Enzymes, Volume IV: Hydrolysis: Other C-N Bonds. Phosphate Esters. Third Edition

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The Enzymes VOLUME IV

HYDR OLYSIS Other C-N Bonds Phosphate Esters Third Edition

CONTRIBUTORS CHRISTIAN B. ANFINSEN ALBERT0 BERNARDI

I. R. LEHMAN ALTON MEISTER

GIORGIO BERNARDI LARRY G. BUTLER NATHAN CITRI

ROBERT C. NORDLIE MARIAN ORLOWSKI S. PONTREMOLI

F. ALBERT COTTON

TED W. REID F. J. REITHEL

PEDRO CUATRECASAS GEORGE I. DRUMMOND FUJI0 EGAMI H. 9.FERNLEY STANDISH C. HARTMAY EDWARD E. HAZEN, J R . VISCENT P. HOLLANDER B. L. HORECKER JOHN JOSSE M. LASKOWSKI, SR.

FREDERIC M. RICHARDS C. H. SWELTER HIROSHI TANIUCHI TSUNEKO UCHIDA IRWIN B. WILSON SIMON C. K. WONG JOHN C. WRISTON, JR. HAROLD W. WYCKOFF MASANOBU YAMAMOTO

c. I. ZIELKE ADVISORY BOARD C. B. ANFINSEN I. R. LEHMAN

ALTON MEISTER STANFORD MOORE

THE ENZYMES Edited by PAUL D . BOYER Molecular Biology Institute and Department of Chemistry University of California Los Angeles, California

Volume IV HYDROLYSIS Other C-jV Bonds Phosphate Esters

T H I R D EDITION

A C A D E M I C P R E S S New York and London

1971

COPYRIGHT 0 1971, BY ACADEMIC PRESS, INC. ALL RIGHTS RESERVED NO PART OF THIS BOOK MAY BE REPRODUCED IN ANY FORM, BY PHOTOSTAT, MICROFILM, RETRIEVAL SYSTEM, OR ANY OTHER MEANS, WITHOUT WRITTEN PERMISSION FROM THE PUBLISHERS.

ACADEMIC PRESS,

M C . 1 1 1 Fifth Avenue, New York, New York 10003

United Kingdom Edition published by ACADEMIC PRESS, INC. (LONDON) Berkeley Square House, London WlX 6BA

LIBRARY OF

CONGRESS CATALOG CARD

LTD.

NUMBER: 75 - 1 1 7107

PRINTED IN THE UNITED STATES OF AMERICA

Contents xi xv

Lisl oj Contributors

P rejuce Contents of Other Volumes 1.

xvii

Ureaser

F. J. REITHEL I. Introduction 11. Isolation and Purification of Jack Bean Urease 111. Molecular Properties IV. Ureases from Other Sources V. Catalytic Properties VI. Summary

2.

1 2

5 13 15 20

Penicillinase and Other p-lactarnares NAT'IAN

CITRI

23 27

I. Introduction 11. Molecular Properties 111. Catalytic Properties IV. Conformation and Function V. Immunological Studies

35

44 46

3. Purine, Purine Nucleoside, and Purine Nucleotide Aminohydrolases C. L. ZIELKE AND C. H. SVELTER I. Introduction 11. Adenine Arninohydrolase 111. Adenosine Aminohydrolasc. IV. 5'-Adenylic Acid Aminohydrolase V. Adenine Nucleoside and Nucleotide Aminohydrolase (Nonspecific) VI. Guanine Aminohydrolase VII. Guanosine Aminohydrolase V

47 51 54

64 73 76 77

vi

CONTENTS

4. Glutaminase and y-Glutamyltransferawr

STANDISH C. HARTMAN I. Introduction 11. Glutaminase of Escherichia coli 111. Other Glutaminases and Glutamyltransferase IV. Concluding Remarks

5.

