273 Gordon, J., 259 Gorecki, M.. 319, 320(5), 342 Gorenstein, D. G., 379, 380, 431 Gorinsky, B. A . , 337, 365(119), 373(119) Gorman, J., 581(58), 582 Goto, J., lll(23, 24), 112, 113(23, 241, 116(23, 24). 119(23) Goto, S., 417 Goto, T., 579, 580(53), 581(52) Goto, Y.,393 Gotoh, S., 487, 496(17), 498 Gottesmann, M., 63(28), 64, 80(28), 83(28), 86(28) Gottesman, M. M., 23, 24(109) Goulian, M., 9. 176, 179 Goux, W. J., 384 Gozes, I., 546, 548(263), 549(263), 552(263) Graessmann, A., 180 Graessmann, M., 180 Graffe, M., 525, 532(67), 539(67), 546 Gragerov, A., 76, 77(114)

AUTHOR INDEX Gralla, J., 139, 140(143), 141(143) Grandi, G . , 385, 413(348), 414(348) Gratzer, W. B., 340, 341(147), 384(147) Gray, C. P., 168 Gray, G. R., 328 Greco, L., 398,399(457), 400(457). 412(457), 421(457) Green, R., 50, 51(64) Green, R. W., 496 Greenberg, R . , 558 Greenblatt, J., 74, 80(91) Greene, J. J., 299 Greenleaf, A . L., 111(17), 112, 113(17), 123, I35 Gregoire, R. J., 56 Greif, R. L., 417, 423 Gregory, M. J.. 341 Grez, M., 221 Gribnau, A . A . M., 417, 423(558) Griffin. B. E., 540 Griffin, J. H . , 355, 368, 369(238), 373(238), 377(305), 379(305), 380, 381, 382 Griffith, J., 168, 170 Grinkevich, V., 74 Grishchenko, V. M., 465 Grit, K. L., 343, 375 Groen, G., 319, 363, 398(462), 399(454), 400(454, 459, 462), 403(472), 404(454), 408(462), 421(454) Groner, Y., 259, 498, 499(71), 548 Grosch, J. C., 528 Grosfeld, H . , 535, 536(142), 547(142), 548(142), 549(276) Gross, C., 65, 74, 77(93), 81(93), 94 Gross, H . J., 185, 188(32), 202(32), 464 Grossman, L., 83 Grossweiner, L. I., 344 Gruber, M., 398, 399(443, 444). 400(443, 444), 405(443, 444), 416, 421(443), 423(555) Grueter, F., 205, 564 Grummt, F., 182 Grummt, I., 148, 498, 499(70) Grunberger, D.,464 Grunberger, D.,578, 579 Grunberg-Manago, M., 518,519(8), 520(15), 521, 522(41), 523, 524(38), 525115, 38), 526(41), 5281781, 529(38), 530( 15, 75, 83),531(15,60,84). 532(67,84), 533(99), 116), 534(8, 99, 1161, 5338, 83, 112),

AUTHOR INDEX 536(15, 83, 84), 537(15), 539(3, 671, 540, 542, 545(8, 991, 546, 547(112) Gubanov, V., 74 Gueguen, P., 536 Guerrier-Takada, C., 475, 476(20) Guha, A., 104 Guida, G., 412 Guilfoyle, T. J., I I1(22), 112, 113(22), 116, 120, 122(58), 123(22, 59), 145(58) Guilley, H., 464 Guissani, A., 521, 523(47). 526(47), 427(80), 528(47, 78), 530(47), 531(80), 535(115), 536(115), 546(47), 547( I 15) Gulyaeva, V. I.. 465, 467(141) Gumport, R., 20 Gumport, R. I., 13, 17. 18(75), 32, 33, 34, 35(14, 22), 32(22), 38, 40(6, 22, 33), 41(6). 4 2 0 , 6 , 7), 43(7), 44, 46(6), 47(6), 54(45), 56(6. 33, 41), 544 Gunlap, A., 568 Gupta, G., 332 Gupta, K. C., 116, 135 Gupta, M. N., 337 Gupta, N. K., 15, 16(62), 28(62) Gupta, R. S., 502,506,507(10), 508(10), 509, 512 Gupta, S. L., 282,284,285(20), 286(20), 287, 289(36), 296(20), 313 Gurevich, A. Z., 329, 335(72), 376(72), 377(72), 381(72), 382(72) Guriev, S., 74 Guschlbauer, W., 448,457(69), 526,530(83), 535(83), 536(83), 541 Guss, J . K., 521 Gussin, G. N., 126, 139(73), 143(73. 146) Gutell, R., 75, 506 Guthrie, C., 471, 476, 494, 49337) Gutmann, H. R., 326 Gutte, B., 357, 359, 360, 432(255) Guy, A., 150 Guynn, R. W., 535 Gyenes, L., 363 Haar, W., 368, 376(306), 377(306), 379(306), 380, 381(306), 461 Haars, R., 123 Haas, B., 56, 543 Haber, E., 391, 394 Hadidi, A., 220, 22503). 233(8) Hadjiolov, A , , 237, 239(63)

597 Haenni, A.-L., 200 Haffner, P. H., 368, 369(307), 376(307). 377(307), 379(307), 386(307) Hagenbuchle, 0.. 147, 150 Hager, D., 66, 67(56), 75(56), 77(56), 94 Hager, D. A., 114 Hager,G. J., 111, 113(7), 116(7), 117(7), 144 Hager, L. P., 180 Hagerman. P. J., 390 Hagiwdra, M., 439, 447(31) Haimovich, J., 363 Hakam, A., 542 Hake, H., 273, 274(37) Halbrook, J. L., 318, 407,408(488), 410(488) Halbwachs. H., 64 Haldenwag, W. G., 62, 66(9, lo), 67(9, lo), 71(9, lo), 72(10, 64), 78(9, lo), 79(10), 80(9, 10) Hall, B. D.. 214 Hall, C., 242 Hall, G. I., 147 Hall, R. H., 568, 575, 576(31), 577(31) Hall, S. H., 498, 499(70) Hall, T. G., 551 Hall, Z. W., 12, 13(45), 14(53), 1358) Hallick, R. B., 152 Halling, S., 65, 76, 132 Halserna. I., 416, 423(555) Halvorson, H. 0.. 543, 549(227) Halvorson, H. R., 390 Harnaguchi, K., 393 Harna-Inaba, H., 176 Hamm, L., 24 Hammes, G. G., 392, 450 Hamprecht, R., 182 Hanawalt, P. C., 22 Handa, H., 148 Hankins, W. D., 579, 581(45) Hanson, A. W., 359 Hansen, U.,71 Hantgan, R. R., 392 Har, T. S., 18 Harada, F., 447 Harbers, E., 185 Harbers, K., 202 Harding, J. D., 130 Hardy, S., 239 Har-El, R., 538 Harmon, G., 577 Harpold, M., 248, 262(15)

AUTHOR INDEX Harriman, P. D., 202 Harrington, W. F., 386 Harris, B., 144, 147(166) Harris, E., 546 Harris, M. A., 572, 573(23), 574(23) Harris, T. J., 36, 496 Hartman, F. C., 327 Hartmann, G., 75, 76, 77( I12), 82( 1 12), 84(112) Hartstein, E., 505 Hartwell, L. H., 26 Haruna, I., 268, 274(4), 276 Harvey, C. L., 13, 14(54), 16(54), 17(54), 28(54, 68) Harvey, E. N., 237 Harvey, R.A., 526,531(84),532(84), 536(84) Haselkorn, R., 145 Hash, J. H., 465, 466(132), 467(132) Hashiguchi, M., 465, 466(135) Hashimoto, J., 465, 466(127, 142). 467(127), 468( 127) Hastings, K., 246 Haugenbuchle, O., 244 Haugen, T. H., 361, 362(275), 397(275) Hausen, P., 140, 141(156), 499 Hauser, H., 240 Hausmann, R., 88, 89(4, 11). 95, 96(41), 97(41), 102(4, 1 1 ) Hautala, J. A., 538 Havinga, E., 345, 347, 348(211, 212, 2131, 349(206), 350(213) Havinga, J., 319, 399(10), 400(10), 403(10), 404(10), 405(10), 409(10) Havron. A,, 344 Hawley, D. A., 268, 270(12) Hawley, D. M., 544 Hayashi, H.. 464 Hayashi, M., 544 Hayashi, R., 357, 358, 359(256) Hayakawa, S., 442 Hayes, F., 511 Hayes, M. B., 366, 367(297), 369(297), 370(297) Haynes, G. R., 195 Hayward, R., 100 Hayward, W. S ., 268, 269(15), 270(15), 276( 15) Hearn, R. P., 346, 352, 355, 388 Heath. E. C., 361, 362(275), 397(275) Hecht, L. I., 184, 1837)

Hecht, S . M., 53, 54(75), 57(75), 259, 551 Heckman, J.. 57 Heidelberger, C., 185 Heil, A., 63, 74(22), 75(22), 76(22), 86(22) Heineman, U., 437 Heinlein, K., 545 Heinrich, J., 202 Heinrikson, R. L., 331. 421(81), 423(81) Heller, S. R.,373 Hemphill, H., 76 Henkens, R. W., 390, 391(396) Henmann, H., 75, 76(107) Henner, D., 244 Henning, D.. 570 Henninger, M., 48, 49(56) Heppel, L. A., 448, 503, 525, 529(73), 535, 537(141), 539(108), 547 Herbert, E., 184, 185(6, II),188(26), 190 (11). 201(11), 202(11) Hercules, K., 48 Hermann, K., 168 Hermans, J., 386 Hermodson, M., 331 Herndley, D. D., 528, 530(100) Herrick, G., 65 Herries, D. G., 462 Herrlich, P., 491 Herzfeld, F., 65. 76(46) Herzog, W. R.,Jr., 390. 391(396) Hey. T., 49, 50(62) Heyden, B., 66 Heyneker, H. L., 16, 20(67), 21(67), 47 Hiatt, W., 77, 78(122), 79, 80(122, 131) Hibner, U., 176, 177(87) Hickey, E., 259 Hicks, M. L., 23, 24(109) Hieter, P. A., 550 Higgins, N. P., 5, 9, 17(19), 28(19), 33, 34. 36(18), 38(16), 41(35), 47(18) Hildebrandt, A,. 120 Hilderman, R. H . , 188, 195(48), 207(48), 214(48) Hill, D., 273, 276(39) Hill, R.,184, 185(12) Hillar, M., 418 Hillel, Z.. 75. 76(104) Hillen, W., 541 Hillenbrand, G., 175 Hilmoe, R. J., 448, 535, 537(141). 547

AUTHOR INDEX Hilmore, R. J . , 525, 529(73). 530. 535(112), 547( 1 12) Hilton, M . , 79, 80(132) Hindley, J., 273 Hinkle. D. C.. 12. 17(50) Hinnebush. A , . 79 Hinton. D. M., 34, 3.5(22), 36(22), 38. 40(22, 33). 44, 54i45), 56(33. 41) Hirai, K . , 519. 521(18), 522(18), 523(18), 524(18), 539, 543. 544 Hirose, S.. 427, 504 Hirs, C. H. W., 318, 325, 328, 358, 375(41), 398, 400(452, 453. 4611, 406(452, 453). 407. 408(488, 489), 410(66. 488, 489). 411. 421(261), 423 Hirsch, J., 75, 136 Hirt, B.. 140, 178 Hishinuma, F., 519, 521(18), 522(IX), 523(18), 524(18), 539 Ho. Y.-K.. 542 Hobbs, J., 190(54), 191, 192(54), 194(54), 197(54), 545 Hobbs. M. B., 22 Hodges. R . S.. 358, 405(260) Hodnett, J . L.. 326 Hodo, J. G.. 1 1 1 , 113(9). 116(9), 120(9), 123(Y) Hoes, C.. 347. 348(217), 350(213) Hoffman, C . H.. 546 Hofmann. K.. 335, 346. 347(105, 106), 350, 351(201), 443 Hofschneider, P. H., 486, 489(13) Hofsteenge, J., 398, 399(449),400(449). 406, 409(449) Hogan, J. J . . 506 Holbrook, D. J . , J r . , 427 Holder, S.. 74. 77(93). 81(93), 94 Holland, 1. B.. 268 Holland, M. J., 144 Hollender. R.. 548, 549(277), 550(277), 552(277) Holley. R. W . , 532. 5331223, 564 Hollingworth, B. R.. 503, 504(301. 50330) Holmes. B. E . , 342. 143(160) Holmes. R.. 507 Holrnes. R. R.. 432 Holmes. S. L., 287, 289(36) Holupirek. M . . 56, 543 Holy. A , . 449. 452(70),464. 467i70) Homandberg. G. A.. 356

599 Honda, B. M . , 149 Hondo, H., 201 Hong. C., 577 Hoogerhout, P., 348(217), 350, 356 Hooker. T . M . , Jr., 383, 384 Hooverman, L. L.. 454, 455 Horgan, W. F., 182 Hori, K.. 268, 273. 278, 279 Horiuchi, K., 169 Horn, V.. 71, 73(83) Horwitz, J., 382, 383(336), 384 Horwitz, M . S., 180, 181, 182(153, 154) Horwitz, S., 64 Hosakawa, S.. 412 Hosoda, J., 22 Hossenlopp, P., 139, l42( 140). l43( 140). 144(148) Houghton, M . , 142, 147 Hovanessian, A. G., 282, 283, 284, 285(12, 19. 2 0 , 286, 287, 288(28, 37), 290(37), 291(17,28), 292(12, 13), 295(13), 300(28, 37), 301(37), 304(80, 811, 305, 308(78, 80, 81), 311(19. 32, 33) Hovemann, B., 214 Howard-Flanders, P., 4 Howarth, 0. W., 389, 395(386) Howell, J. R., 327 Howes. N. K., 579, 580(49), 581(49) Hozumi. T., 20 Hsieh, W. T., 525 HSU,M . - C . , 383 Hu, A , , 535 Hu. M . . 34 Hua, H., 34 Huang, H., 90, 93(24, 371, 94, 98(24), 101(24), 104(37), 10379) Huang, A., 88, 105(7) Huang, W. M . , 176 Huberman, J., 179 Hubert, E., 243 Hubert, E., 539,548(171), 549(171), 550(171) Huebscher. D., 182 Huet, J., 126. 128. 137 Huez, G . . 243. 539, 548(171). 5491171). 550( 171) Huff, N . , 228, 244(39) Hugli, T. E., 341, 426 Huizinga, J . D., 319, 400(7), 406(7) Hummel, J. P., 450 Hunt, R. E., 519

600 Hunt, T., 306 Hunter, T . , 179 Hunter, M . J . , 421. 422 Huppert, J . , 225(25), 226, 233(25) Hurwitz, E., 363 Hurwitz, J.,4, 12(13), 15(13), 16, 17(13,69), 27(12, 13), 31. 32(1, 2), 33(1. 2). 34(12), 3312). 36(12), 40(12). 41(1. 12). 42(2. 12). 45(2), 47(2, 12). 51, 62. 68, 69. 86, 108, 141, 149, 152, 156. 157(11), 158(11), 159(11), 160(11), 161(15), 165(15), 166(11, 15), 168, 169(45, 46), 170, 171(21), 173, 180, 181, 182(151, 154). 185, 227, 247, 248, 249(22), 250(22), 251(22), 252(22, 33), 255(22), 261(33), 262, 263(22), 498, 499(71), 558, 568 Hutchinson, D. W., 519. 521(17), 521(17), 522(17), 524(17), 525(17), 539(17), 540, 546( 177) Hutter, R., 538 Hutton, J . , 233, 240(55) Hyman, R.. 89 Hynie, S . , 528, 534(97)

Iborra. F . , 126, 128, 134(79), 135, 136(113), 137

Ichikizaki, I . , 439, 447(31) Igarashi, K . , 427, 504 Igarashi, S. J., 201 Igo-Kemenes, T . , 202 Iijima, H . , 324, 32338, 39), 334(38, 39) Ikeda, H . , 502, 511(16) Ikeda, J . - I . , 181, 182(151) Ikeda, J.-E., 156, 160, 168, 169(46), 170, 171(21), 181, 182(154)

Ikehara, M., 34, 37, 38(23, 311, 42, 43(44), 44, 54(47), 55(47), 56(79), 541, 544

Ikeuchi, T., 68 lmahori, K., 447, 453, 454, 455, 456, 457 Imahori, T . , 448 Imamoto, F., 80 Immartino, A. J., 343, 344(168) Inagaki, F., 333, 444, 44351). 446(51) Ingles, C. J . , 135 Inman, R., 179 Inouye, H . , 268. 270(1 I) Inoue, N . , 442 Inouye, H . , 540 Ipata, P. L., 447

AUTHOR INDEX Irie, M . , 323, 412, 414, 426, 440, 447. 456, 458(63), 463(63, 93)

Isaksson, L. A., 566 Ishihama, A., 63, 68, 74(18, 211, 75(21), 76(18, 21), 77(113), 79, 81. 82(21, 142), 84, 129 Ishihara, Y . , 284, 285(27) Ishii, S . , 80 Ishikura, H., 502, 51 l(16) Ishiwata, K . , 436 Ishiye, M . , I11(24), 112. I13(24), 116(24) Isomaa, V . , 241

Ison, R . R., 380 Israeli, R . , 282 Itagaki, A., 180 Itagaki, K . , 530 Itakura, K., 20, 52 Ito, H.. 439, 447(31) Ito, K . , 129 Itoh, T., 173 Itoh, Y . H . , 582 lukova, M . N., 467 Ivanova, G. S . , 449, 452, 465, 467(70) Iwakura, Y . , 79, 84 Iwata, T . , 467 Iwatsuki, N., 22 Izumiya, N., 436, 443(9) Izzo. P., 562, 563(22) Jackson, R. L., 358,398,400(452), 406(452), 410, 421(261)

Jacob, S.. 218, 220(3), 221, 222(13), 223(!3), 225(13), 226(13), 230(13), 231(13), 232(3, 13. 42), 233(13), 237(13), 238. 239(7 I1, 242 Jacob, S . T . , 110, 136 Jacob, T., 83 Jacobs, M. F., 147 Jacobson, A , , 496 Jacobson, A. B . , 514 Jacquemin-Sablon, A., 7, 8(29). 9(29), 15, 16(29), 17(29), 18(29), 19(29), 22, 27(29, 60) Jaehning, J . A . , 67, 73(65), 111(29), 112, 113(8, 10, 29), 116(11), 119 Jahnke, P., 545 Jailkhani, B. L., 326. 334, 420, 421(47), 422(47) Jakubowski, H . , 560

AUTHOR INDEX Janekovic, D., 76, 77(116) Janik, B., 545 Janion, C., 541, 543198) Jankowicz, C.. 572, 573(23). 574(23) Janne, O., 241 Janner, F., 582 Janski, A. M., 436 Jantzen, E., 445 Jantzen, H. M., 182 Jardetzky, O., 462, 355, 366(237), 367(237, 300), 368(237, 294), 369(237, 294, 300), 370(237), 372(294, 315). 373(237, 294, 315), 376(294. 300), 377(294, 300), 379(237, 300), 381(300), 386(237). 396075). 429 Jarvis, A. P., 284, 285(20), 286(20), 296(20) Jaskunas, S. R., 494 Jeanteur, Ph., 498. 499(68, 69) Jedlicky, E., 184. 185(10), 186(10), 187(10), 189(10), 190(10), 194(10), 196(10), 201( 10) Jekel, P. A., 398, 400(446), 405(446), 409(446), 42 l(446) Jelinek, W., 104, 235. 236(69), 238, 498 Jendrisak, J. J., 65,66(43), I 1I , 114, I15(35), 116(8), 117(8, 43), 120, 121(61), 122(58), 124(43, 61). 128(11), 130, 132(86), 145(11, 58), 146 Jensen. D. E., 428 Jentoft. J. E., 320, 324(20), 376(37) Jentoft, N., 320, 324(20), 376(20, 34, 37) Jerusalimsky, A. P., 433 Jervis, L., 436 Jewett, P. B., 427 Jewett, S. W., 341 Jiamachello, P. F., 135 Jippsen, P. G. N., 502, 513(23) Jockusch, B. M., 122 Johnsen, E., 363 Johnson, B. C., 146 Johnson, J., 67 Johnson. K. L., 542 Johnson, P. F., 32 Johnson, R. E., 395 Johnson, R. N.. 327 Johnston, D. E., 144, 146(170) Johnston, L. H.. 26 Johnston, M. I.. 288 Joklik, W.. 219, 220(4), 221(4). 224(4),

60 I 225(4), 226(4), 229(4), 236(4), 237(4), 238(4) Joklik, W. K., 152 Jolly, J., 88, 93(10), 98(10), 102(10) Jolly, S. O., 152 Jones, A. S., 540 Jones, G. H., 76, 381 Jones, S. S., 292 Jordan, B. R., 515 Jori, G., 337, 344, 356(117) Jorm, T. M., 63(33), 64,86(33) Jou, M., 228 Jovin, T. M., 4, 5(14) Jungmann, R. A , , 136 Juntt, N . , 520 Juntz, G. P. P., 112 Juodka, B. A., 36, 37(25) Juretschke, H. P., 329, 330(74), 33374). 346(74), 373(74), 374(74) Justesen, J., 287, 292(41), 301(41), 302(41), 303(41, 68) Kaback, H. R., 520 Kabasawa, I., 407, 408(489), 410(489) Kacian, D. L., 275 Kaczkowski, J . , 185(41), 186, 187(41), 189(41), 190(41), 192, 193(56), 194(56), 195(56) Kadesch, T., 75 Kaempfer, R., 282, 548, 549(277), 550(277), 552(277) Kaesberg, P., 270, 274 Kafatos, F. C., 548 Kainuma, R., 22 Kaiser, A. D., 4, 27 Kakefuda, T., 173, 174(68) Kakiuchi, N., 541 Kakulas, B. A . , 416 Kallenbach, N . R., 32, 40(4), 41(4) Kalnitsky, G., 338 Kamen, R. I., 268, 269, 270(9, 20), 272(20), 273(20), 276(20) Kamesaka, Y., 426 Kamikubo, T.. I 1 1(23. 24). 112, I13(23, 24). 116(23, 24), I19(23) Kamm, R. C., 425 Kammen, H. O., 570, 571(15), 572, 580(15), 582(15) Kan, L. S., 52

602 Kanaya, S., 437 Kang, H. S., 32 Kann, H. E., 497 Kaplan, L. J., 382 Kaplan, L. M., 180 Kaplan, R., 505, 506, 512(42), 525, 538(69) Kaptein, R., 330, 335(80), 339(80), 346(80), 366, 367030, 301, 302, 303), 370(80), 371(80), 37260). 372(80), 377(80), 378(80), 386(80), 389(80), 395(80) Kapitza, E. L., 494 Kar, D., 431 Kara, J., 528 Karabachyan, L. V., 323, 429 Karam, J. D., 25, 47 Karkas, J., 223 Karpeiskii, A. M., 528 Karpeiskii, M. Ya., 528 Karpeisky, M. Ya., 323, 329, 335(71, 72, 73), 364(73), 369(71), 376(72), 377(72, 73), 379(71, 72). 381(72), 382(72), 406(73), 415(73), 431 Karpetsky, T. P., 219, 416, 426, 433(544) Karplus, M., 385, 423 Karstadt, M., 84 Kartha, G . , 324, 325(38). 334(38) Kasai, H., 442, 446, 447, 579, 581(52) Kasai, K., 465 Kasai, T., 502, 507(10), 508(10), 509(10), 512 Kassavetis, G., 70, 86(79), 90, 99, 100, 104, 108(76) Kassel. R. L., 328, 433(57) Kastern, W., 260 Kates, J., 88. 105(6) Kato, I., 394 Kato, T., 436, 443(9) Katoh, Y.,519, 525(20) Katz, G.. 564 Katze, J. R., 579. 580, 58100, 54), 582(54) Kaufman, E. E., 572, 573(21) Kaufman, G., 179 Kaufmann, G . , 32, 37, 40(4), 41(4), 42, 52. 58(29), 531, 533(113), 534(113), 535, 536(126, 142), 543(126), 544(127, 228), 5431 13), 546(228), 547( 142). 548(142) Kaufrnan, R. J., 148 Kawaguchi, H., 132 Kawakita, M.,306, 307(86) Kawano, Y.,444, 445(51). 446(51) Kawasaki, K., 443

