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

.

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

.

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 . . . . . .

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

183

. . . . . . . . . . . . . . . . . . . 186 . . . . . . . . . . . . . . . . . . . 192

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

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

.

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

Contents of Other Volumes

This Page Intentionally Left Blank

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


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


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


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


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


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


(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.

DNA LIGASES

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.

1.

DNA LIGASES

21

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.

22


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.

I.

DNA LIGASES

23

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 .

24


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.

26

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.

DNA LIGASES

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.

28


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.

DNA LIGASES

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.


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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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


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


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


(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


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


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.

56


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.

2. T4 RNA LIGASE

57

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.

58


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


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

62

MICHAEL J. CHAMBERLIN

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

63

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.

64

MICHAEL J. CHAMBERLIN

(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

65

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.

66

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 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


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.


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.


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

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.

284

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.

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(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.

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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.

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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.

11.


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.

296

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.

1 I. 2’,5‘-OLIGOADENYLATE SYNTHETASE

297

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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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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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.

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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-

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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.

1I.

305

2’3’-OLIGOADENYLATE SYNTHETASE

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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2’,5’-OLIGOADENYLATESYNTHETASE

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.

312

L. ANDREW BALL

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.

1 I.


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.


Section 111

RNA Nucleases and Related Enzymes


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


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.


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


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.


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 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


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


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


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


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


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


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


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


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 9

38 2.2 1.8 2.9 4.6 7.3 8.1 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


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


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


(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


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.


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


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


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


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.


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.


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


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.


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


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.


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.


535

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


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.


537

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


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.


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


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


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.


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


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.


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.


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.


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.


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


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.


Section IV

RNA Modzficatim


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

558

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


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


(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


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


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


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.


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


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.


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


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


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.


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


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.


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


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


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


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


/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


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


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


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


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

658


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


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

659


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

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

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

Nucleic Acids

Quadruplex Nucleic Acids

Nucleic Acids Isolation Methods

Peptide Nucleic Acids

Extracellular Nucleic Acids (Nucleic Acids and Molecular Biology)

Nucleic Acids (Tutorial Chemistry Texts)

Molecular Modeling of Nucleic Acids

Nucleic Acids (Tutorial Chemistry Texts)

Nucleic Acids Hybridization: Modern Applications

Nucleic Acids (Tutorial Chemistry Texts)

Nucleic Acids in Innate Immunity

The Chemical Biology of Nucleic Acids

Ribonucleases (Nucleic Acids and Molecular Biology 26)

Nucleic Acids: Structures, Properties, and Functions

Nucleic Acids from A to Z

Advanced Organic Chemistry of Nucleic Acids

Nucleic Acids in Chemistry and Biology

Nucleic Acids: Structures, Properties, and Functions

Interplay between Metal Ions and Nucleic Acids

Peptide Nucleic Acids: Protocols and Applications

Peptide Nucleic Acids. Methods and Protocols

Mass Spectrometry of Nucleosides and Nucleic Acids

Functional Nucleic Acids for Analytical Applications

The Enzymes, Vol XVIII: Control by Phosphorylation, Part B (Specific Enzymes), 3rd Edition

Capillary Electrophoresis of Nucleic Acids. Introduction to the Capillary Electrophoresis

Problems of Market Liberalism: Volume 15, Social Philosophy and Policy, Part 2 (Vol 15, Pt.2)

The Grantville Gazette Vol. 15

Guitar - Play Along Vol 15 ~ R&B

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

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

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

Nucleic Acids

Quadruplex Nucleic Acids

Nucleic Acids Isolation Methods

Peptide Nucleic Acids

Extracellular Nucleic Acids (Nucleic Acids and Molecular Biology)

Nucleic Acids (Tutorial Chemistry Texts)

Molecular Modeling of Nucleic Acids

Nucleic Acids (Tutorial Chemistry Texts)

Nucleic Acids Hybridization: Modern Applications

Nucleic Acids (Tutorial Chemistry Texts)

Nucleic Acids in Innate Immunity

The Chemical Biology of Nucleic Acids

Ribonucleases (Nucleic Acids and Molecular Biology 26)

Nucleic Acids: Structures, Properties, and Functions

Nucleic Acids from A to Z

Advanced Organic Chemistry of Nucleic Acids

Nucleic Acids in Chemistry and Biology

Nucleic Acids: Structures, Properties, and Functions

Interplay between Metal Ions and Nucleic Acids

Peptide Nucleic Acids: Protocols and Applications

Peptide Nucleic Acids. Methods and Protocols

Mass Spectrometry of Nucleosides and Nucleic Acids

Functional Nucleic Acids for Analytical Applications

The Enzymes, Vol XVIII: Control by Phosphorylation, Part B (Specific Enzymes), 3rd Edition

Capillary Electrophoresis of Nucleic Acids. Introduction to the Capillary Electrophoresis

Problems of Market Liberalism: Volume 15, Social Philosophy and Policy, Part 2 (Vol 15, Pt.2)

The Grantville Gazette Vol. 15

Guitar - Play Along Vol 15 ~ R&B

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