79

80 93 98

L-Asparaginase

JOHN C. WRISTON,JR. I. Introduction 11. Occurrence 111. Guinea Pig Serum Asparaginase IV. Escherichin coli Asparaginase V. Other Asparaginases VI. Physiological Properties

6.

101 102 105 107 116 117

Enzymology of Pyrrolidone Carboxylic Acid

MARIANORLOWSKIAND ALTON,MEISTER I. Introduction 11. Detection and Determination of Pyrrolidone Carboxylic Acid 111. Natural Occurrence of Pyrrolidone Carboxylic Acid IV. Nonenzymic Formation of Pyrrolidone Carboxylic Acid from Glutamic Acid, Glutamine, and Other Compounds V. Enzymic Formation of Pyrrolidone Carboxylic Acid from Glutamic Acid VI. Enzymic Formation of Pyrrolidone Carboxylic Acid from Glutamine and Glutaminyl Peptides VII. Enzymic Formation of Pyrroline Carboxylic Acid froni y-Glutamyl Amino Acids VIII. Enzymic Formation of Derivatives of Pyrrolidone Carboxylic Acid IX. Pyrrolidone Carboxylyl Peptidase X. Pyrrolidone Carboxylate Metabolism

7.

124 125 127 130 133 139 142 146 147 149

Staphylococcal Nuclease X-Ray Structure

F. ALBERTCOTTON A N D EDWARDE. HAZEN,JR. I. Introduction 11. The Conformation of the Peptide Chain 111. The Binding of Thyniidine-3’,5-Diphosphate and Calcium Ion IV. Some Correlation Studies in Solution V. Some Tentative Comments on Mechanism and Plans for Future Studies

153 159 163 172 174

CONTENTS

8.

vii

Staphylococcal Nuclease, Chemical Properties and Catalysis

CHRISTIAN B. ANFINSEN,PEDROCUATRECASAS, AND HIROSHI TANIUCHI I. Introduction 11. Isolation 111. Covalent Structurc IV. Behavior in Solution V. Substrate Specificity and Catalytic Mechanisms VI. Stereochemical Probes of the Active Site VII. Coniplementation of Fragments VIII. Synthetic Analogs

9.

177 178 180 183 185 195 196 199

Microbial Ribonucleaser with Special Reference to RNarer TI, T2, N,, and U,

TSUNEKO UCHIDA AND FUJIOEGAMI I. Introduction 11. Fungal RNases TI, T,,N1, U,, and U, 111. Other Microbial RNases of Special Interest IV. List of Microbial RNases

10.

205 208 239 248

Bacterial Deoxyribonuclearer

I. R. LEHMAN I. Introduction 11. Exonucleases 111. Endonucleases 1 1.

25 1 252 259

Spleen Acid Deoxyribonucleare

GIORGIO BERNARDI I. Introduction 11. Physical and Chemical Properties 111. Catalytic Properties IV. Distribution, Intracellular Localization, and Biological Role

12.

27 1 272 276 285

Deoxyribonuclease I

>I LASKOWSKI, . SR. I. Introduction 11. Chemical Nature 111. Active Center IV. Inhibitor

289 292 297 299

CONTENTS

viii V. Ions

VI. Kinetica VII. Specificity VIII. Physiological Role

302 303 30% 310

13. Venom Exonuclease

M. LASKOWSKI, SR. I. Introduction 11. Chemical Nature of the Enryme 111. Structural Characteristics of Substrates AIfecting Susceptibility IV. Venom Exonuclease as a Tool for Structural Determination V. Other Venom Enzymes That Hydrolyze Phosphate Esters

14.

313 317 319 324 328

Spleen Acid Exonuclease

ALBERTO BERNARDI AND GIORGIO BERNARDI I. Introduction 11. Isolation, Purity, and Physical Properties 111. Catalytic Properties IV. Distribution and Intracellular Localization

15.

329 330 331 336

Nucleotide Phorphomonoesterases

GEORGE I. DRUMMOND AND MASANOBU YAMAMOTO I. 5’-Nucleotidase 11. 3’-Nucleotidase

16.