AUTHOR INDEX Kay, J., 439 Kayne, M. S., 573 Kaziro, Y., 278 Kedinger, C., 117, 119(45), 120(45), 122(45), 123(45), 124(45), 126(45), 127(45), 138, 139, 148, 150 Keil, T. U., 486, 489(13) Keir, H. M., 18 Keith, G., 563 Keith, J. M., 249, 560 Kellenberger, E., 21, 176 Keller, E.. 577, 578(40) Keller, W., 111(28), 112, 113(28), 148 Kelly, K. H., 122 Kelly, T. J., Jr., 178, 179, 180, 181, 182(150) Kempf, J., 425 Kenkare, U. W., 352 Kennell, D., 494 Kenney, W. C., 465, 466( 128) Kent, S . B. H., 421 Kenzo, O., 498. 499(71) Keren-Zur, M., 364 Kerling, K. E. T., 347, 348(211, 212, 213, 217), 349(206), 350(213), 356 Kerr, D. S., 191, 196(50), 197(50), 207150) Kerr, 1. M., 282, 283, 284, 285(12, 17, 28). 286(17), 287(17), 288(28, 37). 290(37), 291(7, 15, 28), 292(12, 13, IS), 293(52, 53), 295(13), 296(52, 53), 297(53), 298(53), 299, 300(28, 37), 301(37, 49, 64), 302(49.58), 304(82), 305(52,53,54), 306(49, 52, 53, 58), 307(52, 53, 58). 308(79,82), 309(82), 310(58), 312(52,53) Kerr, S . J., 557 Kersten, H., 558, 559, 563(12), 564(9), 565(9) Kersten, W., 559, 563(12) Keshgegian, A . , 244 Kessler, B., 520 Kessler, R. E., 520, 539(26) Key, J. L., 120, 123(59) Keyser, G. E., 540 Khan, A., 545 Khandker, R., 428, 431(631), 432(631) Khechinashvili. N. N., 386 Khorana, H. G., 8, 9(34), 15, 16(62), 19, 20, 21(82), 28(62), 83, 84, 483, 503 Khurshid, M., 545 Kidd, G . H., 117, 119(46), 1 2 3 ~ 6 1 124(46), , 152

AUTHOR INDEX

Kieras, R., 238 Kikuchi, Y., 539, 543, 544(226) Kim, C., 259 Kim, J., 407 Kim, S., 323 Kimball, A. P., 326 Kimchi, A., 282, 287, 290, 298, 301(45), 304(59), 30359, 75). 307(59). 308(59), 309(83), 310(45) Kimhi, V., 68 Kimhi, Y., 520, 521, 522(39), 524(39), 525(39), 529(39), 534(39), 535, 536(139), 547 Kimura, G . , 180 Kimura, S., 444, 445(50, 52), 446(52), 457(50), 463(50, 92) Kindler, P., 486, 489 King, A,, 75, 76(106) King, N . L. R . , 366, 367(299), 370(299), 372 King, T. P., 328 Kingsbury, D. W., 551 Kingston, R., 66. 67(59), 75, 104 Kinscherf, T. G., 511, 537, 538 Kinsley-Lechner, E.. 184 Kirk, K. L., 347, 355(208), 365 Kirschenbaum. A . H., 215 Kirschner, M. W.. 124 Kirveliene, V., 36 Kitamurd, N.. 502, 511(16) Kitano, S.. 53, 54(75), 57(75) Kjellin-Strkby, K., 566 Klapper, M. H., 331 Klee, C. B., 521, 522(44), 523(44), 524(44), 526(53, 54), 527(54), 530(90). 532 Klee. W. A,, 356, 387, 396(380). 405 Klein, T., 42 Kleid. D., 72 Klemperer, H. G., 195 Kleppe, K., 8, 9(34). 14, 16(59), 17(59). 19(59), 21(59), 28 Kleppe, R. K . , 14, 16(59), 17(59), 19(59), 2 I(59) Kline. L., 475 Kline. L. K., 558, 564(6), 566(6), 568, 574(1), 575, 576(30, 31), 577(31) Klita, S., 551 Knable. T.. 56 Knable, T., 533, 543 Knapp, G., 32

603 Knight, M., 292, 293(53), 296(53). 297(53), 298, 305(53), 306(53), 307(53), 3 12(53) Knippers, R., 240 Knoller, S., 282 Knopf, K. W., 7, 8(28), IO(28) Knowles, J. R., 345, 388(188) Knox, J. R., 359 Kobata, A., 410 Kobayashi, N., 137 Kobayashi, R., 398, 400(461) Kobori, J., 160 Koch, G., 231, 232(43) Kochoumian, L., 328 Koehler, K. A., 427 Koerner, J. F., 51 Koester, H., 53 Kofoid, E. C., 56 Kohn, K. W., 497 Kolakafsky, D., 184 Kole, R., 471, 472(9), 473(9, lo), 474(9-12), 568 Koller, T., 275 Kolodner, R., 28, 174, 175(76-78) Komoriya, A,, 365 Konanti, T., 388 Kondo, M., 268, 269, 270(10, 20). 272(10, 20). 273(20), 276(20), 278(22) Konishi. Y., 393 Konrad, E. B., 24 Koopmans, M. A. G., 418, 419 Kop, J . , 506 Kornberg, A., 4, 5(14), 9, 17, 33. 68, 156, 157(9), 158(9, 10). 160, 161(22). 162(22. 23, 30, 36-38), 165(26, 34), 168(5), 170(22, 43). 171(22, 23, 43), 176, 178, 237, 518, 525(5), 529(5), 539(5) Kornberg, R., 107 Kornberg, S . . 68 Korner, A,, 577 Korsten, K., 88, 89(1I), 10211 I ) Koshland, D. E., Jr., 448 Koski. R. A., 473, 474(12. 13), 47313). 476(13). 479(13) Kossel. H., 544 Kotelchuck, D., 355, 385 Koths, K., 162, 165(41) Kotick, M. P., 545 Kozak, M., 243 Kozarich, J . W., 551 Kozinski, A. W., 22, 25(95)

604 Kozinski, P. B., 22 Kraft, N., 416, 423 Krakow, J.. 63(33), 64, 84, 86(33) Kramer, F. R., 275, 276(55), 279 Kramer, R. A., 486, 491, 493(7) Kranias, E. G., 136 Kraus, A. A., 416 Krebs, G., 1 Il(27). 112, I l3(24), I17(27), 123(27) Kreiser, T. H., 545 Krishman, I., 284, 2832.5) Krishna, R. V., 537, 538(153, 159) Kroger, H., 62 Kroger, M., 191. 192(55) Kroger, M., 542 Kroll, M., 227, 237(30), 238(30), 240(30) Kronenberg, H. M., 552 Krug, M., 44, 5348). 56 Krug, R.. 185, 259, 260(54), 265(56) Krug, R. M., 464 Kruger, D., 88 Kubokawa, S., 475, 479(17), 480(17), 482( 17) Kuchino, T., 180 Kuchino, Y.,561, 573 Kuechler, E., 540 Kuentzle, C. C., 182 Kuhn, J., 538 Kumagi, H., 427, 504 Kumar, A., 28 Kung, H.-K., 532, 535(121) Kuninaka, A., 519, 525(20) Kunitz, M., 318, 425 Kuntzel, H., 150, 151(217) Kuo, C. H., 273 Kuper, H., 398, 400(459a) Kupper, H., 93, 103(35), W(35) Kuppers. B., 274 Kuriyama, Y., 439, 447, 467 Kurland, C., 239 Kuroda, Y., 53, 54(75), 57(75) Kuroiwa, A., 137 Kushner, S. R., 538 Kusmierek, J. T., 542 Kutter, D., 67 Kutter, E., 51 Kuwata, T., 286. 311(29), 312 Kwong, T. C., 559, 566 Kyner, D., 423

AUTHOR INDEX Labhardt, A. M., 355, 391(245, 246) LaBone, T., 573 Lacks, S. A., 426, 489 Lacroute, F., 127, 135(77) Laemmli, U. K., 124 LaFiandra, A., 259 Laipis, P. J., 4, 6(15), 17(15), 18(15) Lambert, J. M., 421 La Montagna, R., 398, 399(457), 400(457), 412(457), 414, 421(457) Lan, L. T., 331 Landers, T. A., 268, 269, 270(12, 13), 271(23), 272(27,28), 277(13,27), 278(23) Lane, C., 259 Lang, N., 62, 66(10), 67(10), 71(10), 72(10), 78(10), 79(10), 80(10) Lanka, E., 173, 175 Lapanje, S . , 395 Lapidot, Y.,323, 364, 426(30), 459 Larson, D., 147 La Rue, B. F., 471, 482(7), 483(7) Laskey, R. A., 148 Laskowski, M., Jr., 356 Last, J. A., 34, 35(20), 37, 38(30), 39(30). 42(20), 5330) Laten, H., 58l(58), 582 Lathe, R., 106 Lattke, H., 130, 131(85) Laulhere, J. P., 152 Law, G. R., 436 Lawrence, M., 86 Lawrie, J., 77, 78(122), 80(122) Laycock, D., 249 Lazarow, P. B., 423 Lazdunski, M., 341 Leach, M., 47 Leach. S., 82 Leautey, J., 522 Lebleu, B., 282, 304, 306, 307(86), 31 I Lebowitz, J., 142 Leboy, P. S., 561, 562(19), 563(21) Lechner, R., 179 LeClerc, J. E., 28, 174, 175(76) Leclerq, M., 243, 539, 548(171), 549(171), 550( I7 I ) Lecocq, J. P., 106 Leder. P., 148, 547 Lee, B., 359, 364 Lee, G., 78, 80(127)

AUTHOR INDEX Lee, H. J . . 494 Lee, J . C., 440 Lee. L., 182 Lee, N., 80 Lee, S., 227, 231, 232(46) Lee, Y. F., 496, 537 Lee, W. Y., 363 Lee. Y. M., 331 LeGendre, S. M., 550 Lehman, I . R.,4,5,6(11),8,9(30), 10(30), 12, 13(30, 45). 14(17, 53), 15(24, 57, 58). 16(24, 57), 17(21, 57, 66). 18(1 I), l Y ( 1 I , 21, 57), 22, 24, 28(22. 24). 47, 176 Lehrach, H., 521, 522(48) Leibart, J. C., 26 Leighton, T.. 65 LeGenaar-Van den Berg, G., 398(463), 399, 400(463), 403(472), 404, 406, 407(483), 409(463) Leilausis, A . , 176 Leineweber, M., 185(40), 186, 187(40), 188(40), 189(40), 194(40), 196(40), 502, 507( I I) Leis, J . P., 496 Leis, J.. 31, 32(1), 33(1), 41(1) Lengyel, P., 282, 284, 285( l8), 287, 288(38), 289(38), 290138), 292(38), 294(38), 299(47). 300(38.47), 301(38,47), 303(47). 304(18, 38), 305(38, 76), 306(76), 307(86) Lennette, E. P., 329. 330(70), 331, 335(69) Lennette, E. T., 512 Lenstra, J . A., 330, 335(80), 339(80), 345, 346(80). 367, 370,371,372(80), 373,377, 378, 386(80), 389(80), 395(80), 398, 400(445, 446), 403(445), 404(190), 405(445, 446), 406, 409(446), 421(445, 446) Leonard, N . J., 38, 40(36), 53(36), 540 Leone, E., 398. 399(457), 400(457), 411, 41 2(457), 414(500), 421(457) Leong, K., 259 Le Page, G. A,, 185 Le Pecq, J. B., 425 Lescure, B., 139. 143(144, 145, 149, 150), 144(149, 150), 145(149) Letendre, C. H., 521, 522(43), 523(43) Leung. K.-L., 548 Levens, D., 150, 151(216)

605 Levin. D.. 261 Levin, D. H., 537 Levin, G. J . , 18 Levisohm, R., 273 Levinson, W. E., 131, 132(91), 184, 192 Levinthal, C., 385 Levit, S..349, 350, 351, 357 Levy, C. C., 219, 416, 426, 427, 433(544), 550 Lewin, B., 138 Lewis, H., 494 Lewis. J. A., 304 Lewis. J. B., 181 Lewis, M . K . , 138, 139(133), 141(133). 142(133), 143(133), 144(133) Lezius, A. G . , 65 Li. J., 74, 80(91) Li, H. J . , 578 Li, J . R.-T., 429 Li, L.-K., 395, 396(423) Li, Y., 328 Liang, C.-J., 410 Libertin, C., 436, 452(6) Libonati, M.,412, 413, 414(509, 525). 415, 429(530, 540) Libor, S., 332 Lichy, J., 181 Lichtin, N. N., 342, 343(165) Liebman, K. C.. 185 Liegel, J., 537 Liehr, J. G., 447 Lielausis, A., 21 Lienhard, G. E., 427 Lill, H., 75 Lill, U., 76, 77(112), 82(112), 84(112) Lillehaug, J . R., 8,9(34) Lilley, D. M. J., 142, 147 Lim, V. I., 386, 406 Lin, J . , 74, 77(93), 81(93), 94 Lin, J . J . C., 214 Lin, M. C., 356,357(252),389(252), 418(252), 432 Lin, S.,20 Lin, Y., 546 Lin, Y. C., 237 Lindahl, L., 494 Lindahl, T., 4, 10(16), 12, 17(16), 18(49), 19(16, 80), 203 Lindquist, R. N., 427

606 Link, G., 1 I1(25), 112, I13(25), 117, I19(46), 123(25, 46), 124(46), 137 Link,T. P.,336,337(116),347(116), 356(116) Liorancaite, L.. 36 Lipkin, V. A., 74 Lipmann, F., 249, 257, 261(30) Lipper, C., 152 Lipsett, M. N., 572, 573(21) Littauer, U . , 68, 243 Littauer, U. Z., 37, 42, 58(29), 185, 196(30), 211(30), 212(30), 215, 518, 520, 521, 522(39, 42), 523(42), 524(39, 42), 525(5, 39, 42). 529(5, 39, 42), 531, 533(113), 534(39, 113), 535, 536(126, 142), 539(5), 540, 543(126), 544(127, 228), 5431 13), 546(42,228),547(142),548(140,142,171, 172, 263, 264), 549(140, 171, 262, 263, 264, 274, 276), 550(42, 140, 171, 172, 264). 551(262, 264). 552(263, 264) Littauer, V. Z., 559 Little, J. W., 5, 12, 13(46), 15(20), 17(20), 18(20, 46). 19(20), 27(20) Litvak, S., 184, 185(10), 186(10, 33). 187(10, 33), 189(10), 190(10), 194(10, 33), 196(10), 197(33), 200(33), 201(10) Liu, C.-C., 176, 177(87), 178(100, 101) Liu, D. K., 423 Liu, M. Y., 320, 323(21), 330(21), 335(21), 364(21), 377(21), 406(21), 415(21) Liu, N.-J., 546 Liu, S., 363 Live, T. R., 6, 7, 8(29), 9(29), 15(23). 16(29), 17(29), 18(29), 19(29), 22, 27(29) Livingston, D. M., 132 Llorens, R., 323 Lockard, R. E., 57, 259 Loeb, L., 63(34), 64,7334). 96 Loeb, L. A., 131(93), 132 Loewen, P. C., 8, 9(34) Logan, D. M., 503 Logue, A. D., 352, 391(226) Lohman, T. M., 134 Lomedico, P. T., 397 Long, E., 138 Longacre, S., 227 Longiaru, M., 181, 182(154) Longo, D., 502, 507, 511(12) Lord, R. C., 386 Loring, D., 149, 262 Losick, R., 62, 63(24, 32). 64, 66(9, 10, 32).

AUTHOR INDEX 67(9, lo), 71(9, lo), 72(10), 76(32), 77(111), 78(9, 10, 32). 79(10, 32). 80(9, 10, 130), 83(24, 32), 88, 89, 105 Loviny, T., 490 Low, K. B., 23, 489 Low, R., 160, 161(22), 162(22, 23). 170(22), 171(22, 23) Lowe, P., 66,67(56), 75(56), 76(106), 77156). 94 Lowe, P. A., 114 Lozeron, H. A., 494 Lubben, T. H., 34 Lubochinski, J., 521, 527(51) Lucas, J. M., 526 Lucas-Lenard, J . , 526, 528 Luce, R., 273 Lucas, S. J., 9 Lucas, Z. J., 176 Ludlum, D. B., 540 Ludwig, M. L., 421, 422 Luftig, R. B., 48 Lund, E., 476, 494, 49337) Luse, D., 262, 263(65) Luse, D. S., 115, 147(38), 148 Lustig, A,, 150, 151(216) Luton, B. A., 431 Lyerla, J . R.,Jr., 355, 375 Lyle, H. L., 48, 49(58) Lyn, G., 4, 12(13), 15(13), 17(13), 27(13) Lyons, H., 425

McAllister, W., 90,92,93,98(33), 99(29,33), 100(29,58), 103(35), 104(35), 105(29, 33) McAuslan, B., 88, 105(6) McCalla, J . I., 201 McCauley, J . W., 307 McClain, W. H., 214,215,477,479,480,495 McCloskey, J. A., 447, 568, 572(5), 575(5), 578(5), 580, 581(54), 582(54) McClure, W., 69, 71, 144, 146(170) McConnell, D., 68 McConnell, D. J., 97 McCulley, C., 47 McDonald, C. C., 386 McDonald, M. R., 318 MacDonald, R. J., 361, 427(274) McGann, R. G . , 186, 187(45), 188(45), 190(45), 194(45), 196(45), 199(45), 207(45), 210(45)

AUTHOR INDEX MacGee, J., 57 McGinnis, J., 496 McGrath, J., 88, 89(8), 91(8), 92(8), 94(8), 98(8), 99(8) McHenry, C., 158 McKenzie, J. M., 423 McLaughlin, P. J., 403 McNamara, P., 520 McMacken, R.. 156. 158, 161, 162(30) MacNaughton, M., 498 McPhie, P., 526, 531(86), 533(86), 537(86) Mackey, J. K., 533. 544(128) Maale, G., 231 Mace, D. N., 176 Mache, R . , 152 Machiori, F., 335, 344 Machuga, E., 331 Macklis, J. E., 494 Madison, J. T., 532, 535(121, 122) Magnusson, G., 178 Magazin, M., 174 Magrum, L. J., 506 Mahal, G., 559, 563(12) Maheshwari, R. K., 311 Mahler, H. R., 184 Maidhof, A , , 227, 232(31) Maitra, U., 86,90,93(24, 37). 94,98(24, 36), 100, lOl(24, 67), 102(67), 103, 104(37, 71), 104, 105(79), 247, 490 Maiyorov, V. I., 34, 35(17) Maizel. J. V., Jr., 496 Majors, J., 80 Majumder, H., 103, 104(71) Malathi, V. G., 31, 32(1, 2), 33(1, 2), 34(12), 36(12), 37(12), 40(12), 41(1, 12), 42(2, 12), 45(2), 47(2, 12) Malcolm, S., 123 Malicka-Blaskiewicz, C., 416 Malone, P., 80 Malorni, M. C., 412, 413(517), 414(517) Mamaeva. 0. K., 528 Mandel, J. L., 135, 138, 139(132) Mangel, W., 70 Manjula, B. N., 335, 336(109), 337(110) Manley, J. L., 147, 148, 263 Mans, R . , 228, 231, 244(39) Manthey, A. E., 34. 35(22), 36(22), 40(22) Maor, D., 416, 433(545) Marbaix, G., 243, 539, 548(171), 549(171). 550(171)

Marchenko. T., 74 Marchiori, F., 346, 347, 350, 351(104, 220), 352. 356, 388 Marcu, D., 564 Marcus, F., 320, 323(24) Marcu, K. B., 564 Mardachev, S. R . , 520 Mardiney, M. R., Jr., 416, 433(545) Marians, K. J., 20, 156, 160, 171(21), 173, 174(72) Marinus, M. G., 559, 566 Markelova, N. Yu., 467 Markham, A . F., 55, 56(79) Markham, R.,503 Markle, H. V., 417 Markley, J . L., 366, 367(296), 368, 370(296), 372, 373(316), 376, 377, 379(316) Markova, I., 74 Markuckas, A. Y.. 36, 37(25) Maroney, P. A . , 284, 285(22), 286, 287(22), 293, 298, 299(56), 300(56), 302, 304(66), 306(66), 310(22, 61) Marshall, M. V., 144 Marshall, R. D.. 410 Martelo, 0. J., 136 Martens, P., 179 Martial, J., 133, 134(99) Martin, E. M., 292, 301(49), 302(49), 305, 306(49), 309(84) Martin, P. D., 36.5, 437 Martin, R., 180 Martin, S . , 246.248(11), 249(17, 18). 250(17), 251(18, 19), 252(18), 254(18), 255(19) Martinez, A . , 84 Marx, P. A . , Jr., 551 Marzotto, A., 350, 352 Masamune, Y., 22, 23, 27, 28(129), 174, 175(76) Mase, K., 417, 424 Masiakowski, P., 186, 187(46), 188(46), 193(43, 44), 194(70), 195(44), 196, 197(70), 198(43), 199, 202(46), 203, 204, 205(44), 206, 207(44, 70, 96). 208(70), 210, 21 1(70), 212(44) Massey, V., 346, 352(198) Massoulie, J., 541 Mathelet, M., 18 Matheson, R. R., Jr., 344, 345(184), 386, 388(184, 186), 394(367), 395(186), 396(186, 367, 372), 406, 429

608 Mathews, E., 22 Mathews, M. B., 181 Mathias, A. P., 418, 425(578), 462 Mathis, D. 3.. 137(131), 138, 150 MatouSek, J., 433 Matrisian, P. E., 423 Matsui, T., 11 1(26), 112, 113(26), 119(26), 127, 149 Matsuo, H., 369, 444, 445(50, 52), 446(52), 457(50). 463(50, 92) Matsuura, S., 346, 347(203), 354, 368(203), 374(203, 203a), 436, 443(9) Matthaei, H., 542 Matthews, C. R., 332, 333(93), 369, 389(96, 311) Mattia, C. A,, 413 Mattoccia, E., 147 Matzura, H., 63, 74(20) Maurer, H. R., 122 Maurer, W., 368, 375, 376(306), 377(306), 379(306), 380, 381(306), 461 Maxam, A. M., 464 Maxam, A., 497 Maxwell, E., 260, 265(58) Mayhack, B., 511, 512 Mazetti, G., 242 Mazumdar, S. K., 337, 3631 19), 373(119) Mazus, B., 131, 132(87), 136 Mazzarella, L., 413 Meadows, D. H., 355, 366, 367(237, 300). 368(237, 2941, 369(237, 294, 300), 370(237), 372(294, 315). 373(237, 294, 3151, 376(294, 300), 377(294, 300), 379(237, 300), 381(300), 386(237), 462 Means, G. E., 320, 323(22) Meares, C. F., 65, 132 Mee, L. K., 342 Mehta, J. R., 540 Meilhac, M., 147 Meisenberger, O., 75, 76(107) Meisner, I., 423 Melchior, W. B., Jr., 332 Melechen, N. E., 4, 5(14) Melton, D. A., 215, 475 Meltzer, S., 244 Mendecki, J., 227, 231, 232(46) Mendelsohn, S. L., 425, 427 Mendelson, S., 48 Menke, G., 437 Merrick, W., 259

AUTHOR INDEX Merrifield, R. B., 357, 358(256), 359(256), 360, 405(260), 432(255) Mertens, P., 263 Mery, E., 525 Mertz, J. E., 15, 27(61) Meselson, M., 4 Metz, D. H., 282, 309 Meurs, E., 286, 287, 304(80), 305, 308(78, 80), 31 1132, 33) Meyenburg, K. V., 505 Meyer, B., 72 Meyer, D. H., 416 Meyer, F., 215, 521, 530(87) Meyer, R., 227 Meyer, R. R., 161 Meyer, W. L., 416 Meyhack, B., 57, 58(96) Meyhack, I., 511 Michaels, G., 150, 151(213) Michel, O., 184, 214(18) Michel, R., 184, 214(18) Michelson, A. M., 291, 296(48), 535, 540, 541, 542, 546 Migchelsen, C., 325, 369(42), 376(42), 377(42), 406(42), 407 Mignery, R. E., 564 Mii, S., 521, 523(46), 525, 529(71, 72). 530(46), 531(46), 535, 537(141) Mikhailov, S. N., 529, 534(97) Milchev, G., 237, 239(63) Mildvan, A., 96 Mildvan, A. S., 63(34), 64, 7334). 131(93), 132 Miller, J. F., 386, 387(373) Miller, J. P., 185, 186(35), 187(35), 188(35), 1891351, 191, 192(35, 52), 194(52), 197(52), 199(35), 200(35), 201(52), 202(52, 79). 203(79), 207(79) Miller, J. S., 73 Miller, M. J., 268, 270(12) Millette, R. L., 491 Milliman, G. E., 395, 396(426) Mills, D. R., 268, 273, 275, 276(55), 279 Millward, S., 246, 264 Minkley, E. G., 491 Minks, M. A., 290,293,299(56), 300,301(46), 302(46). 303(46), 304(46, 66). 306(66) Misra, T. K., 476, 483124) Mitra, T., 57 Mitsui, Y., 323, 324, 364(40)