337 352

Nucleoside Cyclic Phosphate Diesterases

GEORGE I. DRUMMOND AND MASANOBU YAMAMOTO I. Introduction 11. Ribonucleoside 2’,3’-Cyclic Phosphate Diesterase with 3’-Nucleotidase Activity from Microorganisms 111. Ribonucleoside 2’,3‘-Cyclic Phosphate Diesterase from Vertebrate Nerve IV. Nucleoside 3’,5’-Cyclic Phosphate Diesterase 17.

355

356 363 365

E. coli Alkaline Phorphatase

TEDW. REIDAND IRWINB. WILSON I. Introduction 11. Molecular Properties 111. Catalytic Properties

373 377 392

CONTENTS

18.

ix

Mammalian Alkaline Phosphaturer

H. N. FERNLEY I. Introduction 11. Molecular Properties 111. Catalytic Properties IV. Mechanism of Enzymic Action 19.

417 422 428 443

Acid Phorphataser

VINCENTP. HOLLANDER I. Introduction 11. Prostatic Acid Phosphatase 111. Red Cell Acid Phosphatase IV. Liver Acid Phosphatase V. Spleen Acid Phosphatase VI. Acid Phosphatase in Serum VII. Miscellaneous Sources

20.

450 455 477

484 493 495 496

Inorganic Pyrophosphatase of Elcherichia coli

JOHN JOSSEAND SIMONC. K. WONG I. Introduction 11. Molecular Properties 111. Catalytic Properties IV. Conclusions

21.

499 501 518 528

Yeast and Other Inorganic Pyrophosphataser

LARRY G. BUTLER I. Introduction 11. Yeast Inorganic Pyrophosphatase 111. Other Inorganic Pyrophosphatases

22.

529 530 539

Glucose-6-Phosphatase, Hydrolytic and Synthetic Activities

ROBERTC. NORDLIE I. Introduction 11. Molecular Properties 111. Catalytic Properties IV. Metabolic Roles and Regulation, in V i m .4ppendix

543 553 565 596

800

CONTENTS

X

23.

F~ctose-1 ,bDiphosphatases

S. PONTREMOLI AND B. L. HORECKER I. Introduction 11. Liver FDPase 111. Kidney FDPase IV. Muscle FDPase V. Fructosediphosphatase of Candida utilis VI. FDPases in Other Microorganisms VII. FDPases in Higher Plants and Blue-Green Algae VIII. Summary and Conclusions

24.

612 616 629 632 635 638 640 644

Bovine Pancreatic Ribonuclease

FREDERIC M. RICHARDS AND HAROLD W. WYCKOFF I. Introduction 11. Isolation and Chromatography 111. Structure IV. Modification of Covalent Structure V. Molecular Properties VI. Catalytic Properties

647 649 653 669 705 746

Author Indez

807

Subject Indez

853

List

of Contributors

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

CHRISTIAN B. ANFINSEN (177) , Laboratory of Chemical Biology, National Institute of Arthritis and Metabolic Diseases, National Institutes of Health, Bethesda, Maryland ALBERT0 BERNARDI (329), Section on Molecular Genetics, National Institute of Child Health and Human Development, National Institutes of Health, Bethesda, Maryland GIORGIO BERNARDI (271, 329), Laboratoire de GhQtique MolCculaire, Institut de Biologie MolQculaire, Facult6 des Sciences, Paris, France LARRY G . BUTLER (5291, Department of Biochemistry, Purdue University, Lafayette, Indiana NATHAN CITRI (23), Institute of Microbiology, The Hebrew University, Hadassah Medical School, Jerusalem, Israel F. ALBERT COTTON (153), Department of Chemistry, Massachusetts Institute of Technology, Cambridge, Massachusetts PEDRO CUATRECASAS (177), Laboratory of Chemical Biology, National Institute of Arthritis and Metabolic Diseases, National Institutes of Health, Bethesda, Maryland GEORGE I. DRUMMOND (337, 355), Department of Pharmacology, University of British Columbia, Vancouver, Canada F U J I 0 EGAMI (205), Department of Biophysics and Biochemistry, Faculty of Science, The University of Tokyo, Hongo, Tokyo, Japan

H. N. FERNLEY (4171, Department of Biochemistry, Institute of Orthopaedics, Stanmore, Middlesex, United Kingdom xi

xii

LIST OF CONTRIBUTORS

STANDISH C. HARTMAN (79), Department of Chemistry, Boston University, Boston, Massachusetts EDWARD E. HAZEN, J R . (153), Department of Chemistry, Massachusetts Institute of Technology, Cambridge, Massachusetts VINCENT P. HOLLANDER (449), Research Institute for Skeletomuscular Diseases, Hospital for Joint Diseases and Medical Center, New York, New York B. L. HORECKER (611), Department of Molecular Biology, Division of Biological Sciences, Albert Einstein College of Medicine, Bronx, New York JOHN JOSSE (499), Syntex Institute of Molecular Biology, Palo Alto, California

M. LASKOWSKI, SR. (289, 313), The Laboratory of Enzymology, Roswell Park Memorial Institute, Buffalo, New York I. R. LEHMAN (251), Department of Biochemistry, Stanford University School of Medicine, Stanford, California ALTON MEISTER (123), Department of Biochemistry, Cornell University Medical College, New York, New York ROBERT C. NORDLIE (543), Guy and Bertha Ireland Laboratory, Department of Biochemistry, University of North Dakota Medical School, Grand Forks, North Dakota MARIAN ORLOWSKI (123), Department of Biochemistry, Cornell University Medical College, New York, New York

S. PONTREMOLI (611), Institute of Biological Chemistry, University of Ferrara, Ferrara, Italy

TED W. REID* (373), Department of Chemistry, University of Colorado, Boulder, Colorado F. J . REITHEL ( l ) , Department of Chemistry, University of Oregon, Eugene, Oregon FREDERIC M. RICHARDS (647), Department of Molecular Biophysics and Biochemistry, Yale University, New Haven, Connecticut

* Present address : Departments of Ophthalmology, Molecular Biophysics, and Biochemistry, Yale University School of Medicine, New Haven, Connecticut.

LIST O F CONTRIBUTORS

xiii

C. H. SUELTER (47), Department of Biochemistry, Michigan State University, East Lansing, Michigan HIROSHI TANIUCHI (177), Laboratory of Chemical Biology, National Institute of Arthritis and Metabolic Diseases, National Institutes of Health, Bethesda, Maryland TSUNEKO UCHIDA (205), Department of Biophysics and Biochemistry, Faculty of Science, The University of Toyko, Hongo, Tokyo, Japan lRWIN B. WILSON (373), Department of Chemistry, University of Colorado, Boulder, Colorado SIMON C. K. WONG (499), Department of Biological Chemistry, Harvard Medical School, Boston, Massachusetts JOHN C. WRISTON, J R . (101), Department of Chemistry, University of Delaware, Newark, Delaware HAROLD W. WYCKOFF (647), Department of Molecular Biophysics and Biochemistry, Yale University, New Haven, Connecticut MASANOBU YAMAMOTO (337, 355), Fisheries Research Board of Canada, Vancouver Laboratory, Vancouver, Canada. C. L. ZIELKE (47), Department of Biochemistry, Michigan State University, East Lansing, Michigan