AUTHOR INDEX Mitsuyasu, N., 436, 443(9) Miura, K., 246 Miura, K. I., 202 Miyake, T., 42, 43(44), 44, 54(47), 55(47), 56(79) Miyazaki, M., 570 Miyazawa, T.. 333, 444. 445(51). 44601) Mizurnoto, K., 249. 257, 261(30) Mizuno, D., 137, 417, 502, 503 Mizuuchi, K., 160 Modrich, P., 5 , 8, 9(30), 10(30), 13(30), 14. 15(57), 16(57), 17(21, 57), 19(21, 57), 24 Modyanov, N.. 74 Moffatt, 381 Mohr, S. L., 464 Moldave, K., 184 Molko. D., 150 Molloy, G. G., 235, 498 Monastryskaia, G., 74 Monier, R., 515 Monroy, G., 149, 248, 251 Montagnier, L., 287, 304(80), 305, 308(78, 80), 311(32, 33) Montibeller, J., 335, 347(106), 350(106), 351(106) Moore, D., 74, 77(93), 81(93, 94) Moore, S., 319, 320. 321(15), 324, 327, 328, 339(15), 357, 358(256), 359(256), 400(15), 405(35), 413, 414(55), 415, 418, 419(574), 420(15, 574), 421, 425(55), 426, 427(59), 432(255), 433, 437, 440(23), 462(23) Moran, C., 78 Moran, L., 176 Morelli, G., 175 Morgan, A., 83 Morgan, M., 246, 259, 260(54) Mori, C., 427 Moriyama, T., 423 Moroder, L., 335, 346, 347, 350(104), 351(104, 220). 352, 356. 388 Morozova, V. G . , 465 Morrice, L. A. F., 18 Morris, C., 92, 93, 98(33), 99(29, 33), 100(29), 105(29, 33) Morris, C. F., 176 Morris, H., 239 Morris, N. R., 559, 566 Morris, P.. 241

609 Morris, R. W., 131, 132(91), 184, 185(11), 190(11), 192, 201(11), 202(11) Morrison, M., 242 Morrow, J. F., 28 Morse, J. W., 214 Morse, L. S., 24 Moseman McCoy, M. I., 32, 34, 35122). 36(22), 40(6, 221, 41(6), 42(6), 46(6). 47(6), 56(6) Moses, R. E., 527, 529(89) Moss, B., 220, 225(6), 228, 230(6), 246, 248(11),249(17, 18, 20),250(17), 251(18, 19, 20). 252(18), 254(18, 23), 255(19), 256(28, 29). 260, 261(10), 262(10), 263, 265(58), 560 Mountain, 1. M., 328, 433(57) Mowbray, J. F., 359(285), 364 Muench, H. J., 578, 579(42) Mukerjee, H., 150, 151(219) Mukerji, S. K., 184, 188(17), 214(17) Mukhopadhyay, A. K., 504 Mulder, H., 398, 399(455), 400(455), 404(455) Mullenbach, G. T., 571, 572 Muller, F., 366, 367(301) Miiller, W. E., 227, 232(31), 237(30), 238(30), 240(30) Munniksma, J., 398, 400(461a) Munoz, R. F., 498 Munro, J. L., 48 Muramatsu, M., 111(26), 112, 113(26), 119(26), 127 Murdock, A. L., 375 Murray, C., 65, 67(44), 72, 76(44), 77(44) Murray, K., 6, 9(26), 47 Murray, N. E., 6, 9(26), 20(39), 47 Murthy, G. S., 336, 337(110) Murthy, P. V. N., 423 Muthukrishnan, S., 246, 247(5), 257(5), 258, 259, 260, 261(5), 262(5), 265(58) Myer. Y. P., 407 Myers, R., 148, 150(191) Myers, T. C . , 528 Nagamine, Y., 137 Nagao, K., 42, 43(44) Nagata, T., 4 Nagawa, F., 475, 478, 479(17, 27), 480(17), 482( 17) Nagle, D. P., Jr., 564, 565(31, 32)

610 Nagyvary, J., 426 Nakada, Y., 86 Nakagawa, E., 56 Nakagawa, S., 432 Nakamura, S ., 278 Nakanishi, K., 578 Nakanishi, Y., 137 Nakashima, S., 423 Nakazato, H., 227 Nakazawa, Y., 440 Nall, B. T., 390 Narang, S. A., 545 Narang, S. A,, 16, 20 Narita, K., 369, 439. 441(27), 442(27), 443(27), 444, 445(50, 52). 446(52), 457(50), 463(50, 92) Nash, H. A., 160 Nasmyth, K. A., 26 Nath, K., 16, 17(69), 69, 141 Nathans, D., 178, 179, 180 Natori. S., 137, 502 Navon, G., 342, 343(160) Nazario, M., 184, 185(12) Neff. N., 67, 70(66), 71(66, 77), 73(66, 77), 75(66), 82(66), 86(90), 93 Nehrotra, B., 84 Neilson, T., 56 Nelson, J., 79, 80(129) Neuberger, A., 410 Neumann, H.. 338 Nevins, J., 219, 220(4), 221(4), 224(4), 225(4), 226(4), 229(4), 236(4), 237(4), 238(4), 248, 262(16) Nevins, J. R., 152 Newman, I., 22 Ng, S. Y., 144, 147(166), 148, 149(197) Nicholls, D. M., 417, 418(570) Nicholson, B., 75, 761106) Nicolay, K., 366, 367(302) Nierlich, D. P., 214 Nierman, W., 67, 69, 70(66), 71(66), 73(66), 75(66, 74), 82(66), 93, 104 Niessing, J., 221, 224, 231, 232(43), 238(23) Nikiforov. V., 76, 77( 114) Nikolaev, N., 485, 486, 487, 496(17), 498, 502 Niles, E., 89, 90(19), 94(19), 95(19), 98, 99(52), 104(52) Nilsen, T. W., 298, 310(60, 61), 311 Nishihara, T., 275, 276(55)

AUTHOR INDEX Nishikawa, S., 34, 37, 38(23,31), 42,43(44), 44, 54(47), 55(47), 56(79) Nishimura, S., 83 Nishioka, Y., 148 Niu, C.-H., 346, 346(203), 354, 368(203), 374(203, 203a) Niveleau, A., 268, 270(12), 276 Nau, F., 558, 562, 564(5) Neu, H. C., 503 Niedballa, U., 542 Nishi, N., 443 Nishimura, S., 447, 557, 561. 564(3), 566, 568, 570(6), 573, 574, 575(5), 578(5, 6). 579, 580(5l, 53), 581(52). 582 Niyogi, S. K., 502 Noguchi, S., 579, 581(52) Noller, H. F., 506 Nomoto, A., 36, 496 Nornoto, M., 465 Nomura, M., 72, 74, 308, 494 Nossal, N. G., 176, 177(91), 178(99), 508, 510

Novelli, G. D., 185, 186(36), 187(36), 193(36), 194(36), 196, 197(36), 210(67) Novik, N., 363 Nowoswiat, E. F., 331 Nozu, K., 268 Nudel, U., 243, 535, 539, 548(140, 171), 549(140, 171, 2771, 550(140, 171, 277), 552(277) Nuret, P., 146 Nusser, I., 227, 237(30), 238(30), 240(30) Nusslein, C., 66 Nutter, R., 232 Oakley, J., 90, 91, 100, 103(21, 22, 27), 486 Obermeier, R., 443 O’Brien, B. M., 536 Ochoa, S., 62, 268, 518, 525, 529(71, 72), 535, 537(141), 539(3) Oda, K., 582 O’Dea, M. H., 160 Ogawa, T., 156, 159(7), 162(7), 174(7), 176 Ogdan, J., 342, 343(15) Ogden, R. C., 32 Ohasa, S., 222, 227(16), 525 Ohe, K., 184 Ohashi, Z., 447, 573 Ohe, M., 369 Ohgi, T., 579, 580(53), 581(52)

AUTHOR INDEX

61 1

Ohnishi, S., 135 Overath, H., 202 Ohnishi, Y., 502, 506, 511(17) Owen, P., 520, 522(23) Ohtaka, Y., 268 Owens, R. A , , 553 Ohtsubo, E., 494 Ozeki, H . , 64,475,479(17), 480(17), 482(17) Ohtsuka, E . , 15, 16(62), 28(62), 34, 37, 38(23,31), 42,43(44), 44,54(47), 55(47), Pace, B., 58 56179). 544 Pace, N. R., 57, 58(96), 268. 273, 475, Okada, N., 579, 580(5l,53), 581(52), 582 476(18) Okazaki, R., 4, 22 Padlan, E. A,, 345 Okazaki, T., 4, 22, 156, 159(7), 162(7), Paddock, G. V., 487, 496(16) 174(7), 176 Paetkau, V., 68, 83 Olden, K., 311 Page, J., 425 Oleson, A. E., 436 Pahler, A., 437 Olomucki, M., 327 Paik, W. K., 323 Olivera, B. M., 4, 6(11, 15), 12, 13(45), Palm, P., 63, 74(22), 75(22), 76(22), 86(22) 14(53), 15(24), 16(24), 17(15, 66), 18(11, Palmenberg, A,, 270, 274 1 3 , 19(11), 28(22, 24) Palmer, R . A., 337, 365( 119). 373( 119) Olson, M. V., 214 Palmieri, M., 415 Omori, A , , 464, 465 Pamula, Z., 418 O’Neill, M., 80 Panasenko, S. M., 8, 13 Onishi, T., 111(26), 112, 113(26), 119(26), Panayotatos, N., 100 I27 Pancha, E., 180 Oobatake, M., 442, 446(42), 456 Pandin, M., 345 Ooi, T., 387, 388, 392, 396(378, 379), 442, Panet, A . , 8, 9(34) 446(42), 456(56) Pao, C. C., 25 Oosterhuis, S., 398(465), 399,400(465), 404, Paoletti, C., 425 405(465) Paoletti, E., 246, 248(11), 249(17), 250(17), Openheimer, N. J., 447 255 Orava, M., 241 Paolozzi, L., 26 Orlandini, G., 242 Paradiso, P., 184 Orozco, E., 152 Pardue, S., 242 Orphanos, P., 81 Parente, A., 398, 399(457), 400(457), Ortiz, P. J., 68, 227, 518 412(457), 413(517), 414(517, 520), Ortwerth. B. J., 417, 418(5) 42 I(457) Osawa, S., 64 Parks, X., 323 Osborn, M., 180 Park, W., 244 Osburne, M., 106 Parson, K. A . , 51 Oshima, T., 436,437,447,448,453126). 454, Pascale, J . , 90, 103(21) 455, 456, 457 Pastan, I., 80 Osterman, H. L., 436,444,452(6), 459, 460, Patchornik, A., 340 461(53), 463(53) Patel. D. J., 330, 335(78), 346(78), 349, Oshinsky, C. K., 5, 7, 12, 13(46), 15(20), 367(78), 368(78, 214), 370(78, 214). 373(78), 374(214) l7(20), 18(20, 461, 19(20), 27(20) Osuna, C., 123 Paterson, A. R. P., 185 Otsuka, H., 447, 466(143), 467, 468(62) Paterson, M., 265 Oudet. P., 137(131), 138, 139, 142(140), Pato, M. L . , 505 I43( 140) Patrzyc, H., 324, 325(39), 334(39) Oura, H., 423 Patthy. L., 333 Ovadi, J . , 332 Paul, A. V., 176 Ovchinnikov, Yu., 74 Paul, J . , 244

612 Paule,M. R., 110, 111(115), 112, 113(13, 14, 1.9, 116(13, 14, IS), 117(13, 14, 15), 119(13-15), 123(14), 128(5), 129(82) Pauling, 61, 24 Pavlakis, G., 57 Pavlovsky, A. G., 329,335(73), 364,377(73), 406(73), 4 1373) Pawelkiewicz, J., 560 Payne, C., 263 Pays, E., 423 Pearson, M. L., 135 Peat, 1. R., 375 Peattie, D. A., 57 Pedrali-Noy, G. C. F., 10 Pedrini, A. M., 10 Peebles, C. L., 32 Peeters-Joris, G., 425 Peggion, E., 346, 388 Penhoet, E. E., 571, 572 Pelham, H. R., 149 Pelichova, H., 362 Pelle, E. G., 479, 495 Pellicer, A., 224, 226(24), 228(24) Penman, M . , 23 1 Penman, S., 231, 498 Pergolizzi, R. G., 579 Perham, R. N., 421 Perlman, D., 179 Perna, P. J., 128, 129(82) Pero, J., 63(24,32), 64,66(32), 76(32), 78(32, 119, 120), 79(32. 119), 80(121, 127, 129, 130), 83(24, 32), 88, 89, 105 Perrin, D. D., 375 Pesce, A , , 107 Peterkin, P. I . , 520 Peterkofsky, A., 575, 577 Peterlin, B. M., 176 Petersheim, M., 332 Peterson, E., 546 Peterson, R. L., 273 Peterson, T. C., 284, 285(23), 302(23), 304(237, 306(23) Petranyi, P., 130, 132(86) Petre, J., 268, 270(1 I ) Petsko, G. A., 365 Pettersson, U., 547 Pettit, N. M., 436 Pfiugfelder, G., 123 F'flumm, M. N., 384, 421 Phelan, J. J., 398, 400(453), 406(453)

AUTHOR INDEX Philipp, M., 543 Philips, G. R., 502, 507(11) Philips, J. H., 566 Philipson, L., 231, 547 Phillipp, M., 56 Phillips, D. C., 397 Phillips, G. R., 185(40), 186(35), 187(35,40), 188(35, 40), 189(35, 40). 191, 192(35, 52), 194(40, 52), 196(40), 197(52), 199(35), 200(35), 201(52), 202(52, 79), 203(79), 207(79) Phillips, W. D., 386 Picard, B., 149 Pietrzak, M., 184, 185(41), 186, 187(41), 189(41), 190(41), 192, 193(56), 194(56), 195(56), 199(13) Pigiet, J., 179 Pignero, A., 411, 414(500) Pilz, I., 75, 76(107) Pincus, M., 331, 332 Piperno, J., 176 Pirotte, M., 416 Piskorz, C., 577 Pitot, H., 497 Planta, R. J., 119, 138(52) Plapp, B. V., 327,329,330(70), 331,335(69) Platt, T., 63(31), 64, 83(31), 86(31) Plautz, G., 476, 483(24) Plotch, S . J., 259, 260, 265(56), 464 Plummer, T. H., Jr., 328, 407, 408(66, 4871, 410(66) Poblete, P., 184, 185(10), 186(10), 187(10), 189(10), 190(10), 194(10), 196(10), 201(10) Pochon, F., 540, 542 Podder, S. K., 464 Polder, L., 171 Polke, C., 168 Pollack, Y.,268, 270(11) Pollard, D. R., 426 Polovnikova, I., 74 Pomerai, D. I., 140 Pongs, O., 453, 455, 457, 458, 462, 463(81) Ponnuswamy, P. K., 385, 406 Porcelli, G., 335, 347(106), 350(106), 35 I( 106) Portier, C., 521, 522(41), 523(58), 524(62), 526(41), 527(80), 531(59, 80), 536, 537(62), 583(155) Post, L., 74, 494

AUTHOR lNDEX

613

Potter, V . R., 185, 424 Potts, J. T., Jr.. 346, 358, 388(197) Poucher, J. S . , 532. 5 3 3 122) Powers, D. M., 577 Preiss, J., 185, 188(29), 211(29), 212(29) Preston, J . F., 146 Pribnow, D., 491 Price, A . , 105 Price, R. P., 498 Pridgen, C., 551 Primakoff, P., 477,479(26). 480(26), 482(26), 502, SIl(15) Pringle, J. R.. 121 Privalov, P. L., 386 Prochiantz, A , , 200 Proudfoot. N., 228 Przybyla, A. E . , 361. 427(274) Ptashne, M., 20, 12 Ptitsyn, 0. B., 365, 385, 386 Puett, D., 340, 356, 357, 411, 421(143) Puigdomenech, P., 323 Pulkrabek, P., 212 Pyle, V. S . , 44, 54(45)

L., 284, 285(18, 20), 286(20), 296(20), 304(18), 306, 307(86) Raue, H. A . , 537, 538(158) Ravetch, J. V . , 169 Ray, R. K., 503 Raziuddin, Chateryi, D., 506 Reali, N ., 242 Recchia, J., 333 Rech, J . , 498, 499(68, 69) Record, M. T., Jr., 134 Reddi, K . K., 503 Reddy, R., 570 Redfield, R. R., 425 Reese, C. B., 292 Regnier. Ph.. 527 Reha-Krantz, L . , 161 Reichard, P., 178, 179, 547, 548(270) Rein, R., 432 Reinberg, D., 156, 157(11), 158(11), 159(11), 160(11), 166(11), 173 Reiner, A. M., 537, 538(151), 539(152) Reinhold, V. N., 410 Reisinger, D. M . , 305, 309(84) Reiter, T. R., 181

Quadritoglio, F., 346. 388 Quattrone, A . J., 335, 347(106), 350(106),

Rekosh, R. M. K., 180 Renart, J., 123 Renugopalakrishnan, V . , 432 Rensing, U., 273, 275, 279150) Resnick, H., 338 Retel, J., 119, 138(52), 144 Rether, B., 185, 186(38), 187(38), 188(38),

35 1( 106)

Quigley, G. J., 195 Raae, A. J., 8, 9(34), 14, 16(59), 17(59), 19(59), 21(59)

Raap, J., 347, 348(21 I), 349(206), 350 Rabin, B. R., 418, 425(578), 462 Rabinowitz, J. C.. 65, 67(44), 72, 76(44), 77(44), 558, 564(8), 565(31, 32)

Rdbinowitz, M.. 150. 151(216), 540 Rabussay, D., 80. 88 Rackwitz, H. R., 145 Raetz, C. R. H.. 320, 323(23), 423(23) RajBhandary, U . L., 28, 57, 463 Rake, A. V . , 202 Ralston, G. B., 428 Ramanarayanan, M., 538, 539(170) Ranson. L., 498 Rapoport, S.. 458 Rashba, H., 244 Rashid, M. A . , 466( l43), 467 Rathinasamy. T. K . , 343 Ratliff, R., 84

Ratner.

189(38), 190(38), 194(38), 197(38), 201(38), 202(38), 207(38), 210, 21 1(98), 21398) Reuter, W., 546 Revel, M., 259, 282, 284, 285(21), 287. 290, 298, 301(45), 302(65), 304(59, 65), 305(59, 65, 751, 306(65), 307(59, 65). 308(59), 309(83), 310(45), 535, 548(140), 549(140), 550(140) Reverman, L. F., 545 Reychler, H., 425 Reynolds, J. H., 326 Reynolds, W. F., 375 Rezelman, G., 423, 424 Rhides, C., 180 Rhodes, D., 246, 262(13) Rhodes, G., 63(33), 64, 70, 86(33), 104 Ric, S . , 538 Rich, A , . 195, 540

614 Richards, F. M., 318, 320, 322, 323(1), 326(1), 327(1), 329, 335(1), 336(1), 337(1), 339(1), 340(1), 342(1), 343(1), 344, 345, 346(1), 347(1), 349(1, 16). 351(1), 352(202), 353, 354, 355(202), 356(1, 118), 358(1), 359(1), 360(1), 364(1, 16). 365(1), 368(1), 372, 373(1), 382(1), 384(1), 388(185), 390(1), 391(226), 404(1, 16), 415(1, 16), 418, 420(1, 16), 424(1), 425, 426(1), 429(1), 431, 432(1) Richards, 0. C., 152 Richardson, C. C., 4, 6(10), 7, 8(29), 9(29), 12, 13(47, 48). 14(54), W I O , 23), 16(29, 54, 63). 17(10,29, 48,50,54,63), 18(29, 48), 19(29, 48, 63), 21. 22, 23, 25(63), 27(29, 48, 60, 90), 28(54, 129), 51, 55(69), 174, 175(76-80) Richardson, D. C., 365 Richardson, D. I., Jr., 430, 431(643) Richardson, J. P., 75, 549 Richardson, J. S . , 365 Richardson, R. W . , 44, 54(45) Richter, G., 111(25), 112, 113(25), 117, I19(46), 123(25, 46). 124(46). 137, 520 Riehm, J. P., 335, 395, 396(423) Rigby, P., 180 Riggs, A. D., 20, 52 Riley, P., 343 Ring, J., 91, 92(26, 28), 98(26), 103(26, 28). 104(26, 28). 105(26) Rio, D., 148, 150(191) Riordan, J. F., 334, 340, 341(146), 342(146), 384(148) Riquelrne, P., 320, 323 Risi, S . , 343 Riva, S., 64, 147 Robbi, M., 423 Robbins, A., 148, 150(191) Roberts, B. E., 552 Roberts, J., 63(27), 64, 83(27), 86(27), 477, 479(26), 480(26), 482(26), 502, 51 I ( 15) Roberts, G. C. K., 366, 367(300), 369(300), 370, 372(315), 373(315), 376(300), 377(300), 379(300), 380, 381(300), 386(3 12), 387(3 13), 389(3 12, 3 l3), 3933121, 396(312, 427), 431, 432(647), 462 Roberts, W. K., 282, 291(7) Robertson, H. D., 259,464,471,472(6), 475,

AUTHOR INDEX 479, 485, 486(4), 487(4), 489(6), 490(6), 494(8), 495(6), 496(16), 499 Robinson, A. J., 180 Robinson, W. S., 145 Rocchi, R., 329, 335, 346, 347, 350( l04), 351(104, 220), 356, 388 Ro-Choi, T. S., 570 Rodeh, R., 559 Rodriguez, R., 64 Rodriguez, R. L., 9, 20(40) Roe, F., 220, 233, 238, 239(71) Roeder, R. G., 110, 112(1, 29). 112, 113(29), 115, 116(40), 119, 124, 127(54, 71), 137(1), 144, 145(1), 147(38, 166), 148, 149(197), 230(51), 232, 262, 263(65) Rogall, G., 150, 151(215) Roget, A., I50 Rohde, W., 145 Rojder, G. C., 148 Rokugawa, K., 519, 525(20) Roller, A., 525 Romano, L., 174, 175(79. 80) Romeo, J. M., 564, 565 Rorner, W., 269, 270(20), 272(20), 273(20), 276(20) Rornovacek, H., 443 Ronda, G. J., 358, 429 Roosemont, J. R., 332 Rosa, M., 100, 101 Rosbash, M., 231 Rose, K., 218, 228(3), 221, 222(13), 223(13), 225(13), 226(13), 230(13), 231(13), 232(3, 13, 42). 233(13), 237(13), 238, 239(71), 242 Rose, J. J., 353 Rose, J. K., 259 Rose, K. M., 1’36 Rosemond-Hornbeak, R., 255 Rosen, H., 282 Rosen, L., 537, 538(153) Rosenbaum, N . N., 575, 576(33), 577(33) Rosenberg, A., 93, 98(33), 99(33), 105(33) Rosenberg, J., 20 Rosenberg, M., 63(29), 64, 80(29), 83(29), 86(29), 98, IOO(48). 101(48), 102(48), 105(48), 265, 486, 491, 493(7) Rosenberg, R., 242 Rosenblurn, E., 220, 225(6), 228, 230(6) Rosenthal, A. L., 426, 489 Ross, A., 506

AUTHOR INDEX Ross, C. A., 462 Roth. J. R., 9 Roth, J. S., 416, 417, 418(547) Rothberg, P. G., 36 Rothman-Denes, L., 88, 107(14, 15) Rothrock. J . W . , 546 Rottman, F., 23 I , 542, 545 Roulland-Dussoix, D., 100 Rowe, M. J., 464 Rowen, L., 156. 157(9), 158(9, 10) Rowlands, D. J . , 553 Roy, A. K., 548, 549(279) Rubin, B. Y.. 287, 289(36) Rubin, G. M.. 570 Rubin, J . R., 464 Riibsamen, H., 428, 431(631), 432(631) Ruch, P., 232 Ruegg, K. J., 275 Ruet, A., 127, 135(77), 521, 527(51) Ruger, A., 76, 77(118) Ruger, W., 76, 77(118) Rundell, K., 180 Runnels, J., 49, 50(62), 51(65) Rupley, J . A., 387, 395, 396(377, 378) Rupprecht, Hecht, S., 259 Rushlow, K., 152 Russchen, F., 398, 400(458) Russell, R. L., 577 Russell, W. G., 180 Riiterjans, H., 329, 330(74), 335(74, 77), 346(74, 771, 366. 369(77), 372(77), 373(74, 77), 374(74), 375, 376, 377(74), 379(77, 237). 380, 436, 437, 444(7), 457, 46 I Riiterjans, H. H., 355, 366(237), 367(237), 368(237). 369(237). 370(237), 373(237), 376(306), 377(306), 379(306), 38 1(306), 386(237) Rutter. W., 227, 241 Rutter, W. J . , 115, 116(40), 119, 126(51), 128, 132(91). 135, 136(112), 144, 152, 192, 214, 361, 427(274) Ryan, M. J., 28 Ryan, T . , 97 Rychlik, I., 212

Saari, B.. 271. 273(29) Sabo. D., 279 Saccone, C., 239

615 Sacharovsky, V. G., 329, 335(71, 72). 369(71), 376(72), 377(72), 379(71, 72). 381(72), 382(72) Sachs, D. H., 373, 393 Sachs, L., 241 Sadler, P. J., 389 Saenger, W., 437 Sagar, A . D., 548, 551(278) Saitoh, T., 76, 77(113), 81, 82(142) Sakabe. K.. 4, 22 Sakaguchi, K., 519, 521(18), 522(18), 523(18), 524(18), 539, 543, 544(226) Sakai, T. T., 195 Sakakibara, Y . , 173, 174(68) Sakano, H., 471, 475(5), 478(5), 479(5,17, 271, 480(5, 171, 482(5, 17) Sakiyama, F., 369, 439, 441(27), 442(27), 443(27) Sakonju, S . , 148 Salas, J., 224, 226(24), 228(24) Salas, M., 224, 226(24) Salditt, M., 498 Salditt-Georgrieff, M., 248, 262( 15) Salganik, R. I., 433 Salomon, R., 535, 546, 548(140, 263). 549(140, 262, 263, 274), 550(140), 551(262), 552(263) Salser, W., 22 Salton, M. R. J., 520, 522(23) Salvo, R., 93, 98(36), 103, 104(71) Salzman, N. P., 179 Samanta, H., 287, 288(38), 289(38), 290(38), 292(38), 294(38), 299(47), 300(38, 47), 301(38, 47), 303(47), 304(38), 305(38, 76), 306(76) Samartsev, M., 346 Samejima, T., 452 Samuel, C. E., 284, 285(23), 302(23), 304(23), 306(23) Sanchez-Anzaldo, F., 65, 132 Sandeen, G., 415 Sander, C., 456 Sandig, L., 564 Sandoval, A,, 358 Saneyoshi, M . , 452 Sanger, F., 463 Sanger, H . L., 145 Sano, H., 16, 227, 231(27), 233(27), 234(27), 235(27), 274 Santer. M., 244

616 Santoro, J., 329, 330(74), 335(74), 346(74), 373, 374, 377 Saragosti, S., 139, 143(142) Sarda, L., 410 Sarkar, P., 90, 93(24), 98(24), 100, lOl(24, 67), 102(67) Sarin, P. S., 145, 147(173), 148(173) Sarrna, R.. 259 Sarris, R., 91, 103(27) Sasaki, A., 466(143), 467 Sasaki, Y., 111(23, 24), 112, 113(23, 24). 116(23, 24). 119(23) Sasse, L., 540 Sassone-Corsi, P., 150 Sato, K., 435, 447(1) Sato, S., 449, 450(72), 452, 457, 465, 582 Sato-Asano, K., 464 Sauer, H. W., 120 Saunders, C. A., 543, 549(227) Saunders, G. F., 397 Saunders, G. R., 144 Sawada, F., 323, 344, 426,452 Sawadogo, M., 137, 144 Sawicki, S . , 236(69), 238 Saxena, V. P., 391 Sbordone, L., 582 Scaife, J., 63, 74(19) Scatturin, A., 350 Schachner, M., 65 Schafer, K., 240 Schafer, K. P., 150, 151(217), 577 Schaffer, S. W., 319,391(406), 392,413(405) Schaffner, W., 275 Schaller, H., 86, 168 Schattenkerk, C., 345, 348(213), 349, 350(213) Schechter, A. N., 355, 368, 369(238), 373(238), 377(305), 379(305), 380(305), 381(305), 382(305), 393 Schedl, P., 477, 479(26), 480(26), 482(26), 502, 511(15) Scheele, G., 423, 424 Scheffer, A. J., 340, 398(464), 399(139), 400(139, 460, 463). 404(139), 407, 409(139, 486), 410(486) Schemer, I. E., 6, 28(22) Scheit, K. H., 63(35), 64,86(35), 464, 521, 522(48), 540, 541, 542 Schekman, R., 156, 168(5) Schellman, J. A., 383, 386

AUTHOR INDEX Scheraga, H. A., 82, 335, 338, 339, 340, 344, 345(184), 355, 362, 363, 366(237), 367(237), 368(237), 369(237), 370(237), 373(237), 379(237), 385(123), 386(237), 387, 388(184, 186), 392, 393(388), 394(279, 367, 388), 395(361), 396(186, 367, 370, 372, 377, 378, 379, 381, 388, 422, 423). 406, 429 Scherzinger, E., 173, 175 Schetters, H., 542 Schibler, U., 150 Schimke, R. T., 284, 285(17), 286( 17), 287( 17) Schimmel, P. R., 320, 322(16a), 470 Schito, G., 88, 107 Schleich, T., 278 Schlagman, S., 160, 171(21) Schleif, R., 75 Schlessinger, D., 485,486,498,502,507(10), 508(10), 509(10), 510(5), 51 l(12, 17), 512, 525 Schlimme, E., 185, 186(34), 187(34), 189(34), 190(34), 191(34), 213 Schmid, F. X., 355, 390(399), 391(244) Schmidt, A., 282, 287, 290, 298, 301(45), 302(65), 304(59, 65), 305(59, 65, 75). 306(65), 307(59, 65), 308(59), 310(45) Schmidt, F. J., 214, 215,480, 482(32), 551 Schmidt, O., 214 Schmidt, W., 558, 564(9), 565(9) Schmukler, M., 427 Schnabel, J. J., 561 Schoenmakers, J. G. G., 417, 423(558) Schofield, P., 184, 185(39), 186, 187(39), 188(39), 1 89(39), I90(39), 19I(39), 192(39), 194(71), 196, 198(71), 201(71) Scholand, J., 359(285), 364 Scholtissek, C., 416 Schray, K. J., 319 Schreier, A. A., 352, 354 Schroeder, C., 88 Schroder, F. P., 363 Schroder, H., 240 Schultz, R. M., 343, 344(168) Schulz, H. H., 461 Schuman, S., 108 Schuster, L., 503 Schwartz, L. B., 124, 127(71) Schwartz, M., 410 Schweiger, M., 491

AUTHOR INDEX Schweppe, J. S . , 136 Scoffone, E., 335, 337, 344, 346, 347, 350(104), 351(104, 220), 356(117, 356, 388) Scofield, R. E., 320 Scopes, D. I. C., 540 Scott, A., 244 Scott, J. F., 170, 184, 185(7) Scott, R. A., 386, 396(370) Scragg, A. H., 150, 151(212) Scrutton, M. C., 132 Seagle, R. L., 341, 342, 384 Sebastian, J., 123 Sebring, E. D., 179 Secemski, 1. I., 427 See, Y. P., 520 Segall, J., 115, 144, 147(38, 166). 149, 262, 263(65) Sehgal, P. B., 313, 548, 551(278, 280) Sehon, A. H., 363 Seibert, G., 227, 232(31) Seidman, J. G., 214, 480, 482(32) Seidman, S., 126, 139(73), 139, 143(73, 146, 147) Seifert, W . , 65 Sekiguchi, M . , 502, 538 Sekimizu, K., 137 Sekiya, T., 464, 483 Sela, I., 546, 548, 549(262, 274), 551(262) Sela, M., 338, 363, 391, 540 Seliger, H., 56, 533, 543, 544 Sells, B. H., 423 Selzer, G., 160 Sen, G. C., 282, 284, 28% 18, 20). 286(20), 296(20), 304(18), 306, 307(86) Senear, A., 276 Sentenac, A., 117, 119, 120(47), 123(47), 126, 127(47), 128, 134(79), 135(77), 136(113), 137, 138, 139, 140, 143(149, 150), 144(149. !SO), 145(149), 521, 527(51) Sethi, S., 220, 225(8), 233(8) Sethi, S . K . , 580, 581(54), 582(54) Setlow, R., 344 Setondji, J., 528 Sgaramella, V., 16, 19,20,21(82,87), 22,28 Shabarova, Z. A., 37 Shafferman, A., 343 Shaila, S . , 282, 304, 306 Shapka, R., 342, 343(161)

617 Shapiro, L., 67, 273 Sharp, P., 263 Sharp, P. A., 147, 148 Shastry, B. S., 148, 149(197) Shatkin, A., 246, 247(5), 257(5), 258(2), 259, 260(54), 261(5), 262(5), 263 Shaw, P. A., 144 Shemyakin, M. F., 494 Shemyakin, M. R., 9 Shenk, T., 180 Sherwood, L. M., 346, 388(197) Shibata, H., 570 Shimada, I., 444, 445(51), 446(51) Shimura, Y., 471, 4 7 3 3 , 478(5), 479(5, 17, 27), 480(5, 17), 482(5, 17) Shimizu, N., 284, 285(26) Shindo, H., 330, 335(79), 339, 346(79), 347(203), 354, 355, 366, 367(297), 368(203), 369, 370(79, 297), 372(135), 373(79, 2391, 374(203, 203a), 379(239), 388(329), 418(79) Shindo-Okada, N . , 579, 580(53), 581(52), 582 Shine, J., 16, 20(67), 21(67), 47 Shiobara, Y . , 530 Shipp, S., 445 Shliapnikov, S. V., 323, 329, 335(71), 369(711, 379(7I) Shlomai, J., 160, 161, 165(26), 170, 171 Shockman, G. D., 302, 303(69) Shorenstein, R., 76, 77(111) Shortle, D., 180 Shortman, K., 416, 417, 418, 423 Shrager, R. I., 373 Shriver, K . K . , 426 Shugar, D., 318,426,534,541,543198, 199) Shulman, E., 182 Shulman, L., 282, 287, 298, 301, 302(65), 304(59,65), 305(59,65), 306(65), 307(59, 65), 308(59) Shum, B. W . - K . , 542 Shuman, S., 152, 248, 249(22), 250(22), 251(22), 252(22, 33), 255(22), 261(33), 262, 263(22), 558, 568 Shure, H., 305, 308, 309(83) Shuster, R., 308 Sidikaro, J., 308 Siebenlist, U., 63(26), 64, 76(26), 83(26), 86(26), 98, 100(47), 102(47), 103(47), 134 Siegel, R., 105

618 Siegel, R. B., 506 Sierakowska, H., 318, 426 Silber, R., 31, 32(1, 2), 33(1, 2). 34(12), 35(12), 36(12), 40(12), 41(1, 12), 42(2, 12), 45(2), 47(2, 12) Silberklang, M., 463 Silberstein, A . , 538 Silengo, L., 485, 486 Silver, L. L., 176, 177(91) Silverman, D. N., 355 Silverman, P. M., 275 Silverman, R. H., 292, 293(53), 296(53), 297(53), 298(53), 305(53), 306(53), 307(53), 312(53) Silverman, S., 214 Silvestri, L. G., 64, 147 Simms, E., 68 Simon, L., 268 Simon, L. N., 528 Simon, M., 99 Simonesits, A,, 57, 464 Simontov, R., 241 Simpson, R., 98, lOO(47). 102(47), 103(47) Simpson, R. B., 63(26), 64, 76(26), 83(26), 86(26) Sims, J., 159, 161(15), 162(39), 165(15, 3942). 166(15, 39, 40), 167(39) Simuth, J., 542 Singer, C. E., 571 Singer, M. F., 502, 503, 507, 508(9), 5 10, 519, 521(13), 522(43), 523(43), 525, 526, 527, 529(73, 89), 530(82, 90). 531(86), 532(114), 533(82, 86). 534, 535(82, 112, 114, 120), 536(82, 114), 537(86, 141), 539(106), 542, 545(131), 547(82, 112) Singh, R., 579, 581(45, 46) Sinha, N. K., 176 Sinsheimer, R., 86 Sinsheimer, R. L., 4, 28 Sippel, A , , 68, 221, 222(10), 225(10), 229(10), 231(10), 233(10), 243(10) Sips, H. J., 398, 400(446), 405(446), 409(446), 42 l(446) Sirothen, K., 50, 51(64, 65) Sittert, O., 390 Sjolin, L., 366 Skehel, J. J., 307 Sklar, V. E. F., 119, 124, 127(54, 71) Skoda, J., 528 Skup, D., 264

AUTHOR INDEX Slater, D., 237, 238, 241(67) Slater, I., 237, 238, 241(67) Slater, J. P., 131 Slattery, E., 304, 305(76), 306(76), 307(86) Slobin, L. I., 268, 270(12) Sloots, B., 398, 400(447), 405(447) Slor, H., 278, 279 Smallcombe, S. H., 355 Smart, J. E., 181 Smellie, R. M. S., 520 Smirnov, Yu., 74 Smith, A., 180 Smith, A. G., 425 Smith, D., 84, 277 Smith, E. L., 333, 403 Smith, G. K., 319, 413 Smith, G. R., 571 Smith, H. J., 151 Smith, J. C., 521, 527(49), 546 Smith, J. D., 213, 471, 472(6), 475 Smith, M., 545 Smith, M. A,, 464 Smith, S. S., 111(18), 112, 113(18), 116(18), 120, 122, 123(18, 55) Smrt, J., 528, 534(97) Smyth, D. G., 320,321(15), 339(15), 400(15), 420( 15) Snechkute, M. A . , 36, 37(25) Sninsky, J. J., 37, 38(30), 39(30), 55(30), 537, 544(150) Snopek, T. J., 9, 32, 34(8), 35(21), 36(18), 37(5), 3801, 39(5). 40(5). 41(5), 42(5), 47( 18). 48(8), 49(8) Snustad, D. P., 51 Snyder, L., 49, 50(62), 51(64) Snyder, L. R., 50, 51(65) Sobell, H. M., 540, 541(189) Soderhall, S., 4, 10(16), 17(16), 18, 19(16, 80) Soderman, G., 179 Soe, G., 519, 521(19), 523(19). 525(19) Sogin, S. J., 543, 549(227) Sokawa, Y., 284, 285(26, 27) Sokolovsky, M., 340, 341, 342(146), 384( 148) So11, D., 214, 470, 475, 543, 547(225), 557, 558, 559, 564(6), 566(6), 568, 571, 572(4), 574(1), 575(4), 576(30, 32), 577(32), 578(4) Soll, L., 582

AUTHOR INDEX Soltis, D., 49, SO(62). 55 Somberg, E., 249 Someno, K., 544 Somers, D. G., 135 Sommer, H., 540 Sommer, R., 168 Sommer, R. G., 545 Sonenberg, M., 395 Sonenberg, N., 259 Sonenshein, A., 106 Sonnenbichler, J., 123 Sopori, M. L., 282 Soreq, H . , 243, 521, 522(42), 523(42), 524(42), 52342). 529(42). 53.5, 539, 546(42), 547. 548(140, 171, 172. 263. 264, 2681, 549( 140, I7 I , 262, 263, 264, 274, 276, 2771, 550(42. 140. 171, 172, 264, 277), 551(262, 264, 278, 280). 552(263, 264. 277, 278) SBrm, F., 464, 528 Sormova, Z . , 528 Sorrentino, S . , 414, 415, 429(540) Southern, E. M., 73 Spadari, S . , 10 Spahr. P. F., 496, 502,503. 504(30), 505(30), 507(8), 510(5), 512, 513(23). 514 Spelzhaus, A , , 205, 564 Spencer, E., 108, 149, 152, 248, 251. 262, 263 Spengler, S. J . , 570, 571(15). 580(15), 5821 15) Sperling, J., 344 Sperling, R., 338. 339(130), 344 Speyer. J. F., 25 Spiegelman, G., 77.78(122), 79.80(122, 131) Spiegelman. S.. 268, 273, 274(4), 275, 276(55), 279 Spierings, T.. 146 Spindler,S.R.. 111(15),112, 113(13, 14, 15), 116(13. 14, IS). 117(13. 14, IS), 119(13IS), 123(14), 137 Spoor, T. C.. 326 Sprague, K. U., 147 Spremulli. L. L., 561 Springgate, C. F.. 131(93), 132 Springhorn, S. S., 448 Spnnzl, M., 184, 190(54), 191, 192(54), 194(54), 196, 197(54), 200, 205(97). 215(3), 545, 564 Squires. C., 74

619 Squires, C. L.. 74 Srinivasan, P. R., 538, 539(170), 559 Sridhara, S., 111(19), 112, 113(19), 117(19), 123(19) Staehelin, M., 538, 539(169) Stahl, D. A., 57, 58(96), 506 Stahl, S . , 70, 94, 95(38) Stalter. K., I14 Stamfer, M., 88, 105(7) Stark, B. C.. 473, 474(l I , 12). 475 Stark, G. R., 284, 285(17), 286(17), 287(17), 336, 337(116), 347(116). 356(116) Staros, J . V., 344, 388(185) Starr, J. L., 185, 191, 196(50), 197(50), 207(50) Staudenbauer, W. L., 173 Stawinski, J., 20 Steczko, J., 331 Stefanos, S., 286 Steffen, R.. 240 Stein, G., 231, 244, 342, 343(160, 161, 165, 166) Stein, J . , 244 Stein, W. H . , 320, 321(15), 339(15), 341, 400(15). 413. 420(15), 421, 425, 437, 440(23), 462(23) Steinberg, C. M., 21, 176 Steinberg, I. Z., 338, 339(130), 344(130) Steiner, D. F., 397 Steiner, R. F.. 519, 528, 530(100) Steinschneider, A., 265 Steitz, J. A . , 72, 244, 276, 485, 486, 491(10, I I ) , 493(11), 494(11), 495(11), 502, 513(23) Stellwagen, E., 132 Stender, W., 75 Stephenson, G. P., 540 Stephenson, M. L., 184, 185(7) Stern, R., 426 Sternbach, H . , 65. 185, 186(34), 187(34), 189(34), 190(54), 191(34), 192(54, 55). 194(54), 196, 197(54), 198, 545 Sternglanz, R., 176 Stetler, DIA , , 136 Stetter. K.. 65. 67(47), 76, 77(47, 1 15, 116). 82(47) Stevens, A , , 62 Stevenson. K. J.. 319 Stewart, G . R., 319 Stewart, M., 496

620 Stewart, W. E., 11, 284,286(16), 312(16) Stewart, W. E., 312 Stillmann, B. W., 181 Stinchcomb, D., 66 Stob, S., 330, 335(80), 339(80), 346(80),

367(80), 370(80), 371(80), 372(80), 372(80), 377(80), 378(80), 386(80), 389(80), 395(80) Stollar, B. D., 540 Stols, A. L. H., 424 Storey, H.T., 443 Stratford, I. J., 546 Straub, F. B., 397 Streaty, R. A , , 405 Strehlke, P.,542 Strickland, E. H., 382,383(336),384(342) Stringfellow, L. E., 271,273(30), 278(30) Strominger, J. L., 503 Strothkamp, R.,90,91,103(22,27) Strycharz, G.D., 494 Studencki, A., 56 Studier, F., 93,98(33), 99(33), 100(53), 101, 105(33,56) Studier, F. W., 10, 23,485,486(1), 489(1), 491(5, 12),493(5), 495 Stukacheva, E. A., 494 Stunnenherg, H. G., lll(20, 21), 112, 113120,21), 116(20), 117(20), 146 Sturtevant, J. M.,346,352(202), 355(202), 388 Sueoka, N.,4 Sugano, H.,424 Sugden, B., 1 1 1(28), 112, 113(28), 148 Sugimoto, K.,4,22 Sugino, A., 4,9,16,20(67),21(67), 22,32, 34, 35(21), 36(18), 37(5), 38(5), 39(5), 40(5), 41(5),42(5), 47(12) Sugino, Y., 80 Sugiura, M., 34,37,38(23, 31). 42,43(44), 55, 56(79)

Sugrue, S., 156 Suhadolnik, R.J., 302,303(69) Suito. F., 414 Sumegi. J., 139,144(148) Sumidar Yasumoto, C., 156, 170 Summers, W., 89,90(19),94(19), 95(19), I05 Sumper, M.,273,274 Sundherg, R.J., 332 Surks, M.,248, 249(22), 250(22), 251(22),

252(22), 255(22), 263(22)

AUTHOR INDEX Suryanarayana, T., 490,506 Surzycki, S. J., 111(16), 112, 113(16),

116(16), 126,139(73), 143(73, 146,147) Surzycki, J. A., 139,143(146) Susman, M., 21,176 Sutcliffe, J. G., 71 Suzuki, H., 398, 399(457),

400(457),

412(457), 42l(457) Suzuki, M., 34,38(23) Suzuki, T., 362 Suzuki, Y., 417 Sverdlov, E., 74 Swan, R. J., 426 Swetly, P.,240,241(76) Swierkowski, M., 534 Swift, T. J., 112 Synder, A. L., 497 Szafranski, P., 551 Szekely, M., 463,490 Szer, W., 541 Szeto, K.S., 543 Szkopinska, A , , 551 Szurmak, B., 136 Szybalski, W., 494 Tabak, H. F., 168 Tabor, M.W.,57 Tabor, S., 175 Tait, R. C., 9,20(40) Takagi, Y., 185 Takahashi, K., 333, 334(99), 433,436,437,

438, 439(3), 440(3, 23). 441(3, 24), 442(3), 444(36,37). 445(20, 51), 446(40, 511, 447(20, 23, 31), 449,450(26, 3 3 , 451.452(3,29,453(25, 26,74), 462(23, 80), 463(25,36,37,74). 464(65), 466( 14, 142),467 Takahashi, S . , 81, 82( l42), 388, 392,442, 446(42), 456(56) Takahashi, Y., 417,424 Takai, N ., 465 Takanami, M., 202 Taketo, M., 76,77( 113) Takeya, T., 483 Tal, J., 215 Tal, M.,538 Talkington, C., 76,78(121), 80(121) Talkington, C. A , , 148 Tamaoki, H.,439,441,442(27),443(27) Tamire, A., 131

AUTHOR INDEX Tamburno, A. M.,337, 338, 350, 356(117), 388( 127) Tamiya, N., 464, 465 Tamm, I., 548 Tanaka, K . , 465 Tanaka, S . , 5 5 , 56(79), 544 Tanaka, T., 56 Taniguchi, T.. 80, 279, 414 Taniuchi, H., 340. 356, 357(142), 359, 394(142),421(142, 263) Taniyama, Y . , 56 Tararskaya, R. I . , 449, 452(70), 467(70) Tarasov, A. P., 520 Tarnowski, G. S.. 328, 433(57) Tarrago-Litvak. L., 200 Tatarskaya, R. I . , 465 Tavale, S. S . . 540, 541(189) Tavitian, A , , 497 Taya, Y.,566, 574 Taylor, A., 75 Taylor, E. K., 564 Taylor, A. L., 489 Taylor, G. T., 355 Taylor, H. C., 345, 347, 365 Taylor, M. M., 273 Taylor, M. W., 116, 135 Taylor, T., 343 Tazawa, S., 545 Tazawa, I., 541, 545 Teeter, M. M., 195 Tegtmeyer, P., 180 Teh, J. S., 366, 367(298),370(298) Teissere, M.. I l l , 113(10), 116(10),117(10) Tener, G. M., 202, 292 Teoule, R., 150 Terada, S., 436, 443(9) Terao, T., 28, 407, 408(490), 450 Terenna, B., 454, 455 Testa, D., 259 Teuber, M., 503 Thach, R. E., 464, 542 Thang, D. C., 521, 522, 523, 527(50, 51) Thang, M. N., 287, 292(41), 301(41), 302(41), 303(41, 681, 519, 521, 522(14, 4 I ), 523, 524, 525, 526(41, 50, 5 1 ), 527(79),530(83,87),531(60,84), 532(67, 84). 535(83), 536(83, 84). 537, 539(67), 540, 546( 117) Thatch. R. E.. 54 Thayer, D. W., 519