This Page Intentionally Left Blank

Preface This volume, the fourth of the treatise, continues the coverage of enzymes catalyzing hydrolyses. More is known about the hydrolases than any other group of enzymes. They thus command a major space allotment. Volume I11 was devoted entirely to enzymes catalyzing hydrolysis of peptide bonds. This volume deals principally with enzymes catalyzing hydrolysis of phosphate ester bonds. Also included is a shorter portion on the hydrolysis of other C-N bonds. Coverage of hydrolysis will be completed with Volume V, which includes cleavage of sulfate and carboxylate esters and both hydrolysis and phosphorolysis of glycosidic bonds. I n these volumes an attempt is made to present an authoritative compilation for those enzymes about which considerable information is now available. Hence, a number of lesser studied enzymes are not included, although often information is given about them in chapters covering their better studied relatives. Authors have responded well to the guideline limiting coverage to molecular aspects of the enzyme or of the reaction catalyzed. As a result some chapters are short. In some instances, however, the amount of information is striking. For two of the phosphodiesterases, pancreatic ribonuclease and staphylococcal nuclease, this volume includes the latest structural information from X-ray analysis. Pancreatic ribonuclease is indeed one of the best understood of all enzymes; information contained in the chapter dealing with this enzyme has broad implications. With this volume we are approaching midstream in planning the treatise. Occasionally I have pondered my wisdom, or lack thereof, in assuming the editorial responsibility. But the completion of each volume is a gratifying experience, and the quality of the product an addicting reward. For planning this volume, as well as Volume 111, thanks are due to xv

xvi

PREFACE

the Advisory Board members C. B. Anfinsen, I. R. Lehman, Alton Meister, and Stanford Moore. For the quality of the volume, much appreciation is extended to the staff of Academic Press, as well as to Lyda Boyer for her editorial assistance. Their interest, enthusiasm, and encouragement were invaluable assets. PAULD. BOYER

Contents of Other Volumes Volume I: Structure and Control

X-Ray Crystallography and Enzyme Structure David Eisenberg Chemical Modification by Active-Site-Directed Reagents Elliott Shuw Chemical Modification as a Probe of Structure and Function Louis A . Cohen Multienzyme Complexes Lester J . Reed and David 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, Jr. Mechanisms of Enzyme Regulation in Metabolism E . R . Stadtman Enzymes as Control Elements in Metabolic Regulation Daniel E . Atkinson Author Index-Subject

Index xvii

xviii

CONTENTS OF OTHER VOLUMES

Volume II: Kinetics and Mechanism

Steady State Kinetics W . W . Cleland Rapid Reactions and Transient States Gordon B. Hammes and Paul R. Schimmel Stereospecificity of Enzymic Reactions G . PopjcEk Proximity Effects and Enzyme Catalysis Thomas C . Bruice Enzymology of Proton Abstraction and Transfer Reactions Irwin A . Rose Kinetic Isotope Effects in Enzymic Reactions J . H . Richards Schiff Base Intermediates in Enzyme Catalysis Esmond E. Snell and Samuel J . Di Mari Some Physical Probes of Enzyme Structure in Solution Serge N . Timasheff Metals in Enzyme Catalysis Albert S. Mildvan Author Index-Subject Index

Volume 111: Hydrolysis: Peptide Bonds

Carboxypeptidase A Jean A. Hartsuclc and William AT. Lipscomb Carboxypeptidase B J . E . Folk Leucine Aminopeptidase and Other N-Terminal Exopeptidases Robert J . DeLange and Emil L. Smith Pepsin Joseph S. Fruton


Chymotrypsinogen : X-Ray Structure J . Kraut The Structure of Chymotrypsin D . M . Blow Chymotrypsin-Chemical George P . Hess

Properties and Catalysis

Trypsin B . Keil Thrombin and Prothrombin Staffan Magnusson Pancreatic Elastase B. S. Hartley and D . iM.Shotton Protein Proteinase Inhibitors-Molecular Aspects Michael Laskozcski, Jr. and Robert W . Sealock Cathepsins and Kinin-Forming and -Destroying Enzymes Lowell M . Greenbaum Papain, X-Ray Structure J . Drenth, J . iV. 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. Marleland, Jr. and Emil L. Smith Streptococcal Proteinase Teh-Yung Liu and S. D . Elliott The Collagenases Sam Seifter and Elvin Harper Clostripain William M . iMitchel1 and William F . Harrington