62 1 Thelander, L., 158 Thi, L. L., 331, 332(86) Thiebe, R., 202, 578, 579(42) Thomas, G., 202 Thomassen, M. J . , 184 Thompson, A., 12, 13(48), 17(48), 18(48), 19(48),27(48) Thompson, J. C., 380 Thompson, S. T., 132 Thoren, M. M., 179 Thrall, C., 244 Tigges, M. A., 51 Timasheff, S., 382 Timchenko, A. A., 365 Tinoco, I., Jr., 52, 464 Wan, R., 66, 76, 78(119), 79, 80(I 19, 130). 148, 149, 150(191), 180 Tobien, M., 76, 77(115) Tocchini-Valentini, G. P., 147 Todd, A , , 540 Todling, G., 56, 543 Tokunaga, T., 56 Tolbert, G., 502, 507, 508(9) Tomasz, J . , 246, 24701, 257(5), 261(5), 262(5) Tomizawa, J.-I., 4, 160, 173, 174(68) Tomkiewicz, C., 88, 89(11), 97, 102(11) Tomlinson, G., 375 Toniolo, C., 347 Torrence, P. F., 288,541,542,545(200,217) Torri, K ., 323 Totsuka, A., 227, 237(30), 238(30), 240(30) Touw, J., 416, 423(555) Tovell, D. R., 282 Towle, H., 88, 93(10), 98(10), 102(10), 231 Trachsel, H., 259 Tramontano, D., 432 Tranquilla, T., 475 Travers, A. A., 62, 63, 71(8), 74(8), 75(8), 77(8), 79(17) Trip, E. M., 545 Trotter, C. D., 491 Troutt, A., 57 Tsai, M. J., 150, 151(213) Tseng, B., 179 Tsernoglou, D., 359, 364 Tsiapalis, C., 221, 222(12), 223(12), 224(12), 225(12), 226(12), 227(12), 229(12, 15), 230(12), 231(12), 233(12), 236(12), 237(12), 243(12)

622

AUTHOR INDEX

Tsiapalis, C. M., 16 Van den Berg, A., 398, 399(448, 4491, Ts'o, P. 0.P., 52,299,449,450,452,456, 400(447-449, 450). 403(472), 404,

545 Tsong, T. Y., 388 Tsou, K. C., 542 Tsugita, A., 222,227(16), 525 Tsuji, I., 427 Tsukada, K., 423 Tsuruo, T., 407,408(490) Tulinsky, A., 437 Tumerman, L., 538 Tushinski, R., 227 Tutas, D., 255 Tutas, P. J., 51 Tysper, A,, 147 Uchida, H., 64 Uchida, T., 435,436(2),437,440(2), 441(2),

442(2), 444(2), 445(2), 447, 448(2), 449(2), 450(2), 458(2), 464(65), 465(2), 466(2, 127),467(127), 468(127), 501 Ueda, K., 156,158, 161,162(30) Uemura, H.,34, 38(23), 42, 43(44), 44, 54(47), 5347). 56 Uenishi, N., 447 Ueno, K., 137 Uhlenbeck, O., 20 Uhlenbeck, 0. G.,32,33, 34(14),37,38(32), 40(34,36).41(40), 42(3,7,32,34),43(7), 51, 52, 53(34, 36), 55(32, 48,68). 56, 57(40,42), 58(42), 276,544 Uhlmann, A,, 86,168 Ukita, T., 407,408(490), 450,464 Ulbricht, T. L., 426 Ulpino, L., 131, 132(87) Urneda, T., 423 Umeyama, H., 432 Urata, Y.,323 Uriel, J., 524 Usher, D. A., 430,431 Valentine, R. C., 522,531(60) Valenzuela, P., 11.9,126(51), 128, 131, 132(91), 133, 134(99), 135, 136(112),

192,214 Valiukaite, R. V., 465 Vallee, B. L., 131, 132(87), 340,341(146),

342(146),384(148) Van Batenburg, 0. D., 347, 348(211), 349(206),350(213), 352,353

405(448),407(450),409(449,450) Van den Berg, M., 57 Van den Broek, H., 146 Van den Broek, J. W. J., lIl(20). 112,

113(20), I16(20), 117(20) Van den Broek, W., 418,419 Vandenbussche, P., 286,311(29), 312 Van den Hende-Tirnmer, L., 398,399(449),

400(449,450),407(450), 409(449,450) Van den Rijn, I., 520,539(26) Von der Helm, K., 84 VanderLaan, K.,107 VanDerLijn, P., 540 Van der Meide, P. H., 271,273(29) Vanderslice, R. W., 48 Van der Zee, R., 341,342.372 Van de Sande, J. H., 8, 9(34), 14, 16, 19,

20(55), 21(55), 28 Van Dijk, B., 403(472), 404,405(447) Van Dijk, H., 398,400(447) Vang, N. H., 531,546(117) Van Keulen, H., 119,138(52), 138,144 Van Kraaikarnp, M.,417 Van Rapenbusch, R., 521, 522(41), 526(41) Van Schagen, C. G., 366,367(301) Van Vliet, D.L., 332 Van Wart, H.E., 344,345(184),388(184) Vaquero, C. M., 286,295,304(57) Vasilenko, S. K., 34,35(17) Vassart, G., 423 Vasseur. M., 511 Vaughan, M., 227 Vecchini, P., 412 Vekstein, R., 52 Venegas, A., 133, 134(99),214 Venetianer, P., 397 Veniyaminova, A. G., 34,3317) Venkatesan, S., 248,249(20), 251(20),256, 26I Venkov, P., 502,507,511(12) Venkstern, T.V., 465,559 Vennstrom, B., 547 Verhaegen, M., 286,311(29) Verhaegen, N.,312 Vescia, S., 433 Vicuna, R., 156,157(l I ) , l58(l I ) , lS9(I I ) ,

160(11), 166(11),168,169(45,46) Vidali, G., 350

623

AUTHOR INDEX Vince, R., 540, 541(191) Vincent, J. P., 341 Vinograd, J., 139, 142 Visser, A. J . W. G., 366, 367(301) Visser, J. P., 335, 347(105). 350( 105) Vithayathil, P. J., 335, 336(109), 337(l lo), 347, 356(118) Voelker, R. A , , 135 Vogeli, G., 582 Vogt, V. M., 140 Vokert, W. A., 344 Volkin, E., 412 Vollenweider. H. J. 275 von Borstel, R. C., 26 von der Haar, F., 185, 186(34), 187(34), 189(34), 190(34), 191(34), 196, 198, 213 von Hippel, P. H., 415, 428 von Minden, D. I.., 447 Vorbruggen, H., 542 Vosberg, H.-P., 28 Voskuyl-Holtkamp, I., 345, 348(213), 349, 350(213) Vournakis, J. N.. 57, 275, 548 Vretsky, S. C., 497 ~

Wachsrnan, J., 261 Wachsman, J . T., 17, 18(75), 32 Wacker, W. E. C.. 341 Wade, H. E., 502, 505 Waechter, C. J., 311 Wagner, G., 386 Wahba, A. J., 268, 270(12). 276 Waittiaux-De Coninck, S., 150, 151(21I ) Wakabayashi, T., 55, 56(79) Waku. K., 440 Waki, M., 436, 443 Walerych, W., 184 Walker, D. E., 331 Walker, E. J., 428 Walker, G. C., 32, 42(3), 544 Walker, R. T., 540 Wall, R., 227, 231 Wallach, D . , 284, 285(21) Wallace, S.. 168. 169(45) Walsh, K. E., 123 Walter, B., 325, 326 Walters, B. L., 341 Walters, D. E., 330, 335(75), 374, 375(75) Waltle, G., 182 Walz, F. G., Jr., 429, 436.437,444,447(49),

450, 452(6), 454, 455, 459, 460, 461(53), 463(53, 98) Warnpler, J., 384 Wandzilak. T. M., 131 Wang, A., 34 Wang, D., 319, 327, 328, 414(55), 415, 425(55), 427(59), 433(47) Wang, F.-F. C., 406, 407, 411 Wang, J. C., 14, 28, 142 Wang, J. H., 368, 369(307), 376(307), 377(307), 379(307), 386(307) Wang, T. P.. 56 Waqar. M., 179 Ward, S., 48 Warme, P. K., 385 Warner, H. R., 22 Warner, V., 48 Warren, B.. 327 Warwick, P. E., 542 Waskell, L., 88, 89(8), 91(8), 92(8), 94(8), 98(8), 9918) Waslyk, B., 148, 150 Wasylishen, R. E., 375 Washington, M. E., 427 Watanabe, K., 333 Waters, J. A., 541, 545(200) Watkins, J. B., 338 Watson, N., 489 Watson, R., 142 Watt, C. D., 346, 352(202), 355(202) Wattiaux, R., 150, 151(211) Weatherall, I.. 380, 381(329) Weaver. D. L., 385, 423 Weaver, R. F., 119. 120(50) Weber, H., 15, 16(62), 28(62), 275, 279 Weber, K., 67, 180, 268, 270(12, 13). 271, 272(27), 276, 277(13, 27) Weber. L., 259 Webster, D. A., 425 Webster, R. E., 485. 486(4), 487(4) Weeks, J. R., 135 Wegnez, M., 149 Wehrli. W., 538, 539(169) Wei, C., 246, 261(10), 262(10) Weickrnann, J. L., 319. 394(11), 406 Weigle, J. J., 4 WeiL P. A,, 115, 144, 147(38, 166), 149,262, 263(65) Weil, P. A., 115, 144, 147(38, 166), 149 Weill, D., 62

624 Weinberg, E., 243 Weinberg, F., 119, 126(51), 128, 214 Weinberg, R. A., 498 Weiner, A., 156, 168(5) Weiner, J. H., 158, 161 Weingartner, B., 148, 181 Weinstein, I. B., 578 Weinstein, L. I., 344, 345(184), 387, 388(184), 396(381) Weiss, B., 4, 6(10), 7, 8(29), 9(29), 12, 13(47, 48). 15(10, 23), 16(29), l7(10, 29, 48), 18(29, 481, 19(29, 48), 22, 27(29, 48) Weiss, S., 61, 62 Weiss, S . B., 115, 145, 572,573(22), 574(22) Weissbach, A., 154 Weissmann, C., 222, 2 4 3 l8), 268, 269, 270(10, 20), 272(10, 20), 273(20), 275, 276(20), 278(22), 279 Welling, G. W., 319,341,342(155), 345,363, 372(155), 398(462, 469, 399(9, 454, 4551, 400(7, 9, 45 I , 454, 455, 459, 460, 462, 4651, 403, 404(9, 190, 454, 455, 469, 4039, 4 6 3 , 407, 408(462), 409(451, 486), 410(486), 414, 421(454) Wells, J. R., 244 Wells, R., 100 Wennekes, L. M. J., 111(20), 112, 113(20), 116(20), I17(20), 146 Wensley, C. G., 134 Werenne, J.. 31 I Werner, R. P., 320 Wertheimer, A., 263 Wernitz, M., 437 Wertz, G. W., 496 Weser, U.,130, 131(85) West, D. K,, 284, 285(22), 287(22), 299, 310(22) West, R. W., Jr., 9, 20(40) Westergaard, O., 17 Westmoreland, D. G., 333,369,389(96,311) Westphal, H., 496 Wetlaufer, D. B., 391(406), 392, 413(405) Whichard, L. P., 427 White, C., 284, 285(20), 286(20), 296(20) White, C. N., 284, 285(24), 287, 291(24), 292(24), 295, 296(24), 297(24), 302(40, 581, 303(40), 304(40), 306(58), 307(58, 73), 310(58, 73) White, F. H., Jr., 338, 391 White, M. D., 323, 364, 426, 458

AUTHOR INDEX Whiteley, H., 76, 77, 78(122), 79, 80(122, 131, 132) Whitfield, P. R., 458, 459 Wiberg, J. S., 48 Wickens, M. P., 148 Wickner, S., 69, 141, 156, 157(8), 158(8), 159(8), 160(2, 8), 161(8) Wiebers, J. L., 533, 544(128) Wiegand, R. C., 284, 285(18), 304(18) Wieland, O., 146 Wieland, T., 133. 146 Wiener, J., 107 Wierbicka, H., 560 Wierenga, R. K., 319, 400(7), 406(7) Wiggs, J. L., 62, 65(11), 66(11, 12). 67(1 I), 70(12, 66), 71(66), 72(12), 73(11, 12, 65, 661, 75(66), 77(12), 78(11), 80(11), 82(66), 93, 105 Wilchek, M., 319, 320(5), 342, 384 Wilcox, G., 80 Wilczek, J., 426 Wilkinson, D. S., 497 Wille, H., 522, 526(57), 527(57), 530(81) Williams, B. R. G., 283. 292, 293(52), 296(52), 299, 301(64), 302(58), 304(14, 82),305(52,54), 306(52,58), 307(52,58), 308(79, 82), 309(82), 310(58), 312(52) Williams, D., 84 Williams, G. H., 423 Williams, J. G . , 498 Williams, K. R., 185(39), 186, 187(39), 188(39), 189(39), 190(39), 191(39), 192(39), l94(71), 196, 198(71), 201(71) Williams, R. C., 75 Williams, R. J. P.. 372, 373(316a) Williamson, V., 139, 143(149), 144(149), 1 4 s149) Williams, F. R., 521, 524(38), 525(38), 529(38) Wilson, D. P., 545 Wilson, G., 327, 328,414(55), 418,419(574), 420(574), 425(55), 433(57) Wilson, G. G., 9, 20(39) Wilson, J. H., 48 Wilt, E. M., 13, 14(54), 15, 16(54,63), 17(54, 63). 19(63), 25(63), 28(54) Wilt, F., 237, 238, 241(66, 68) Wimmer, E., 36, 496 Winnacker, E. L., 178, 181 Winsor, B., 127, 135(77)

625

AUTHOR INDEX Winters, M. A., 218, 219(2), 221, 222(11), 223(2, I ] ) , 224(11), 225(2, 11). 226(11), 228(2), 229111). 230(11), 231(11), 233(11), 234(11), 236(11), 237(11), 238(1 I), 243(1I ) Wintersberger, E., 150, 151(214, 215) Witkop, B., 541, 542, 545(200, 217) Witney, F. R., 111(16), 112, 113(16), 116(16), 139, 143(146, 147) Wittschieber, E., 360 Witzel. H., 330, 33377). 346(77), 366, 369(77), 372(77), 373(77), 376, 379(77), 428, 431(631), 432(631), 457, 458, 459 Wlodawer, A., 365, 366, 390(292) Wodak, S. Y., 320, 323(21), 330(21), 335(21), 364(21), 377(21), 406(21), 415(21) Woese, C. R . , 506 Wold, F., 320, 325, 326, 327, 328, 352, 354, 355 Wolfson, J., 174 Wong, T., 572. 573(22, 23). 574(22, 23) Wood, D. L., 31 1 Wood, J. N., 284, 285(19), 304(80, 81), 305, 308(78, 80, 81). 311(19), 519, 521(17), 522(17), 523(17), 524(17), 52317). 539( 17) Wood, W. B., 32, 34(8), 48(8), 49(8, 56, 58) Woodfin, B. M., 346, 352(198) Woods, P. S., 111(29), 112, 113(29) Woody, R. W., 339, 383 Woodward, C. K., 349, 367, 368(214), 370(2l 4). 374(214) Worst, R., 57 Wrathall, D. P., 388 Wreschner. D. H., 292, 293(53), 29603). 297(53), 298(53), 305(53), 306(53), 307(53), 312(53) Wright, R . , 16, 28(68) Wriston, J . C., 410 Wu, A. M., 184 Wu, C. W., 67, 75, 76(104), 132 Wu, G. J., 147, 150, 151(218) Wu, H., 99 Wu, J., 302, 303(69) Wu, R., 20, 27, 545 Wunderli, W., 538, 539(169) Wyckoff, H . W., 318. 320, 322, 323(1, 21), 325, 326(1), 327(1), 329, 330(21), 335(1, 21), 336(1), 337(1), 339(1), 340(1),

342(1), 343(1), 344(1), 346(1), 347(l). 349(l, 16), 351(1), 356(1), 358(1), 359(1), 360(1),!364(1, 16,21,40), 365(1), 36"). 372, 373(1), 377(21), 382(1), 384(1), 390(1), 404(1, 16), 406(21), 415(1, 16, 21). 420(1, 16), 424(1), 426(1), 429(1), 431, 432(1) Wyers, F., 117, 120(47), 123(47), 127(47). 137 Wyneken, U., 134 Wynvicz, A. M., 379, 380 Xue, C., 34 Yajima, Y., 359, 360(265) Yakobson. E., 282, 308 Yakovlev, G. I., 329, 335(71, 72), 369(71), 376(72), 377(72), 379(71, 72), 381(72), 382(72), 431 Yamada, Y., 502, 511(16) Yarnada, T., 28 Yamagata, S., 462 Yarnashita, J., 519, 521(19), 523(19), 52319) Yamashita, K., 410 Yamashita, S., 407, 408(490) Yamazaki, H., 548 Yamkovoy, V. I., 34, 35(17) Yamuguchi, N., 180 Yanaihara, C., 443 Yanaihara, N., 443 Yang, H. H., 519 Yang, K.-Y., 546 Yang, S . K., 571 Yang, S. P., 519 Yaniv, M., 139, 143(142, 144, 145) Yankeelov, J . A., Jr., 333 Yanofsky, C., 71, 73(83), 86 Yarbrough, L. R., 69, 141, 144, 145(167) Yarranton, G., 170 Yarus, M., 582 Yasuda, S., 161, 162(36) Yasuda, T., 579 Ybarra, D. M., 324 Yeates, D. G. R., 337, 365(119), 373(119) Yegian, C. D., 48 Yehle, C. D., 475 Yehle, C. O., 499 Yip, K. F., 542 Yon, J., 525 Yoneda, M., 388

626 Yoshida, H., 436, 449, 452, 464, 465, 466( 135) Yoshida, K., 465 Yoshida, M., 464 Yoshida, N., 447,459,466(143), 467,468(62) Yoshikawa, M., 504 Yoshino, H., 519, 525(20) Yot, P., 536 Young, D. A., 425, 427 Young, D. M., 358, 391 Young, E. T., 4 Young, R. A., 271, 272(31),486,491(10, I I ) , 493( I I ) , 494(1 I), 4 9 3 I I ) Young, H . , 18 Young, R., 72 Yudelevich, A., 104, 170 Yuki, A., 502, 511113, 14) Yuki, R.. 449, 452 Yura, J., 68 Yura, T., 63, 64,74(18), 76(18) Zabel, P., 551 Zabinski, M., 459, 461, 463(98) Zaborsky, 0. R., 395, 396(426) Zachau, H. G., 202, 526, 530(83), 535(83), 536(83) Zagari, A., 413 Zagarska, L., 551 Zahn, R., 227, 232(31), 237(30), 238(30), 240(30) Zaldivar, J., 133, 134(99)

AUTHOR INDEX

Zamecnik, P. C., 184, 185(7) Zarbl, H., 264 Zaron, de Behrens. G., 521, 522(45), 526(45), 527(45), 547(225) Zechel, K., 64, 156 Zehring, W., 88, 107, 227 Zelipka, J., 465 ZelinkovB, E., 449, 452(70), 465, 467(70) Zendzian, E. N., 398 Zenebergh, A,, 328, 43308) Zhenodarova, S . M., 465, 467(141) Ziff, E., 464 Zilberstein, A., 282, 290, 301(45), 302(65), 304(65), 305(65, 7 3 , 306(65), 307(65), 3 lO(45) Zilinskiene, V. J., 36, 37(25) Zillig, W., 63, 64, 65, 67(47), 74(22), 75(22), 76(22, 46), 77(47, 115, 116), 82(47), 86(22) Zimmerman, S. B., 5, 7, 12, 13(46), 15(20), 17(20), 18(20, 46), 19(20),27(20) Zinder, N. D., 169,268,270(16), 485,486(4), 487(4) Zinn, K., 94, 95(38) Zipursky, S. L., 156, 173, 174(72) Zivin, R., 107 Zmudzka, B., 541, 545(198, 199) Zucchelli, G. C., 447 Zutra, A., 533, 543, 544(127, 228), 546(228) Zweers, A,, 417, 418 Zwiers, H., 398(464), 399, 400(464)

Subject Index

A A . castellanii RNA polymerases, subunit structure, 128, 129 A. vinelandii PNPase, 529, 530 Abortive initiation reaction, eukaryotic RNA polymerase, 144-145 Acylation, pancreatic RNase, 325 Acetylation, tyrosine residues, 340 Adenosine derivatives N6-isopentenyladenosine,575-577 2-methylthio-N6-isopentyladenosine, 577 modifying enzymes, 575-578

N-[(9-p-~-ribofuranosylpurin-6-y~)carbamoyl]threonine, 577, 578 structure, 576 Adenosine diphosphate P-substituted, 54 derivatives, 42-44 adenosine triphosphate replacement, 159, 160 Adenosine monophosphate acceptance, transfer RNA, 202 anomalous incorporation, 210, 21 1 Cpc, acceptor, 205, 207 residue, T4 RNA ligase, 36, 37 Adenosine triphosphate analogs, poly(A) polymerase, 227 inhibitor. 23 1 627

binding subsites, 207-209 cofactor, phosphodiester formation, 17, 18 derivatives, 303 requirements, 2‘,5’-oligoadenylate synthetase, 300 transfer RNA nucleotidyltransferase, 196, I97 ATP-independent reaction, T4 RNA ligase, 42-44 S-Adenosylmethionine, transcription stimulation, 263 Adenovirus DNA replication, 182 specific binding, 143 synthesis priming, 180-182 transcription, 147, 148 Aden yl ylation donor 3‘-phosphate, RNA ligase, 45 stereoisomer, 39 substrate specificity, 38 T4 RNA ligase, 37-39 2’,5’-oligoadenylate synthetase, 303, 304 ADP-Pi exchange reaction, activators, 529-530 AF/103, poly(A) polymerase, inhibitor, 232 Affinity chromatography pancreatic RNase, 318, 319

SUBJECT INDEX poly(A) polymerases, 220, 221 RNase T I , 436 transfer RNA nucleotidyltransferase, 188 Affinity labeling, eukaryotic RNA polymerase, 132-134 Aggregation, S-protein, 354, 355 Alcohols, polyhydric, 114 Alkylation pancreatic RNase bromoacetate, 331 carboxymethyl derivatives, 331 kinetic constant, rate, 331 iodoacetamide, 330 methionine residues, 336, 337 reductive, 323, 324 RNase A amino groups, 323, 324 histidine residues, 329-331 RNase T I , histidine residues, 441 Amber mutation E coli CTrSx, 50 RNA ligase, 48, 49 Amanitin, RNA polymerase IIinhibitor, 133 Amanitin, resistant RNA polymerase, 135 Amanitin, concentration, 146 Amanullin, eukaryotic RNA polymerase, inhibitors, 145, 146 Amatoxins, eukaryotic RNA polymerase inhibitors, 145, 146 Amidination lysine residues, cytoplasmic RNase inhibitor, 422 pancreatic RNase, 326, 327 Amino acid composition cytoplasmic RNase, 419, 420 DNA ligase, 10 guanyloribonucleases, 466, 467 PNPase, 523, 524 T7 RNA polymerase, 94, 95 sequence allelic polymorphism, 399 amino acid substitutions, 404, 405 evolutionary history, 403, 404 half-cystine residues, 406 heterogeneity, 399 histidine residues, p K,, 406, 407 pancreatic RNase A, 321 proline substitutions, 404 residues, role, 405 secondary structure, 406

species variation, 397-41 I termination codon, mutations, 399, 403 3-(3-Amino-3-carboxypropyl)uridine, 569, 513, 574 Amino groups pancreatic RNase, 320-329 acylation, 325 alkylation, reductive, 323, 324 amidination, 328, 329 amino acid residue sequence, 321 CDNCB, 324, 325 cross-linked dimers, 327, 328 glycosamination, 328, 329 inactivation, 326 +NH2 groups, 325 poIy(A), 326 pyridoxal phosphate, 320, 323 Schiff bases, 320, 323 structure, 322 RNase T ,, 439, 440 Antibiotics, poly(A) polymerase, inhibitor, 232 ApC, CMP acceptor, 205 Aqueous fractionation methods, poly(A) polymerases, 237 Arabinose, RNase inhibition, 426 Archybacteria, RNase polymerase, 77 Arginine residues pancreatic RNase, 333-335 RNase TI, 441 Arsenolysis, PNPase, 536 Asparaginyl residues, S-peptide-S-protein interaction, 346 Aspartate, His-I 19-Asp- 12I interaction, 374, 37s Aspartate residues, pancreatic RNase, 335, 336 Aspergillus nidulans, amanitin-resistant RNA polymerase 11, 146 Assay activity, 112-1 15 DEAE filter, 114 DNA ligase, 5-7 dnaG gene product, 156, 157 methyltransferase enzymes, 558, 559 novel specificity, RNA polymerase, 7173 poly(A) polymerase, 226 pseudouridylate synthase, 571 Qf3 replicase, 269, 270 qualitative, RNA polymerase, 72

SUBJECT INDEX

629

DNA-dependent, 70-72 quantitative DNA-dependent RNA polymerase, 69, 70 eukaryotic RNA polymerase, I15 T7-like bacteriophage DNA-dependent RNA polymerase, 93 TCA precipitation, 114, 115 RNA, 114 RNase, 424-426 RNase I, 504 RNase 11, 506, 507 RNase D, 481 sulfurtransferase, 572, 573 T4 DNA ligase, 8. 9 T4 RNA ligase, 33, 34 T7-like bacteriophage DNA-dependent RNA polymerase, 91-94 transfer RNA : guanine transglycosylases, 580 transfer RNA nucleotidyltransferase, 186 ATP(CTP)-transfer RNA nucleodyltransferase, see Transfer RNA nucleodyltransferase ATP-PP, exchange reaction, T4 RNA ligase, 36, 37

B B . subfilis PBS2 RNA polymerase, 105, 106 B . subtilis RNA polymerase, 78, 79 B. subrilis transfer RNAm”’, 7-methylguanosine, 563 Base Q,579-582 structure, 579 Base Y,578, 579 Binding constants poly(A) polymerase, 233 RNase T I , 453 S-peptide-S-protein interaction, 352, 353, 355 transfer RNA nucleotidyltransferase, 28, 194, 196, 197 Biosynthesis, RNase, 361, 362 Borohydride, reduction alkylation, RNase A, 324 C

C5 protein, RNase P, reconstitution, 472, 473

Calf thymus DNA, 114 template activity, 139, 140 Calf thymus RNA polymerase I, 126, 127 Calorimetry, S-peptide-S-protein interaction, 352, 353 Cap binding proteins, 259 Capping enzyme, see RNA Guanylyltransferase Capping reaction, characteristics. 258 CarMV RNA, 3’ sequence, 552 Carbohydrate moities glycosidated RNase, 408, 409 glycosylation sites, 410 side chains, effects, 41 1 variations, 407-41 1 2-Carboxy-4,6-dinitrochlorobenzene, pancreatic RNase, 324, 325 Carboxyl groups, modification, RNase T I , 437-439 Carboxy methylation cytoplasmic RNase inhibitor, 421 RNase T , , 437 COOH termini, pancreatic RNase, 345-359 see also S-Peptide-S-protein interaction Cell cycle, poly(A) polymerase, 240 differentiation, poly(A) polymerase, 241, 242 whole, 2’,5’-oligoadenylates, 307, 308 cores, 309 Chloroplast RNA polymerase, 151, 152 Chick embryo cells, interferon treated, purification, 288, 289 Chromatography, see specific types Circular dichroism RNase A, 382-384 RNase T I , 456, 457 Circularization reaction, T4 RNA ligase, 41, 42 CMP anomalous incorporation, 210 ApC, acceptor, 205 transfer RNA-N, 209 tfansfer RNA nucleotidyltransferase, 204, 205 Column chromatography, eukaryotic RNA polymerase, 116-1 17 COOH terminus, modification near, 356359 CpC, AMP acceptor, 205, 207 Crystallization, RNase, 364, 365

630

SUBJECT INDEX

CTP binding subsites, 207-210 transfer RNA nucleotidyltransferase, 196, I97 Cyanoborohydride, reductive alkylation. RNase A, 324 Cysteine residues PNPase, 523, 524 RNase refolding, 392 Cytidine, 574 CMP incorporation, 204 structures, 575 Cytoplasm, polyadenylation, 235-237 Cytoplasmic RNase, inhibitor amidination, 422 amino acid compositions, 419, 420 carboxymethylation, 421 chemical properties, 416-423 contact points, 422 in vitro protein synthesis, 423, 424 lysine residues, 421 placental inhibitor, 418-420 purification, 417

D 3'-dATP, poly(A) polymerase, inhibitor, 231, 232 DEAE filter assay, RNA, 114 Deamidation, RNase A, 336 Denaturation, eukaryotic RNA polymerase, I30 Deoxynucleoside triphosphates, primer formation, 158 Deoxyribonucleotides, polymerization, PNPase catalyzed, 545 Depolymerization, RNase, 429, 430 Dinucleotide phosphates, analogs, RNase, 380-382 Diribronucleoside monophosphates, adenylyl transfer reactions, 303 Dissociation constants RNase T I fluorescence, 456 gel filtration, 450, 451 UV-difference spectroscopy, 453-455 transfer RNA nucleotidyltransferase, 199 Disulfide bonds

pancreatic RNase, 337-339 RNase T,, 442 Disulfides, RNase refolding intact, 390 reduced, 391, 392 Divalent cation, see also MgZ+;Mn2' eukaryotic RNase, requirements, 498 RNase 11, requirements, 507 RNase 111, requirements, 487 DNA acceptor, phosphodiester bond forming, 40 bacteriophage dnaG-dependent priming, 162- I63 binding protein, 160, 161 a3, 162, 163 4x174, 162, 163, 165 pathways, 161, 162 calf thymus, 114 double-stranded priming, 169-174 fragments, joining, 15, 16 leading strand synthesis, 170-172 mechanism, 173, 174 minus strand synthesis, 171-173 recombination, 4 relaxed, unnicked, 142 restricted, 139 single-stranded, replication, 160- 165 protein requirements, 164 4x174 DNA, replication, 169 +XI74 R F DNA replication, 171-173 pathways, 172 4x174 RFI DNA, replication, in vifro, 170-172 RNA polymerase, 166-169 template activity, 140 catalytic reactions, 83 dnaG-dependent pathways, 165-167 eukaryotic RNA polymerase, 138-140 unnicked, 141, 142 DNA-adenylate intermediate, 13, 14 DNA-AMP intermediate, 13, 14 DNA-directed reaction, RNA polymerase, 83-86 DNA ligases, 1-29; see also E. coli DNA ligase; T4 DNA ligase amino acid composition, 10 assays, 5-7 biological activity, detection, 6

SUBJECT INDEX choice of, 7 partial reaction, 6, 7 phosphodiester bond formation, 6 bacterial, physiological requirements, 25,26 bacteriophage, physiological requirements, 25, 26 bacteriophage-induced, 21-23 catalytic properties, 10-21 blunt-end joining, 19-21 phosphodiester formation, see Phosphodiester formation reactions, reversal, 14-15 E. cofi-induced, 4, 5 in vivo, role, 21-26 isolation, 5-10 joining reaction cofactor requirement, 17, 18 divalent cation, 18, 19 pH optimum, 18 ligase-adenylate intermediate, 12, 13 molecules recombinant, preparation, 28, 29 structure, study, 28 mutants, 26 physical properties, 5- 10 amino acid composition, 10 molecular weight, 9, 10 purification, 8, 9 research applications, 26-29 TCinduced, 4, 5 DNA-negative mutants, 34 dnaC protein, 156-160 ADP, ATP replacement, 159, 160 assay, 156-157 catalytic properties, 157-160 de novo synthesis, 157, 158 deoxynucleoside triphosphates, addition, 158 molecular weight, 157 primer formation, 159 priming pathways, 160-165 a3, 162, 163 6x174, 162, 163. 165 purification, 157 stoichiometry, 159 template interactions, 165- I67 Donor blocking group, oligonucleotide synthesis, 55 Duplex structure, RNA ligase reaction, inhibitor, 40, 41

63 I E

E. coli cr, 67

dnaG protein, see dnaC protein infection, T4, 49-50 RNA ligase source, 34 RNases, see specific RNase transfer RNA nucleotidyltransferase, biological role, 213, 214 transfer RNA processing enzymes, see specific RNases E. cofi bacteriophage N4 RNA polymerase, 106- I08 E. coli BN, 482 E. coli CTrSx, T4 infection, 49, 50 E. coli DNA ligase, 23-25, see also DNA ligase joining activity, destruction, 13 joining reaction, pH optimum, 18 mutations, 23, 24 physiological requirement, 25 purification, 8 reversal reaction, 14, 15 T4 gene 30 mutants, 23 E. coli N464, RNase 11, 51 I E. coli RNA polymerase cr, 67 core, 66, 67 holoenzyme, 66 extinction coefficients, 81, 82 preparation, 64, 65 renaturation, 130 subunit structure, 74-76 E. gracilis RNA polymerase I1 inhibition, 132 EF-Tu .Ts, QP replicase, 272,273, 277, 278 Ehrlich ascites tumor cells ATP requirements, 300-301 2’,5’-oligoadenylate synthetase, purification, 288 dsRNA concentration, 299 specific activity, 301 Elongation PNPase, 532 QP replicase, 278-279 EMC virus, infection, interferon and, 307308, 311 Endometrial cancer cells, human, interferon treatment, 311

632 Endoribonuclease, see also Specific RNases activation, 2’3’-oligoadenylates, 305-307 substrate specificity, 306, 307 Enzyme-AMP formation, T4 RNA ligase, 36-37 Esterification, aspartate residues, 335, 336 Estrogen, poly(A) polymerase, activity and, 240, 241 3-Ethoxy-2-ketobutanol, arginine residues, 334 Eukaryotic DNA replication, priming, 178182 Eukaryotic messenger RNA cap, structure, 246, 247 Eukaryotic RNA polymerase, 109-153 catalytic properties DNA template, 138-140 nicked, initiation, 140, 141 sequence-specific initiation, 143, 144 unnicked, initiation, 141, 142 DNA binding, 138-145 nicks, 140, 141 sequence-specific, 143, 144 template, 138-140 unnicked, 141, 142 inhibition amanin, 133 amatoxins, 145, 146 1,lO-phenenthroline, 131 pyridoxal 5’-phosphate, 133, 134 rifomycin derivatives, 146, 147 mutant, 134, 135 priming, 141 purification, 1 11-1 17 activity assays, 112-115 column chromatography, 116, 117 nucleic acid removal, 116 polyhydric alcohols, role, 114 procedures, 115-1 17 purity, 117 solubilization, 115, 116 sources, I 1 1-1 I3 yields, 111-113 stimulatory factors, 137 subunit functions, 128-137 active site, 132-134 affinity labeling, 132-134 mutant polymerases, 134, 135 phosphorylation, modification, 135, 136 purified, 128

SUBJECT INDEX reconstitution studies, 129, 130 renaturation studies, 129, 130 zinc, role, 130-132 subunit structure, 117-128 active enzyme, 126, 127 determination, 120-122 largest peptide, 123-126 polymerase I, 127, 128 polymerase 11, 123-128 polymerase 111, 127, 128 polypeptide composition, 118, 119 quantitation, 122 SDS-polyacrylamide gel electrophoresis, 120-122 size, 119 transcription extract systems discovery, 147, 148 fractionation, 148, 149 in vivo, mimicing, 149, 150 Eukaryotic RNA polymerases I1 largest polypeptide, 123- I26 stoichiometrics, 131 Eukaryotic RNase, RNA processing, 497499 Eukaryotic viral RNA, secondary site cleavage, 496 Eukaryotic virus, RNA cap synthesis, 246248 Exogenous DNA, transcription extracts, 147 Exonucleases, see also specific RNases activity, measurement, 27 Extinction coefficients, RNA polymerase holoenzyme, 81, 82

F fd DNA, replication, RNA polymerase, 169 Fluorescence dissociation constants, RNase T I , 456 tyrosine residues, 384, 385 Folding pathway, RNase, 385-397 antigenic sites, 394 C-2 proton resonances, 373, 374, 387 conformation native stability, 386-388 protein, 389 transition, 373-374 cysteine residues, 392 disulfides, 390-392

SUBJECT INDEX

633

equilibrium studies, 386-389 histidine residues, 373, 374, 387 immunochemical approach, 389, 393-397 in vivo, 397 isomerization, 390, 391 kinetic studies, 390-397 proline isomerization, 390, 391 properties, unfolded, 388, 389 refolding, 390-392 S-carboxymethylation, 392, 393 S-peptide, 391 S-protein, 388, 394 sulthydryl intermediates, 392, 393 terminal segments, 388 unfolding stages, 387, 388 thermal, 395, 396 Fractionation bacterial RNA polymerase, 65 chromatin, 238, 239 nucleoplasmic, 238, 239 transcription extracts, 148, 149 Fragment condensation, RNase T,,synthesis, 443 Friend cells cycle, 240 differentiation, 241 G

GCRFI DNA, template, 171 Gel filtration, RNase T I , 449-452 Gene 4 protein, lagging-strand DNA synthesis, 174 Guanine-7 methylation, 260, 261 Guanine-specific RNase, 465-468 amino acid composition, 466 Guanosine derivatives, modifying enzymes, 578-582 base Q, 579-582 base Y,578, 579 Glutamate residues pancreatic RNase, 335, 336 RNase T I , 437. 439 Glutamine residues, pancreatic RNase, 335, 336 Glycosamination, reductive, 328, 329 Glycosylation, pancreatic RNase, 408-410 GMP intermediate messenger RNA capping, 253, 254

transguanyl ylation, 26 I Goldberg-Hogness box, 150 GTP-PP, exchange, 252 Guanidination, lysine amino groups, 375 Guanine derivatives, binding, RNase T,, 450, 452, 453 Guanosine triphosphate, Qp replicase, initiation, 273-275 Guanyloribonucleases, amino acid composition, 466, 467 Guanylylation, guanine-7 methylation, relationship, 260, 261 Guanylyltransferase, see RNA guanyly Itransferase

H Heart, metabolic activation, poly(A) polymerase, 242 HeLa capping reaction, 256 HeLa cell ATP, requirements, 300 guanylyltransferase, 256, 257 pyrophosphate inhibition, 257 5-methylcytidine, 560 poly(A) polymerases, 238 Holoenzyme, preparation, 66 Histidine residues C-2 proton resonances, 376, 377, 387 NMR, 366, 367, 369 RNase T I , 444 conformational transition, 373, 374 deprotonation, 374, 375 His-1 19-ASP-121 interaction, 373, 374 hydrogen exchange rate, RNase T I , 445446 mononucleotide phosphates, effect, 376378 NMR, 366-372 C-2 proton resonances, 366, 367, 369 chemical shifts, 371 ionization, 368 spectra, 377, 378 pancreatic RNase, 329-333 bromoacetate, 331 carboxymethyl derivatives, 33 I iodoacetamide, 330 kinetic constants, 332 PKO changes, 377-379

634

SUBJECT INDEX

values, 406, 407, 444, 445 RNase T I , 440, 441, 444, 445 S-peptide+protein interaction, 347-349 titration curves, 373, 374, 444, 445 Tyr-25, 373, 374 Hormones, sex, poly(A) polymerase, 240, 24 1 Hydrogen exchange rate, histidine residues, 445, 446 Hydrogen ions, poly(A) polymerase, 230 Hydrolysis assay, RNase, 425, 426 RNase 11, 508 RNase D, 481 step, RNase catalysis, 430 transfer RNA, 211-213 I Immunochernistry, RNase, 362-364 folding, 389, 392-397 Immunological analysis, PNPase, 524, 525 Influenza virus, transcription, cap role, 259, 260 Inhibition eukaryotic RNApolymerase, 131, 133, 134 amanin, 133 amatoxins, 145, 146 1 ,lo-phenanthroline, 131 pyridoxal 5'-phosphate, 133, 134 rifomycin derivatives, 146, 147 HeLa cell capping enzyme, 257 2',5'-oligoadenylate synthetase, 302 phosphodiester formation, 19 PNPase, 528, 529 poly(A) polymerase, 231-233 QP replicase, 278 RNA ligase, 40, 41 RNA polymerase I, 133, 134 RNase, 416-420, 427 transfer RNA nucleotidyltransferase, 198 Initiation abortive, eukaryotic RNA polymerase, 144, 145 at nicks, 140, 141 unnicked, 141, 142 sequence-specific, 143, 144 Intercalating drugs, poly(A) polymerase, 233 Interferon

2',5'-oligoadenylate synthetase, induction, 286, 287 2'3'-oligoadenylate system, 309-3 12 treated whole cells, 307, 308 treated messenger RNA, 309, 3 10 Intermolecular forward reaction, T4 RNA ligase, 33, 36-41 Intermolecular reaction, circularization reaction, 41, 42 Iodination, RNase A, 339 lodoacetamide inactivation RNase A, 330 RNase T I , 440, 441 Ion exchange, poly(A) polymerases, 219, 220 Ion exchange chromatography, eukaryotic RNA polymerase, 120 Isoelectric point, PNPase, 523, 524 Isomerization, proline, RNase, 390, 391 N6-isopentenyladenosine, 575-577 structure, 576 K KiBabl mouse fibroblast, interferon treatment, 31 I Kethoxal, arginine residues, 334 L Lagging-strand synthesis, bacteriophage T4, 177 Lambda DNA, bacteriophage nicked, 141 unnicked, 142 LiCI, denatbration, eukaryotic RNA polymerase, 130 Ligase-adenylate intermediate, DNA ligase, 12-13 Ligase-AMP compounds, DNA ligase, 13 Ligase reaction, reversal, 14-15 Ligases, see s,pecific ligases Lowry protein method, RNA polymerase concentration, 82 Lymphocyte DNA, template transcription, 140 Lysine amino group, 375-376 cytoplasmic RNase inhibitor, 421 amidination, 422

SUBJECT INDEX

635 M

5-methylcytidine, 559-561 I-methylguanosine, 559, 561 MI RNA, RNase P, reconstitution, 472,473 7-methylguanosine, 559, 563 MDV-I , elongation, 279 N2-methylguanosine, 559, 562, 563 Messenger RNA ribothymidine, 559, 564, 565 breakdown, 2'.5'-oligoadenylate system, structures, 558, 559 309, 310 Methyltransferase reaction, characteristics, cap role, 258, 259 254-255 capping Mg2+ mechanism, 252, 254 DNA ligase, joining activity, 18, 19 multiple poly(A) polymerases, 223, 224 reaction sequence, 253 nuclear poly(A) polymerase, 237-239 degradation, non-interferon-treated, 3 12, 2',5'-oligoadenylate synthetase, 300-301 3 I3 PNPase requirements, 525 interferon treated, 309, 3 10 poly(A) polymerase, 229, 230 3'-noncoding sequence, 552 RNA synthesis, 112 poly(A) transfer RNA nucleotidyltransferase regulatory role, 550, 551 removal, 548, 549 activity, 193, 194 Missense mutants, RNA ligase, 48 poly(A) polymerase, 243 Mitochondria1 RNA polymerase, 150, 151 translation, 295, 296 Methionine residues Mitogen, poly(A) polymerase, 240 Mn2+ pancreatic RNase, 336, 337 S-peptide-S-protein interaction, 346-347 DNA ligase, joining activity, 18, 19 I-Methyladenosine, 561 multiple polymerases, 223, 224 structure, 559 PNPase requirements, 525 5-Methylaminomethyl-2-thiouridine, 565, poly(A) polymerase, 229, 230 566, 574 priming, inhibition, 141 structure, 569 QP replicase, template specificity, 274 Methylation, see olso Guanine-7 methylRNA synthesis, 112 ation specific binding studies, 143 capped RNA, 248 transfer RNA nucleotidyltransferase RNA, enzymes, see Methyltransferase activity, 195 enzymes Molecular weight 5-Methylcytidine, 559-561 DNA-dependent RNA polymerase, substructure, 559 units, 74, 77 I-Methylguanosine, 561, 562 dnaG protein, 157 structure, 559 DNA ligase, 9, 10 7-Methylguanosine, 563 poly(A) polymerase, 225 structure, 559 QP replicase, 271 N'methylguanosine, 562, 563 T7 RNA polymerase, 94 structure, 559 transfer RNA, 187, 190, 191 Methylhistidines, S-protein binding capacMononucleoside diphosphates, capping, ity, 348-350 25 I 2-Methylthio-Nb-isopentenyladenosine, Mononucleotides, RNase T I inhibition, 576, 577 458 Methyl transferase enzymes, 559-566 Monovalent cations assay, 558, 559 poly(A) polymerase, 230 I-methyladenosine, 559, 561 QP replicase, 274 5-methylaminomethyl-2-thiouridine, 565, RNase 11, 507-509 566, 569, 574 RNase 111, 487

636

SUBJECT INDEX

N

0

N4 RNA polymerase, bacteriophage, 106- 2',5'-oligoadenylates, cores, whole cells 108 effects, 309 NDP-Pi exchange reaction, 534-535 2',5'-oligoadenylate synthetase, 281-3 I3 Neuroblastoma cells, differentiation, 241, 2'-adenylylation, 303, 304 242 biological role, 304-313 NH2 terminus, pancreatic RNase, 345-359, 2',.5'-oligoadenylate cores, 309 see also S-peptide-S-protein interacinterferon action, 309-312 tion non-interferon-treated cells, 312, 3 13 NH4, DNA ligase. joining reaction, 19 nuclease activation, 305-307 Nitration, tyrosine residues, 340-342 occurrence, whole cells, 307, 308 Nuclear magnetic resonance diribonucleoside monophosphates, 303 RNase, 366-382 interferon treatment, 268-288 chemical shifts, 371 levels, 286, 287, 310 histidine residues, see Histidine resmolecular weight, 289 idues nucleotidyl donation, 303 ionization, 368 occurrence, 285, 286, 307, 308 proton resonance, 366, 367 polypeptide composition, 289 RNase A, titration, 346 properties, 289, 290 RNase T I , 457 purification, 287-289 Nuclease reactions catalyzed, 290-304 activation, 2',5'-oligoadenylates, 305analytical methods, 292 307 assay, 293-297 poly(A) polymerase contaminant, 222, ATP requirements, 300 223 inhibitors, 302 Nucleic acid kinetics, 301, 302 modification, RNA ligase, 57, 58 Mg2+, requirements, 300-301 removal, eukaryotic RNA polymerase, nuclease activation, 294-296 1 I6 oligoadenylate activity, 294-297 Nucleophilic displacement, phosphodiester oligoadenylate synthesis, 290-302 bond formation, 39 phosphatase-resistant core, 291 Nucleoside diphosphate product heterodispersity , 292, 293 functional regions, 543 product size, 301 modified, polymerization, 540, 541 product stability, 302 oligonucleotide synthesis, 543-544 product structure, 290-292 radiolabeling, 547, 548 protein synthesis, inhibition, 294-296 substrates, modification, 533, 534 prototype reaction, 290 Nucleoside triphosphates radiobinding assay, 297, 298 cleavage, 255 radiochemical assay, 293, 294 nucleotidyl donation, 303 radioimmune assay, 297, 298 transfer RNA, 196, 197 reaction mechanism, 301, 302 Nucleotides reaction requirements, 297-301 modifcation. 567-582 dsRNA, requirements, 297-299 adenosine, 575-578 Oligodeoxyribonucleotide, joining, RNA cytidine, 574, 575 ligase, 56 guanosine, 578-582 Oligonucleotides uridine, 568-574 circularization reaction, 41, 42 synthesis, radiolabeled, 547-548 fingerprinting, 547, 548

SUBJECT INDEX

637

PNPase, primers and inhibitors, 529 size. phosphodiester bond formation, 40

synthesis defined sequence, 543, 544 donor blocking group, 55 RNA ligase, 53-56 Oligoribonucleotide primer, 175 de n o w synthesis, 157, 158 Organelle-coded RNA polymerase, 150, I52

chloroplast, 151, 152 mitochondrial, 150, 151 Oxidation methionine residues, 337 periodate, RNA decapping, 265 Oxovanadium ion, RNase inhibition, 427