XiX

xx


Other Bacterial, Mold, and Yeast Proteases Hiroshi Matsubara and Joseph Feder Author Index-Subject

Index

Volume V (Tentative): Hydrolysis (Sulfate Esters, Carbowyl Esters, Glycoside Bonds) , Phosphorolysis, and Hydration

Plant and Animal Amylases John A . Thoma, Joseph E . Spradlin, and Stephen Dygert Neuraminidases Alfred Gottschalk and A . S . Bhargava Cellulases

D. R . Whitaker The Hydrolysis of Sulfate Esters A . B. Roy Arylsulfatases R. G. Nicholls and A . B. R o y Phage Lysozymes and Other Lytic Enzymes Akira Tsugita Bacterial and Mold Amylases Toshio Takagi, Hiroko Toda, and Toshizo Isemura Carboxylic Ester Hydrolases Klaus Krisch Hyaluronidases Karl Meyer Phospholipases Donald J . Hanahan Glycogen and Starch Debranching Enzymes E . Y . C . Lee and W . J. Whelan Yeast and Neurospora Invertases J . Oliver Lampen


xxi

L-Glucan Phosphorylases-Chemical and Physical Basis of Catalysis and Regulation Donald J . Graves and Jerry H . Wang Acetylcholinesterase Harry C . Froede and Irwin B. Wilson Dehydrations Requiring Vitamin H,z-Coenzyme Robert H . Abeles Dehydration in Nucleotide Linked Deoxysugar Synthesis L. Glaser and H . Zarkowsky Aconitase Jenny Pickworth Glusker P-Hydroxydecanoyl Thioester Dehydrase Konrad Bloch Purine Nucleoside Phosphorylase R . E . Parks, Jr. and R . P . Agarwal Enolase Finn Wold Fumarase and Crotonase Robert I,. 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


The Enzymes VOLUME IV

H Y D R O L YSlS Other C-N Bonds Phosphate Esters Third Edition


Ureases F. J . REITHEL I . Introduction . . . . . . . . I1. Isolation and Purification of Jack Bean Urease Enzymic Activity Measurement . . . I11. Molecular Properties . . . . . . A . Molecular Weight Determinations . B . Other Physical Properties . . . C . Chemical Composition and Behavior . D . Urease Derivatives . . . . . E . Immunological Behavior . . . . IV . Ureases from Other Sources . . . . V . Catalytic Properties . . . . . . . A . Mechanism . . . . . . . B . Substrate Specificity . . . . . C . Kinetic Studies . . . . . . D. Active Site Studies . . . . . VI . Summary . . . . . . . . .

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1 2 4 5 8 10 11 12 13 13 15

15 16 18 20 21

.

1 Introduction

The title retains the trivial name for enzymes with the systematic name of urea amidohydrolase and the Enzyme Commission code number of EC 3.5.1.5. Ureases are hydrolases acting on C.-N bonds (nonpeptide) in linear amides and thus belong to a group that includes glutaminase, formamidase, and formyltetrahydrofolate deformylase . The title is plural to emphasize that urease activity may be exhibited by several protein species. Urease, singular. has come to mean by common usage. that particular enzymic protein first crystallized by Sumner from jack bean 1

2

F. J. REITHEL

meal. An excellent account of the earlier work was provided by Sumner in the first edition of this compendium (I), and in a well-known book ( 2 ); a more recent review was assembled by Varner (3). Although the present review recognizes the multiplicity of ureases, the bulk of our knowledge derives from the crystalline protein isolated from the jack bean Canavalia ensiformis. This seed has little utility for other purposes, and it is not known what function is served by urease in the economy of the plant. Since both urea and urease occur widely in the plant kingdom there have been numerous studies of variations in the concentrations of both components ( 4 ) .There seems to be no obvious correlation. A recent ontogenetic study on the jack bean ( 5 ) has not revealed conclusive evidence of an important function for urease. To date no one has demonstrated urease to be an enzyme to engage the enthusiasm of the investigator on the basis of its biological function. As a hydrolase it has been notable for specificity. This, together with an outstanding catalytic efficiency, has justified a search for the details of the hydrolytic mechanism and has potentiated numerous investigations of urease activity in a wide spectrum of biological material. I n the recent past urease has stimulated interest as a model in the study of subunit interactions. The preparative molecular weight of 489,000 ( 6 ) appears to be the result of a systematic arrangement of single polypeptide chain subunits of 30,000 daltons ( 7 ) .

II. Isolation and Purification of Jack Bean Urease

I n view of the isolation of the crystalline enzyme from this source (chosen because of the high concentration) 45 years ago, it might be supposed that a discussion of preparative procedures would be unnecessary. On the contrary, vigorous contentions concerning the purity and activity of crystalline urease have been a hallmark of its literature right to the present. Several factors contribute to this situation. The urease content of jack bean meal varies according to the origin and 1. J. B. Sumner, “The Enzymes,” 1st ed., Vol. 1, Part 2, p. 886, 1951. 2. J. B. Sumner and G. F. Somers, “Chemistry and Methods of Enzymes,” 3rd rev. ed. Academic Press, New York, 1953. 3. J. E . Varner, “The Enzymes,” 2nd ed., Vol. 4, p. 247, 1959. 4. E. G.Bollard, S y m p . SOC.E z p . Bid. 13, 304 (1959). 5. P.P. Sehgal and A. W. Naylor, Botan. Gaz. 127, 27 (1966). 6. F.J. Reithel and J. E. Robbins, ABB 120, 158 (1967). 7. C. C. Contaxis, Ph.D. Thesis, University of Oregon, 1970.

1. URENSES

3

age of the seed and the method used in preparing the meal ( 8 , 9 ) . The acetone extraction must be performed precisely as directed or some other extraction procedure adopted. The enzymic activities of various preparations may be difficult to compare because assay conditions have not been uniform and because the activity is sensitive to minute concentrations of heavy metals. The propensity of the enzyme to dissociate, and to form oligomers, compounds the difficulty of assigning a specific activity. Finally, there are anomalies (10) noted during assay that suggest variations in activity with conformational change and this further adumbrates specific activity as a criterion. As Mamiya and Gorin (11) note, Sumner occasionally had difficulty in preparing crystalline urease from some jack bean meals and this is a technical problem that has received attention in each recent improved method of preparation (11-15). Modification of Sumner’s extraction procedure by including p-mercaptoethanol (11, 14) t o diminish aggregation and ethylenediaminetetraacetate (EDTA) (15,15) to maintain a low concentration of metal ions has been helpful. The procedure used in this laboratory with consistent success over a period of 4 years, and involving several students, is essentially that of Mamiya and Gorin (11) but with EDTA included in the buffers. [There is an error in Mamiya and Gorin (11). The concentration of acetone used was 32%, not 360/0.] A method for the isolation of urease from hydrated jack beans developed by Sehgal and Naylor (15) employs DEAE-cellulose as a final purification step. A method intended for purification of urease from mammalian sources (16) was developed using jack bean meal as a standard source. It employed no acetone but is a combination of (NH,) &30, precipitation and Sephadex column separations. [Note Fish8. The prime supplier of jack beans for many years was Mr. Ernest Nelson, Waldron, Arkansas. His business is being continued by his son, Winton R. Nelson. Companies offering jack beans and jack bean meal at. the present time include Sigma Chemical Co., St. Louis, Mo.; General Biochemicals Co., Chagrin Falls, Ohio; and Worthington Biochemical Co., Freehold, N. J. 9. W. N . Fishbein, Ann. N . Y . Acad. Sci. 147, 857 (1969). 10. Workers in several laboratories have noted that the activity of urease uppears to Increase upon standing a t room temperature. Until this is understood assay prowdures cannot be assumed to yield precise values. 11. G. Mamiya and G. Gorin, BBA 105, 382 (1965). 12. K. K. Lynn, BBA 146, 205 (1967). 13. R. I,. B1:ikeley. E . C. Webb, and 13. Zerner, Biochemishy 8, 1984 (1969). 14. F. J. Itpithel, J. E. Robbins, and G . Gorin, ABB 108, 409 (1964). 15. P. P. 8ehg:il a d A . W. Naylor, Plant Physiol. 41, 567 (1966). 16. E. J. Conway, 0. Fitzgerald, and I10.5 when urease solutions were irradiated with 280 or 296 nm radiation. Addition of urea increased the fluorescence 49. I. Matsuo and G. Mamiya, Keio J . M e d . 17, 189 (1968). 50. J. B. Sumner and D. B. Hand, JACS 31, 1255 (1929).