P Pancreatic RNase, 317-433, see also specific RNase bovine, 414 catalytic properties, 424-433 activators, 427 arabinose, inhibition, 426 assays, 424-426 depolymerization, 429, 430 hydrolysis, step, 430, 431 inhibitors, 426, 427 kinetics, 428-430 mechanism, 430-433 mechanistic models, 43 1-433 substrate conformation, 43 1 transphosphorylation, 430, 43 I chemical properties, 320-364 acetylation. 340 alkylation, methionine residues, 336, 337

amino k i d sequence, see Amino acids, sequence amino groups, see Amino groups, pancreatic RNase antigenicities, 363 arginine residues, 333-335 asparagine residues, 335, 336 aspartate residues, 335, 336 biosynthesis, 361, 362

carboxypeptidase Y, 358 chemical synthesis, 359-361 COOH terminus, modification, 356-359 deamidation, 336 disulfjde bonds, 337-339 elongation, disulfide bond, 339 equilibrium constants, 353, 354, 357 esterification, 335-336 functional group modification, 320-345 glutamate residues, 335, 336 glutamine residues, 335, 336 histidine residues, see Histidine residues hydrogen bond, 358, 359 immunochemistry, 362-364 iodination, 339 methionine residues, 336, 337 NH2 terminus, residue role, 345-359 nitration, 340-342 radicals, reactions with, 342-345 radiolysis, 342, 343 S-peptide-S-protein interaction, see Speptide-S-protein interaction tyrosine residues, see Tyrosine residues U V irradiation, 343-345 glycosidated, carbohydrate moieties, 408, 409

inhibitor activity, 418 physical properties, 364-397 circular dichroism, 382-384 conformational transition, 373, 374 dinucleotide substrate analogs, interactions, 380-382 fluorescence, 384-385 folding pathway, see Folding pathway histidine residues, see Histidine residues His-lI9-Asp-12I interaction, 374, 375 hydrogen bond, 373 lysine amino groups, 375, 376 mononucleotide phosphates, 376, 377 NMR, 366-382 optical properties, 382-385 phosphate group interactions, 380 pK,, phosphate group, 380 pK, changes, histidine, 377-379 refolding, see Folding pathway spin-lattice relaxation time, 380

SUBJECT INDEX substrate analogues, interactions, 376382 substrate resonances, 379 tyrosine residues, see Tyrosine residues unfolding, see Folding pathway U V absorption spectra, 382 X-ray diffraction, 364-366 preparation, 3 18-320 affinity chromatography, 318, 319 pup-Sepharose, 319 research applications, 433 species variation, 397-41 1 amino acid sequence, see Amino acids, sequence carbohydrate moieties, 407-41 1 PBS2 RNA polymerase, bacteriophage, 105, 106 Penodate oxidation, RNA decapping, 265 PH optimal DNA ligase, joining reaction, I8 titration curves, histidine residues, 373374 transfer RNA nucleotidyltransferase, 193-195 I$K intercistronic region, secondary structure, 167 1,lo-Phenanthroline, enzyme inhibition, 131 Phenylalanine residues, 357 Phosphate, inorganic, poly(A) polymerase, inhibition, 231 Phosphate group, RNase interaction, 380 Phosphodiester, formation ATP, cofactor, 17, 18 at blunt ends, 11, 19-21 DNA-adenylate intermediate, 13, 14 DNA ligase activity, 6 ligase-adenylate intermediate, 12, 13 mechanism, DNA ligase, 10-14 at nicks, 10-12, 15-19 activators, 19 cofactor requirement, 17, 18 divalent cation requirement, 18, 19 DNA substrates, 15, 16 inhibitors, 19 pH optima, 18 RNA-DNA hybrids, 16, 17

RNA substrates, 16, 17 sulfhydryl requirements, 19 nucleophilic displacement, 39 oligonucleotide size, 40 reaction kinetics, 14 T4 RNA ligase, 35-36, 39-41 Phosphorylation eukaryotic RNA polymerase, 135, 136 poly(A) polymerases, 242 RNA polymerase, modification, 134, 135 Phosphorol ysis PNPase catalyzed, 530, 535-537 poly(A), removal, 548-551 proteolyzed enzyme, 527 3’ sequences, 549 synchronous, 548, 549 Photoinactivation, RNase A, 343, 344 Photooxidation, RNase, T I , 440 PK, histidine residues Cm-RNase T I , 444,445 RNase T,,444, 445 Placenta, human, inhibitor in vitro translation, 423, 424 RNase, 418-420 Plasmid DNA, supercoiled, 139, 142 transcription, analysis, 144 PNPase, 5 17-553 insoluble, 546, 547 physiological functions, 537-539 properties, 5 19-530 activators, 528, 529 amino acid composition, 523, 524 immunological analysis, 524, 525 inhibitors, 528, 529 intracellular distribution, 519, 520 isoelectnc point, 523, 524 metal ion requirements, 525 molecular weight, 522, 523 occurrence, 519, 520 oligonucleotides, 529 polyamines, 529, 530 primers, 529 proteolysis, 526-528 stability, 525, 526 subunits, 522, 523 purification, 520-522 reactions catalyzed, 518, 519, 530-537 arsenolysis, 536 de novo synthesis, 530-532

SUBJECT INDEX deoxyribonucleotide polymerization, 545 elongation, 532 equilibrium, 533 NDP-Pi exchange reaction, 534, 535 phosphorolysis, 530, 535-537 polymeriza:ion, 531-534 substrates, modified, 533, 534 transnucleotidation, 537 research applications, 539-553 conjugation. insoluble matrix, 546, 547 deoxyribonucleotide polymerization, 545 labeling, PNPase-directed, 553 NDP functional regions, 543 NDP modifications, 540, 541, 543 3' nonencoding sequence, 552 3'-OH region, regulatory function, 550552 oligonucleotide fingerprinting, 547, 548 oligonucleotide synthesis, 543, 544 phosphorolysis, synchronous, 548, 549 poly(A) removal, 550. 551 polynucleotide synthesis, 539-543 radiolabeled nucleotides, synthesis, 547 ribosomal RNA, 551 3' termini, 551 transfer RNA, 551 viral RNA, 551, 552 Poliovirus RNA, second site cleavage, 496 PofyiAt adding enzyme, see Poly(A) polyrnerases diphosphate-terminated, HeLa capping enzyme, 257 effect, amidination, RNase A, 326 elongation reaction, 236-237 removal, 550-551 messenger RNA, 548-549 Polyadenylation, poly(A) polymerase, 228, 235, 236, 243, 244 Polyamines PNPase, 529, 530 poly(a) polymerase, inhibitor, 233 transfer RNA nucleotidyltransferase activity, 195 Polyanions, QP replicase, inhibition, 278 Poly(A) polymerases, 217-244 biological role, 235-239 cytoplasmic, 235-237

639 de novo synthesis, 236, 237 elongation reaction, 236, 237 nuclear, 236, 238, 239 polyadenylation, 235, 236 subcellular localization, 236-238 inhibitors, 23 1-233 antibiotics, 232 intercalating drugs, 233 product analog, 231, 232 substrate analog, 231, 232 multiple, 223-225 polyadenylation, 228, 235, 236, 243, 244 properties, 219-223, 225, 226 assay, 226 ion requirements, 229, 230 kinetics, 233-235 molecular weight, 225 primer requirements, 227-229 specific activity, 221, 222 purification, 219-223 affinity chromatography, 220, 221 contaminating enzymes, 222, 223 ion exchange, 219, 220 purity, criteria, 221, 222 reactions definition, 218 mechanism, 233-235 regulation, 239-242 cell cycle, 240 cell differentiation, 241, 242 hormonal stimulation, 240, 241 metabolic activation, 242 mitogen stimulation, 240 research applications, 243, 244 stoichiometry 227 substrates, 227 Polyethyleneimine precipitation, RNA polymerase, 90 Polymerization de novo synthesis oligoribonucleotide primer, 157, 158 poly(A) polymerase, 236, 237 PNPase, 527, 529-532 PNPase activators, 529, 530 de novo 'synthesis, 527, 529-532 deoxyribonucleotides, 545 elongation, 532 equilibrium, 533 substrates, modified, 533, 534

640 Polynucleotide chains, property alteration, 5, 6 Polynucleotide phosphorylase, see PNPase Pol ynucleotides PNPase-directed labeling, 553 synthesis, 539-543 heteropolymers, 541-543 homopolymers, 539-541 NDP modifications, 540, 541 Polyoma DNA elongation process, 179 supercoiled form, 139 Polyoms, specific binding, 143 Polypeptide, composition, eukaryotic RNA polymerase, 118, I19 Poly(rA).poly(rU), synthesis, 84 Poly(rG), synthesis, 86 Poly(rIC), synthesis, 84 PP, exchange reaction, 252 Postribosomal supernatant fraction, 2',5'-oligoadenylate synthetase, 288 Precipitation assay, RNase, 425 Primase, see dnaG protein Priming eukaryotic DNA replication, 178-182 multiple pathways, 160-165 dnaG-dependent , 162- I65 protein requirements, 164 poly(A) polymerase, requirements, 227229 Priming enzymes, 155-182, see also specific enzymes phage-encoded, 174-178 bacteriophage T4, 176-178 bacteriophage T7, 174-176 Progesterone, poly(A) polymerase, activity and, 241 Proline isomerization, RNase folding, 390, 391 RNase refolding, 392 substitution, amino acid sequence, 404 Protein cap binding, 259 in v i m synthesis, cytoplasmic RNase inhibitor, 423, 424 synthesis, elongation factors, see EFTU.TS Proteolysis during enzyme isolation, 123 PNPase. 526-528

SUBJECT INDEX PseT mutants, RNA ligase, 50, 51 Pseudouridine, 570-572 structure, 569 Psuedouridylate synthetase I, 570, 571 pup-Sepharose, RNase purification, 319 Purification capping enzyme, rat liver nuclei, 257, 258 cytoplasmic RNase, 417 dnaG protein, 157 DNA ligase, 8, 9 eukaryotic RNA polymerase, 1 11-1 17 activity assays, 112-1 15 column chromatography, 116, I17 nucleic acid removal, 116 polyhydric alcohols, role, 114 procedures, 115-1 17 purity, 117 solubilization, 115, I16 sources, 111-113 yields, 111-113 HeLa cell capping enzyme, 256, 257 N'-isopentenyladenosine, 575-577 2',5'-oligoadenylate synthetase, 287-289 PNPase, 520-522 poly(A) polymerases, 219-223 affinity chromatography, 220, 221 contaminating enzymes, 222-223 ion exchange, 219, 220 purity, criteria, 221, 222 pseudouridylate synthetase I, 570, 571 QP replicase, 269, 270 RNase 1, 504 RNase 11, 506, 507 RNase 111, 486, 487 RNase IV, 512, 513 RNase D, 480 RNase 0, 478 RNase P, 471, 472 RNase P2, 477, 478 RNase TI, 436, 437 PNPase, 520-522 T4 DNA, 8 , 9 T4 RNA, 33-35 T7 RNA polymerase, 89, 90 transfer RNA, 186-190 first partial, 185 purity, 188-190 source, 187 transfer RNA:guanine transglycosylases, 580

64 1

SUBJECT INDEX transfer RNA nucleotidyltransferase, 186-190 vaccinia virus capping enzyme, 249 Pyridoxal phosphate RNase, 320, 323 yeast RNA polymerase I, inhibition, 133, I34 Pyrmidine nucleotide monophosphates, RNase A binding, 377 Pyrophosphate, inhibition HeLa cell capping enzyme, 257 poly(A) polymerase, 231 Pyrophosphorol ysis messenger RNA capping, 253 transfer RNA, 21 1-213

Q QP replicase, 267-279 catalytic properties, 273-279 binding sites, 276 EF-Tu’Ts, 277, 278 elongation, 278, 279 heterologous templates, 273-275 homologous templates, 273-275 host factor, 276-277 inhibition, 278 reactions, 273 S I , 276, 277 termination, 279 properties, 270-273 molecular weight, 271 purification, 269-270 subunits, 268-273 Ef-Tu .Ts, 272, 273 identification, 270, 271 relationships, 271, 272 SI-11, 272, 273 Queuosine, 579-582 structure, 579

R R17 RNA, cleavage, RNase IV, 513, 514 Radiobinding assay, 2‘,5’-oligoadenylates, 297, 298 Radiochemical assay, 2 ’ 3‘-oligoadenylates, 293, 294 Radiolysis, RNase, 342, 343

Radioimmune assay, 2’,5’-oligoadenylates, 297, 298 Radiolabeling NDP, 547. 548 nucleotides, synthesis, 547, 548 2’,5’-oligoadenylates, 293, 294 RNA, 57, 113, 114 Rat liver nuclei, capping enzyme, 257, 258 Reconstitution eukaryotic RNA polymerase, 129, 130 RNase P, 472, 473 Reduction, RNase A, 338, 339 Refolding, see Folding pathway Renaturation eokaryotic RNA polymerase, 129, 130 PNPase, 526 Reovirus guanylyltransferase, activity, 264 Restriction enzymes, DNA digestion, 139 Reverse exchange reaction, RNA ligase, 46 Reverse transfer reaction, T4 RNA ligase, 44, 45 Ribonuclease, see specific RNase Ribosomal RNA 3‘ termini, role, 551 turnover, RNase I, 505,506 Ribothymidine, 564, 565 structure, 559 Rifamycin, inhibition eukaryotic RNA derivates, 146, 147 poly(A) polymerase, 232 cap function, in virro, 258-260 reaction sequence, 247, 248 structure, 246, 247 transcription, relationship, 262, 263 chain elongation, T7 RNA polymerase, 104

chain termination, T7 RNA polymerase, 105 cleavage, 58 double-stranded cleavage, RNase 111, 489, 490 processing, eukaryotic RNase, 497-499 heterologous templates, QP replicase, 273-275 homologous template, QP replicase, 275 ligase, see RNA ligase messenger, see Messenger RNA methylation, see Methylation 3’-OH region, regulatory role, 550-552

642 polymerases, see specific polymerases priming proteins. 177, 178 radiolabeling, 57 replication, see specific replicases sequence determination, use of RNase T I , 463-464 3’ sequences, phosphorolysis, 549 single-stranded, cleavage primary sites, 491-495 RNase 11, 508 RNase 111, 490-497 secondary sites, 495-497 substrates, phosphodiester form, 16, 17 synthesis, divalent cation presence, I12 terminal riboadenylate transferases, see Poly(A) polymerases 3’ terminus, events, 57 5’ terminus, events, 58 transfer, see Transfer RNA triphosphate terminus, 247, 248 5 S RNA, transcription, 148, 149 RNA-DNA hybrids, phosphodiester formation, 16, 17 RNA (guanine-7-)methyltransferase,vaccinia virus capping enzyme, 254-255 RNA guanylyltransferase, 245-265 purified systems, 248, 249 rat liver nuclei, 257, 258 role in vivo, 258-264 activity control, 264 cap function, 258-260 capping versus transcription, 262, 263 guanine-7 methylation, 260, 261 guanylylation, 260, 261 transguanylylation, 261, 262 vaccinia virus, see Vaccinia virus capping enzyme RNA ligase active nucleoside 3’3’-bisphosphates, 53 biological role, 47-52 mutants, 48 oligodeoxyribonucleotide joining, 56 reactions ATP-independent , 42-44 circularization, 41, 42 inhibitors, 40, 41 intermolecular forward, 36-4 I reverse exchange, 46 reverse transfer reaction, 44,45 TFA activities, 48-49

SUBJECT INDEX RNA polymerase, see also specific polymerases bacteriophage structure, 96, 97 templates, 97 priming, 166-169 radiolabeling, 73 RNA polymerase I, 126, 127 affinity labeling, 134 inhibition, 133, 134 RNA polymerase 11, 124 capping enzyme and, 262, 263 RNA triphosphatase, vaccinia virus capping enzyme, 255 RNase, see also specific RNases guanine-specific, 465-468 amino acid composition, 466 RNase I, 501-506 action, mechanism, 505 applications, 506 biological role, 505, 506 intracellular location, 503 physical properties, 504, 505 purification, 504 RNase 11, 502, 503, 506-512 action, mechanism, 508-510 degradation, processive, 509, 510 hydrolysis, 508, 509 monovalent cation, 508, 509 single-strand specificity, 508 applications, 512 biological role, 5 10-5 12 physical properties, 507 purification, 506, 507 RNase Ill, 476-477, 485-499 double-stranded RNA, cleavage, 489,490 eukaryotic cells, 497-499 molecular weight, 487 purification, 486, 487 single-stranded RNA, cleavage, 490-497 bacteriophage T7 early, 491-495 double cleavage, 494 primary sites, 491-495 30 S preribosomal, 491, 493, 494 salt concentration, 496, 497 secondary sites, 495-497 structure, 487-489 RNase IV, 502-503, 512- 515 action, mechanism, 513 applications, 5 15

SUBJECT INDEX

643

biological role, 5 15 chemical synthesis, 360 properties, 512, 513 circular dichroism, 382-384 purification, 512. 513 histidine residues, 329 RNase A structure, 322 alk ylation X-ray diffraction, 364, 365 amino groups, 323, 324 RNase T , , 435-468 histidine residues, 329-331 chemical properties, 437-446 kinetic constants, 332 amino groups, 439, 440 amidination, 329 arginine residues, 441 antigenicities, 363 carboxyl groups, 437-439 chemical shifts, 371 dissociable groups, 444, 445 chemical synthesis, 359-361 disulfide bonds, 442 circular dichroism, 382, 384 enzymatic modifications, 442, 443 conformational transition, 373, 374 functional groups, modification, 437cross-linked dimers, 327, 328 443 crystal structure. 365, 366 histidine residues, see Histidine resdeamidation, 336 idues fluorescence, 384, 385 hydrogen exchange rate, 445, 446 kinetics, 428, 429 physical parameters, 444-446 constants, 332 primary structure, 438 methionine residues, 336, 337 purification, 436, 437 NMR reactions catalyzed, 447-463 spectra, 370, 377, 378 best-fit parameters, 460 titration, 346 binding, 453-455 pK, changes, 379 catalytic specificity, 448, 449 proton resonance, 366, 367 circular dichroism, 456 pyrmidine nucleotide monophosphates, dissociation constants, 450, 451, 453binding, 377 455 reduction, 338-339 enzyme inhibition, 458 structure, 322 fluorescence, 456 U V irradiation, 343, 344 gel filtration studies, 449-452 RNase D, 479-482 k,,,, 459, 460 biological role, 482 mechanism, 461-463 hydrolysis, 48 I NMR, 457 properties, 480 pH dependence, 450, 452, 460 purification, 480 spectral data, 452-458 reaction catalyzed, 480-482 steady-state kinetics, 458-461 RNase-DNase, preparation, 328 substrate analogs, interaction, 449RNase 0, 478, 479 458 RNase P, 471-476 transesterification, 459-461 biological role research applications, 463-465 from non-E. coli organisms, 475, 476 sequence determination, 463, 464 properties, 471, 472 spectral studies, 445, 446 purification, 471, 472 synthesis, 443 reaction catalyzed, 473-475 thermal transition, 446 structure, 472, 473 trypsin residues, 442, 443 RNase P2?477-479 tryptophan residues, 441, 442, 445, 446 biological role, 478, 479 tyrosine residues, 442, 445, 446 purification, 477, 478 rT, formation, ribothymidine catalyzed, RNase S 564-565

644

SUBJECT INDEX

S SI, Qp replicase, 276, 277 SI-11, Qp replicase, 272, 273, 277, 278 S-carboxymethylation, RNase folding, 392, 393 S factor, 152 S-peptide, RNase refolding, 391 S-peptide-S-protein interaction, 345-356 activating ability, 348 aggregation, 354, 355 amino acid substitutions, 404, 405 bhding activity, 351 capacity, 348 constants, 352, 353, 355 properties, 351 calorimetric studies, 352, 353 dissociation, 353, 354 equilibrium constants, 353, 354 glutamate residues, 350, 351 hisddine residues, see Histidine residues methionine residues, 346, 347 methylhistidines, substitution, 348-350 tyrosine residues, 346 van? Hoff enthalpy, entropy, 353, 354 S-protein RNase conformational stability, 388 RNase folding, 394 28 S ribosomal RNA, 2',5'-oligoadenylates, 308 30 S preribosomal RNA, cleavage, primary sites, 491, 493, 494 30 s ribosomes, RNase 1, 503, 506 6 S RNA MDV-I elongation, 279 replication, 275, 276 Salmonella fyphimurin, RNA polymerase, 91 Schiff bases, amino groups, pancreatic RNase, 320, 323 SDS-polyacrylamide gel electrophoresis eukaryotic RNA polymerase, 120- 122 nuclease activity, 426 RNase 111, 487-489 T7 RNA polymerase, 94 Seminal plasma RNase aggregation, 413 bovine, 41 1-415 disulfide reduction, 412, 413

double-stranded substrates, 415 hydrolysis, 415 subunits, 412, 413 tyrosine residues, 413, 414 Sequence-specific binding, DNA, 143, 144 Solubilization, eukaryotic RNA polymerase, 115-1 17 SP6 bacteriophage RNA polymerase, 96 DNA sequence, 101 promoters, 101, 102 Species I RNA, secondary site cleavage, 496,497 Spin-lattice relaxation time, RNase A, 380 Stepwise solid-phase method, RNase T , , 443 Stoichiometry, poly(A) polymerase, 218, 227 Streptomycin sulfate precipitation, T7 RNA polymerase, 89 Sulfhydryl, DNA ligase, joining aetivity, 19 Sulfhydryl inhibitors. transfer RNA nucleotidyl transferase, 191, 192 Sulfhydryl intermediates, trapping, RNase refolding, 392, 393 Sulfurtransferase, 572, 573 SV40 DNA elongation process, 179 nicks, binding, 140, 141 replication, priming, 178-180 specific binding, 143 supercoiled form, 139 unnicked, transcription, 142 SV40 messenger RNA, 308

T T3 bacteriophage RNA polymerase promoters, DNA sequence, 101 purification, 90-91 T4 DNA ligase, see also DNA ligase assay, partial reaction, 7 bacteriophage, 21, 22, 25 blunt end joining, 19, 20 joining activity, 16 joining reaction, pH optimum, 18 priming enzymes, 176-178 purification, 8, 9 RNA-DNA hybrids, 16-17 T4 gene 30 mutation, 22, 23 T4 gene 63, 48

SUBJECT INDEX T4 polynucleotide kinase 3'-phosphastase, 50, 51 T4 RNA ligase, 31-58 applications, 52-58 assays, 33, 34 gene locations, 50 intermolecular forward reaction, 33, 3641 adenylyated donor formation, 37-39 enzyme-AMP formation, 36, 37 phosphodiester bond formation, 39-41 joining, stimulation, 20 nucleic acid modification, 57, 58 oligonucleotide synthesis, 53-56 phosphodiester bond formation, 39-41 physical properties, 35 properties, 33-35 protein isolation, 34 purification, 33-35 reactions catalyzed, 35-47 T5 DNA ligase, 27 T7 bacteriophage RNA polymerase promoters, 99-102 DNA sequence, 101 RNA chain termination, 105 T7 DNA as template, 70 polymerase promoters, 99-102 priming enzymes, 174-176 template, 72, 73 T7 DNA ligase bacteriophage, 23 joining reaction, pH optimum, 18 T7 early RNA bacteriophage, cleavage primary sites, 491-494 secondary sites, 495 template activity, 140 T7-like bacteriophage DNA-dependent RNA polymerase, catalytic properties, 98-105 template specificity, 97 transcriptional maps 98-102 T7 RNA polymerase amino acid composition, 94, 95 assay, 91-94 molecular weight, 94 promoter binding, 103 purification, 89, 90 RNA chain elongation, 104 synthesis, 91

645 Tail fiber attachment, RNA ligase, 48, 49 Takadiastase, 435 T antigen, eukaryotic systems, 180 TATA box, 150 TCA precipitation assay, RNA, 114, 1 I5 Template calf thymus DNA, I14 dnaG interactions, 165-167 eukaryotic RNA polymerase, 138-140 G4-RFI DNA, 171 lymphocyte DNA, 140 QP replicase, 273-275 RNA polymerase, 97 specificity, T7-like bacteriophage polymerase, 98-102 well-defined, 72, 73 TFA protein, purified, 48.49 Thiolated pyrimidines, 573 4-Thiouridine, 572, 573 structure, 569 Titration, curves, histidine, 373,374,444,445 TMV RNA, 3' sequence, 551 Transcription bacteriophage N4 RNA polymerase, 107I08

cap sites, 149. 150 criteria, 80 factors, DNA-dependent RNA polymerase, 79-81 maps, T7-like bacteriophage RNA polymerase, 98-102 nicked DNA, 140, 141 random, 73 RNA capping, relationship, 262, 263 specific versus random, 143 T7 DNA, 70, 72 unnicked DNA, 141, 142 well-defined templates, 72, 73 Transcription cycle, 85, 86 T7-like bacteriophage DNA-dependent RNA polymerase, 92 T7-like bacteriophage RNA polymerase. steps, 102-105 Transcriptor extracts, eukaryotic systems adenovirus DNA, 147, 148 discovery, 147, 148 fractionation, 148, 149 in vivo, mimicing, 149, 150 Transesterification, RNase T , , catalyzed, 459, 460

SUBJECT INDEX Transfer RNA methyltransferases, see Methyltransferase enzymes precursor molecules, interaction with processing enzymes, 470 processing enzymes, 469-483, see also specific RNase E. coli BN, 482 interaction with precursors, 470 RNase 11, 3‘ termini, 551 Transfer RNATyrprecursor, 481 RNase 11, 511 Transfer RNA:guanine transglycosylases, properties, 581 Transfer RNA nucleotidyltransferase, 183215 assay, 186 biological role, 213-215 catalytic properties, 192-213 ADP, 198 AMP, 198, 202, 205 anomalous nucleotide incorporation, 208-21 I ATP, 196, 197, 206-209 binding, 201, 202 binding constants, 28, 194, 196, 197 -C-C-A sequence, 207-210 cation effects, 193-195 CMP, 204, 205, 209, 210 CTP, 196, 197, 206-210 dead-end inhibition studies, 198 dissociation constants, 199 forward versus reverse reactions, 212 hydrolysis, transfer RNA, 21 1-213 kinetic constants, 203 kinetic mechanism, 198, 199 Mg2+, 193, 194 misincorporation, nucleotides, 210, 21 1 MnZ+, 195 model acceptors, 202-207 model, catalysis, 208, 210 nucleoside triphosphate donors, 196, 197 nucleotide incorporation, 205-208 pH effects, 195 polyamines, 195 pyrophosphorolysis, transfer RNA, 211-213 recognition, 202-207

recognition regions, 204, 205 RNA acceptors, 199-207 specificity, 199-201 transfer RNA recognition, 202-207 -C-C-A sequence regeneration, 184 chemical properties, 191, 192 metalloenzyme, 192 partial reaction, 184 physical properties, 187, 190, 191 molecular weight, 187, 190, 191 research applications, 215 specific activity, 187-190 Transguanylylation, mechanism, 261, 262 Transnucleotidation, PNPase catalyzed, 537 Transphosphorylation assay, RNase, 424-425 step, RNase catalysis, 430, 431 5’-O-Triphosphoryladenylyl(2’ ,5)adenylyl(2’,5’)adenosine, 283 Tyrosine bovine seminal plasma RNase, 413, 414 circular dichroism bands, 382-384 fluorescence, 384-385 NMR, 372, 373 proton resonances, 367 UV absorption spectra, 382 Tyrosine residues pancreatic RNase, 339-342 RNase T I , 442 S-peptide-S-protein interaction, 346 spectral studies, RNase TI, 445, 446 Trypsin residues PNPase, 527 RNase T I , 442, 443 Tryptophan residues PNPase, 523, 524 RNase T , , 441, 442 spectral studies, 445, 446

U Uridine, modification, 568-574 3-(3-amino-3-carboxypropyl)uridine,573, 574 derivatives, structures, 568-570 5-methylaminomethyl-2-thiouridine, 574 modifying enzymes, 570-574 pseudouridine, 570-572 thiolated pyrimidines, 573 4-thiouridine, 572, 573

647

SUBJECT INDEX UV absorption spectra, tyrosine residues, 382 UV difference, RNase T,, 4.52, 453 UV irradiation, pancreatic RNase, 343-345

Viral RNA, 3’ sequences, 55 1, 552 Viral-coated RNA polymerase, 152 VSV infection, interferon and, 311

w V Vaccinia virus guanyltransferase, 248-255 covalent enzyme-guanylyate intermediate, 252-254 enzymatic reactions, 250 GTP-PP, exchange, 2.52-254 mechanism, 252-254 methylation, 260, 261 molecular properties, 249, 250 purification, 249 reactions catalyzed, 250-265 research applications, 264, 265 RNA (guanine-7-)-methyltransferase, 254, 255 RNA guanylytransferase, 250-252 RNA polymerase, 152 RNA triphosphatase, 255

Wheat germ RNA polymerase RNA polymerase I, 127 RNA polymerase 11, 124-125 denaturation, 130 SDS-polyacrylamide gel electrophoresis, 120- 122 Wyeosine, 578, 579

X X-Ray diffraction, RNase, 364-366 Z

Zn2+ eukaryotic RNA polymerase, 131, 132 T7 RNA polymerase activity, 96

Contents of Other Volumes Volume k Structure and Control

X-Ray Crystallography and Enzyme Structure David Eisenberg

Chemical Modification by Active-Site-Directed Reagents Elliott Shaw

Chemical Modification as a Probe of Structure and Function Louis A . Cohen

Multienzyme Complexes Lester J . Reed mnd Duvid J . Cox Genetic Probes of Enzyme Structure Milton J . Schlesinger Evolution of Enzymes Emil L. Smith The Molecular Basis for Enzyme Regulation D . E. Koshland. J r . Mechanisms of Enzyme Regulation in Metabolism E. R . Studtman

Enzymes as Control Elements in Metabolic Regulation Daniel E. Atkinson Author Index-Subject Index

Volume II: Kinetics and Mechanism

Steady State Kinetics W. W. Cleland Rapid Reactions and Transient States Gordon B. Hammes and Puul R. Schirnmel Stereospecificity of Enzymic Reactions G. Popjak Proximity Effects and Enzyme Catalysis Thomas C . Bruice 648

CONTENTS OF OTHER VOLUMES

Enzymology of Proton Abstraction and Transfer Reactions Intin A . Rose Kinetic Isotope Effects in Enzymic Reactions J . H . Richurds Schiff Base Intermediates in Enzyme Catalysis Esmond E. SneII and Sumuel J . DiMuri Some Physical Probes of Enzyme Structure in Solution Serge N . Timushrff Metals in Enzyme Catalysis Albert S. Mildvan Author Index-Subject Index

Volume 111: Hydrolysis: Peptide Bonds

Carboxypeptidase A Jean A . Hurtsuck rind William N . Lipscomb Carboxypeptidase B J . E. Folk Leucine Aminopeptidase and Other N-Terminal Exopeptidases Robert J . DeLunge nnd Emil L. Smith Pepsin Joseph S. Fruton Chymotrypsinogen: X-Ray Structure J . Kraut The Structure o f Chymotrypsin D. M . B l a ~ i Chymotrypsin-Chemical Properties and Catalysis George P. Hess Trypsin B . Keil Thrombin and Prothrombin Staffun Mugnusson Pancreatic Elastase B. S. Hartley and D. M. Shotton Protein Proteinase Inhibitors-Molecular Aspects Michael Luskowski, Jr., and Robert W. Sealock Cathepsins and Kinin-Forming and -Destroying Enzymes Lowell M . Greenbaum Papain, X-Ray Structure J . Drenth, J . N . Jansonius, R . Koekoek, and B . G. Wolthers Papain and Other Plant Sulfhydryl Proteolytic Enzymes A . N. Glazer and Emil L . Smith Subtilisin: X-Ray Structure J . Kraut Subt ilisins: Primary Structure, Chemical and Physical Properties Francis S. Markland. Jr., und Emil L . Smith

649

CONTENTS OF OTHER VOLUMES

Streptococcal Proteinase Teh- Yung Liu wid S. D. Elliott The Collagenases Sam Seifter orid Elvin Hlirper Clostripain William M . Mitchell and William F. Harrington Other Bacterial, Mold, and Yeast Proteases Hiroshi Matsubarri and Joseph Feder Author I nde x-S ubject Index

Volume IV: Hydrolysis: Other C-N Bonds, Phosphate Esters

Ureases F. J . Reithel Penicillinase and Other P-Lactamases Nathun Citri Purine, Purine Nucleoside, Purine Nucleotide Aminohydrolases C. L . Zielke and C. H . Suelter Glutaminase and y-Glutamyltransferases Stundish C. Hurtmuti L- Asparaginase John C. Wriston, Jr. Enzymology of Pyrrolidone Carboxylic Acid Marian Orlowski and Alton Meister Staphylococcal Nuclease X-Ray Structure F. Albert Cotton and Edward E. Hazen, Jr. Staphylococcal Nuclease, Chemical Properties and Catalysis Christian B. Anfinsen. Pedro Cuatrecasas, und Hiroshi Einiuchi Microbial Ribonuclease with Special Reference to RNases T,,T2,N,, and Uz Tsuneko Uchida and Fujio Egami Bacterial Deoxyribonucleases 1. R. Lehmcin Spleen Acid Deoxyribonuclease Giorgio Bernardi Deoxyribonuclease I M . Laskowski, Sr. Venom Exonuclease M. Laskoliiski, Sr. Spleen Acid Exonuclease Albert0 Bernnrdi und Giorgio Bern(irdi Nucleotide Phosphomonoesterases George I . Drummond and Mascinobu Yumrrmoto Nucleoside Cyclic Phosphate Diesterases George I. Drummotid cind M(isrinobi4 Y m w n o t o E. coli Alkaline Phosphatase Ted W. Reid and Initin 6 . Wilson

CONTENTS OF OTHER VOLUMES

65 1

Mammalian Alkaline Phosphatases H. N. Ferti1e.v Acid Phosphatases Vitirent P. H o Ilutider Inorganic Pyrophosphatase of Escherichiu coli John Josse rind Simon C. K. Wong Yeast and Other Inorganic Pyrophosphatases Lurry G. Butler Glucose-6-Phosphatase, Hydrolytic and Synthetic Activities Robert C. Nordlie Fructose- 1,6-Diphosphatases S . Pontremoli arid B . L . Horecker Bovine Pancreatic Ribonuclease Frederic M. Richnrds and Hurold W. Wvckofl Author Index-Subject Index

Volume V: Hydrolysis (Sulfate Esters, Carboxyl Esters, Glycosides), Hydration

The Hydrolysis of Sulfate Esters A . B. Roy Arylsulfatases R . G. Nicholls und A . B. Roy Carboxylic Ester Hydrolases Klaus Krisch Phospholipases Donald J . Hunuhun Acetylcholinesterase Hnrty C. Froede cind lnvin B. Wilson Plant and Animal Amylases John A . Thomus, Joseph E. Sprudlin, und Stephen Dygert Glycogen and Starch Debranching Enzymes E . Y. C. Lee and W. J . Whelan Bacterial and Mold Amylases Toshio Takagi, Hiroko Todu, and Toshizo lsemura Cellulases D.R . Whitaker Yeast and Neurospora Invertases J . Oliver Lunipen H yaluronidases Kurl Me-yer Neuraminidases Alfred Gottschalk and A . S.Bhurguvu Phage Lysozyme and Other Lytic Enzymes Akira Tsugita Aconitase Jenny Pichvorth Glusker

652

CONTENTS OF OTHER VOLUMES

/3-Hydroxydecanoyl Thioester Dehydrase Konrad Bloch Dehydration in Nucleotide-Linked Deoxysugar Synthesis L. Glaser and H . ZarkowsAy Dehydrations Requiring Vitamin B,, Coenzyme Robert H. Abeles Enolase Finn Wold Fumarase and Crotonase Robert L . Hill and John W. Teipel 6-Phosphogluconic and Related Dehydrases W. A. Wood Carbonic Anhydrase S. Lindskog, L . E. Henderson, K . K . Kannan, A . Liljas, P. 0. Nyman, and B. Strandberg Author Index-Subject Index

Volume VI: Carboxylation and Decarboxylation (Nonoxidative), lsomerization

Pyruvate Carboxylase Michael C. Scrutton and Murray R. Young Ac yl-CoA Carboxylases Alfred W. Alberts and P. Roy Vagelos Transcarbox yIase Harland G . Wood Formation o f Oxalacetate by COz Fixation on Phosphoenolpyruvate Merton F. Utter and Harold M . Kolenbrander Ribulose- 1,S-Diphosphate Carboxylase Marvin 1. Siegel, Marcia Wishnick, and M . Daniel Lane Ferredoxin-Linked Carboxylation Reactions Bob B. Buchanan Amino Acid Decarboxylases Elizabeth A. Boeker and Esmond E. Snell Actoacetate Decarboxylase Irwin Fridovich Aldose-Ketose Isomerases Ernsf A. Noltmann Epimerases Luis Glaser Cis-Trans Isomerization Stunley Seltzer Phosphomutases W. J. Ray, Jr., and E. J . Peck, Jr. Amino Acid Racemases and Epimerases Elijah Adams

CONTENTS OF OTHER VOLUMES

65 3

Coenzyme B,,-Dependent Mutases Causing Carbon Chain Rearrangements H. A . Barker BI2 Coenzyme-Dependent Amino Group Migrations Thressa C . Stadrman

IsopentenylpyrophosphateIsomerase P. W. HoNoI\ 'ay Isomerization in the Visual Cycle Jorum Heller

AS-3- Ketosteroid Isomerase Paul Tululuy und Anri M. Berison

Author Index-Subject

Index

Volume VII: Elimination and Addition, Aldol Cleavage and Condensation, Other C-C Cleavage, Phosphorolysis, Hydrolysis (Fats, Glycosides)

Tryptophan Synthetase Charles YunofsX? und Irving P. Crtrnfiford Pyridoxal-Linked Elimination and Replacement Reactions Leodis Duvis and David E. Metzler The Enzymatic Elimination of Ammonia Kenneth R. Hunson und Evelyn A . Huvir

Argininosuccinases and Adenylosuccinases Sarah Riitner

Epoxidases William B. Jukoby und Thorsteri A. fiellstedt Aldolases B. L. Horecker, Orestes Tsolus, und C. Y. Lrii Transaldolase Orestes Tsolus and B. L. Horecker 2- Keto-3-deoxy-dphosphogluconicand Related Aldolases W. A. Wood Other Deoxy Sugar Aldolases David Sidney Feingold und Putricia Anti Hofee

SAminolevulinic Acid Dehydratase David Shemin

8-Arninolevulinic Acid Synthetase Peter M . Jordun und David Shemin

Citrate Cleavage and Related Enzymes Leonard B. Spector

Thiolase Ulrich Grhring und Feodor Lynrn

Acyl-CoA Ligases Malcolm J . P. Higgins, Jack A . Kornblutt, w i d H u r v Rudney a-Glucan Phosphorylases-chemical and Physical Basis of Catalysis and Regulation Donald J . Gruves and Jerry H . Wung

CONTENTS OF OTHER VOLUMES

Purine Nucleoside Phosphorylase R . E. Purks, Jr., und R . P. Agunvul Disaccharide Phosphorylases John J . Mieyul and Robert H . Abeles Pol ynucleotide Phosphorylase T. Godefrqv-Colburn und M . Grunberg-Manugo The Lipases P. Desnuelle P-Galactosidase Kurt Wullenfels und Rudolf Weil Vertebrate Lysozyrnes Tuiji Imoto, L . N.Johnson, A . C. T. North. D. C . Phillips, und J . A . Rupley Author Index-Subject Index

Volume VIII: Group Transfer, Part A: Nucleotidyl Transfer, Nucleosidyl Transfer, Acyl Transfer, Phosphoryl Transfer

Adenylyl Transfer Reactions E. R . Studtmun Uridine Diphosphoryl Glucose Pyrophosphorylase Richard L. Turnquist and R . Guurth Humeri Adenosine Diphosphoryl Glucose Pyrophosphorylase Jack Preiss The Adenosyltransferases S. Harvey Mudd Acyl Group Transfer (Acyl Carrier Protein) P. Roy Vugelos Chemical Basis of Biological Phosphoryl Transfer S. J. Benkovic und K . J . Schruy Phosphofructokinase David P. Bloxhum und H e n n A . Lurdy Adenylate Kinase L . Nodu Nucleoside Diphosphokinases R . E. Purks, Jr.. and R. P. Agurwul 3-Phosphoglycerate Kinase R . K . Scope Pyruvate Kinase F. J . Kuyne Creatine Kinase (Adenosine 5'-Triphosphate-Creatine Phosphotransferase) D. C. Watts Arginine Kinase and Other Invertebrate Guanidino Kinases J . F. Morrison Glycerol and Glycerate Kinases Jeremy W . Thorner und H e n n Puulus Microbial Aspartokinases Puolo Truffu-Buchi

CONTENTS OF OTHER VOLUMES

655

Protein Kinases Donul A . Wulsh and Edwin G . Krebs Author Index-Subject Index

Volume IX: Group Transfer, Part 6: Phosphoryl Transfer, One-Carbon Group Transfer, Glycosyl Transfer, Amino Group Transfer, Other Transferases

The Hexokinases Sidney P. Colowick Nucleoside and Nucleotide Kinases Elizubeth P. Anderson Carbamate Kinase L. Ruijrnun und M . E. Jones

N5-Methyltetrahydrofolate-HomocysteineMethyltransferases Robert T. Tuylor und Herbert Weisshuch Enzymic Methylation of Natural Polynucleotides Splviu J . Kerr und Ernest Boreh Folate Coenzyme-Mediated Transfer of One-Carbon Groups Jeunne I. Ruder and F. M . Huennekens Aspartate Transcarbamylases Gury R . Jwobson und George R. SturX Glycogen Synthesis from UDPG W. Stulmuns und H.G. Hers Lactose Synthetase Kurt E. Ebner Amino Group Transfer Alexander E. Bru unstein Coenzyme A. Transferases W. P. Jenrhs Amidinotransferases Jumes B. WuIAer N - Acet ylglutamate-5-Phosphotransferase Gezu Denes Author Index-Subject Index

Volume X: Protein Synthesis, DNA Synthesis and Repair, RNA Synthesis, Energy-linked ATPases, Synthetases

Polypeptide Chain Initiation Severo Ochou und Rujarshi Muzumder Protein Synthesis-Peptide Chain Elongatian Jeun Lucus-Lenurd und Laszlo Beres Polypeptide Chain Termination W. P. Tute und C . T. Cuskey Bacterial DNA Polymerases Thornus Kornberg und Arthur Kornberg

656

CONTENTS OF OTHER VOLUMES

Terminal Deoxynucleotidyl Transferase F. J . Bollum Eucaryotic DNA Polymerases Lawrence A . Loeb RNA Tumor Virus DNA Polymerases Howard M . Temin and Satoshi Mizutuni DNA Joining Enzymes (Ligases) I. R . Lehmun Eucaryotic RNA Polymerases Pierre Chumbon Bacterial DNA-Dependent RNA Polymerase Michael J . Chamberlin Mitochondria1 and Chloroplast ATPases Harvey S. Penefshy Bacterial Membrane ATPase Adolph Abrums and Jeffrey B. Smith Sarcoplasmic Membrane ATPases Wilhrlm Husselbach Fatty Acyl-CoA Synthetases John C. Londesborough und Leslie T. Webster, J r . Aminoacyl-tRNA Synthetases Dieter Sol1 and Puul R . Schimmel CTP Synthetase and Related Enzymes D.E. Koshlund, Jr., and A . Levitzki Asparagine Synthesis Alton Meister Succinyl-CoA Synthetase Williium A . Bridger Phosphoribosylpyrophosphate Synthetase and Related Pyrophosphokinases Robert L . Switzer Phosphoenolpyruvate Synthetase and Pyruvate, Phosphate Dikinase R. A . Cooper und H . L. Kornberg Sulfation Linked to ATP Cleavage Harry D. Peck, Jr. Glutathione Synthesis Alton Meister Glutamine Synthetase of Mammals Alton Meister The Glutamine Synthetase of Escherichiu coli: Structure and Control E. R. Studtmun and A . Cinsburg Author Index-Subject Index

Volume XI: Oxidation -Reduction, Part A: Dehydrogenases (I), Electron Transfer ( I )

Kinetics and Mechanism of Nicotinamide-Nucleotide-LinkedDehydrogenases Keith Dalziel

CONTENTS O F OTHER VOLUMES

657

Evolutionary and Structural Relationships among Dehydrogenases Michael C .Rossmunn. Antlers Liljus. Curl-hur Briinden. rind Leonrrrtl J . Bunusz.uk Alcohol Dehydrogenases Curl-lvur Briinden. Huns Jiirnvull. Huns Eklunti. und Bo Furugren Lactate Dehydrogenase J . John Holbrook. Anders Li!iu.s, Steven J . Steindel. und Michuel C . Rossmunn Glutamate Dehydrogenases Emil L . Smith, Briuti k1. Austen, Kenneth M . Blumenthrrl, untl Joseph F. Nyc Malate Dehydrogenases Leonard J . Bonuszuk und Ralph A . BrudshuLi) Cytochromes c Richurd E. Dickerson m d Russell Timkovich Type b Cytochromes Bunji Hugihuru. Nobuhiro Stito. unrl Tuteo Yumrrnuku Author Index-Subject Index

Volume XII: Oxidation-Reduction, Par?B: Electron Transfer (ll),Oxygenoses, Oxidases (I)

Iron-Sulfur Proteins Gruhum Pulmer Flavodoxins and Electron-Transferring Flavoproteins Stephen G. Mayhew und Martha L . Ludwig Oxygenases: Dioxygenases Osumu Huvuishi, Mitsirhiro Nozuki, und Mitchel T. Abbott Flavin and Pteridine Monooxygenases Vincent Mussey unil Peter Hemmerich Iron- and Copper-Containing Monooxygenases V. Ullrich und W. Dirppel Molybdenum Iron-Sulfur Hydroxylases and Related Enzymes R . C. Bruy Flavoprotein Oxidases Hurold J. Bright und Duvid J . T. Porter Copper-Containing Oxidases and Superoxide Dismutase B. G. Mulmstriim, L.-E. Andreussoti, und B. Reinhummur Author Inde x-Subjec t Index

Volume XIII: Oxidation-Reduction Port C

Glyceraldehyde-3-phosphate Dehydrogenase J . Ieuuri Hurris und Michuel Wuters Nicotinamide Nucleotide Transhydrogenases J . Rydstrom, J . B. Hoek, und L . Ernster

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CONTENTS OF OTHER VOLUMES

Flavin-Containing Dehydrogenases Churles H. Williams, Jr. Metal-Containing Flavoprotein Dehydrogenases Youssef Hnteji rind Diana L. Stiggoll Cytochrome c Oxidase Winslow S. Caughey, William J . Wulluce. John A . Volpe, t i n d Shitzyu Yoshikciwu Cytochrome c Peroxidase Takashi Yotietmii Catalase Gregon R. Schonbaum untl Britton Chunce Author Inde x-Subjec t Index Index for Volumes I-XI11

Volume XIV: Nucleic Acids, Part A

SECTION I. DNA POLYMERASES A N D RELATED ENZYMES DNA Polymerases-A Perspective Arthur Kornberg DNA Polymerase I of Escherichici coli I. Robert Lehman DNA Polymerase I11 Holoenzyme Charles MeHenry and Arthur Kornberg T-Phage DNA Polymerase I. Robert Lehman Cellular and Viral-Induced Eukaryotic Polymerases Arthur Weissbach Reverse Transcriptase Inder M. Vrrma Terminal Deoxynucleotidyltransferase Robert Ratlif SECTION 11. DNA NUCLEASES A N D RELATED ENZYMES Deoxyribonucleases: Survey and Perspectives Stuart Linn Type I Restriction Enzymes Brian Endlich and Stuart Linn Type I1 Restriction Enzymes Robert D. Wells, Ronald D. Kleiti, and C. K . Singleton Endonucleases Specific for Single-Stranded Polynucleotides I. Robert Lehtnun. Exodeoxyribonucleases of Escherichici coli Bernard Weiss recBC-like Enzymes: Exonuclease V Deoxyribonucleases Karen M. Telunder Muskavitch und Stuart Linn Enzymes That Incise Damaged DNA Errol C. Friedberg, Tllomus Bonura, Eric H . Rudany, und Jack D . Love

CONTENTS OF OTHER VOLUMES

Pancreatic DNase Stanford Moore

III. DNA MODIFICATION SECTION Bacteriophage T4 Polynucleotide Kinase Charles C. Richurdson Eukaryotic DNA Kinases Steven B. Zimtnerman und Burbaru H . Pheiffer Type I DNA Topoisomerases Jumes C. Wung DNA Gyrase and Other Type I1 Topoisornerases Murtin Gellert DNA Unwinding Enzymes Malcolm L. G e f e r Single-Stranded DNA Binding Proteins Stephen C. Kowiilczykowski, David G . Beur, and Peter H . von ffippel The recA Enzyme of Escherichirr coli and Recombination Assays Kevin McEntee und George M . Weinstock Site-Specific Recombination Protein of Phage Lambda Howurd A. Nash Photoreactivating Enzymes Betsy M . Siitherlund DNA Methylation Stunley Huttman DNA Base-Insertion Enzymes (Insertases) Zvi Livtieh utid Joseph Sperlitig DNA Glycosylases Bruce K . Duncan Author Index-Subject Index

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