51. E. A. Chernitskii, V. M. Mazhul, and S. V. Konev, Biofizika 13, 581 (1968).

11

.. UREASES

md addition of guanidine-HC1 abolished the maxima. Fishbein et al. :9,39,56)interpreted changes in mobility in gel electrophoresis, followng repeated recrystallizations, as the formation of conformer isoenzymes. The ultrasonic absorption coefficient of urease has been measured (53) md found to increase in the presence of urea.

>. CHEMICALCOMPOSITIONAND

BEHAVIOR

The amino acid composition (6,54) of urease has no unique features but does provide a guide for molecular weight considerations. The two itations are not in agreement and hence further work is in order. Of )articular interest are the values of 82 (6,43) and 64 (54) half-cystines ier 485,000 MW. This is unexpectedly less than the value of 29 cystine dus 47 cysteine per 473,000 reported by Gorin et al. ( 5 5 ) . However, uch values necessarily reflect the technique used to estimate the protein oncentration. In view of the wide discrepancies in absorbance one night expect a corresponding scatter in sulfhydryl content. In a later nvestigation Gorin and Chin (56) found, titrating with N-ethylmalimide, 21 sulfhydryl groups that reacted rapidly with no loss of enymic activity, 7-8 groups that reacted more slowly with the loss of 0% of the activity, and 16-20 groups that reacted slowly. Andrews (43) ound that estimation either with N-ethylmaleimide or 5,5'-dithiobis2-nitrobenzoic acid) indicated the following numbers of sulfhydryl roups per (16n) : 26-28 rapidly react without loss of activity, 7-9 titrate :ss rapidly with loss of activity, and 45-50 more become available fol)wing unfolding. Twenty years ago. Desnuelle and Rovery (57) prolosed that the loss of activity during such titrations was not a result of 'locking groups and modifying their chemical behavior but rather that he loss of activity reflected a change in structure. The use of "S$beled tetraethylthiuram disulfide (58) has indicated 19 readily titratble sulfhydryl groups per mole of protein and a total of 53 groups itratable when the protein was dissolved in 0.25% SDS. The protein sed was a commercial sample of indeterminate age and submaximal ctivity. There is no compelling evidence for the existence of interchain 52. W.N.Fishbein, C. L. Spears, and W. Scurzi, Federation Proc. 28, 468 (1969). 53. M.Pancholy and T. K. Saksena, J . Acoust. Soc. Am. 44, 639 (1968). 54. J. M.Milton and I. E. P. Taylor, BJ 113, 678 (1969). 55. G. Gorin, E. Fuchs, L. G. Butler, S. I,. Chopra, and R. T. Hersh, Biochemistry , 911 (1962). 56. G. Gorin and C. C. Chin, BBA 99, 418 (1965). 57. P. Desnuelle and M. Rovery, BBA 3, 26 (1949). 58. A. H.Neims, D. S.Coffey, and L. Hellerman, JBC 241, 3036 (1966).

12

F. J . REITHEL

disulfide bonds in the (16n) species, and there is no evidence for intrachain disulfide bonds. Without doubt disulfide bonds do form in (32n) and (4%) forms and similar oligomers, but there is no evidence that such formation is mandatory. When 40% of the groups are blocked in unfolded preparations reassociation (43) is strongly inhibited. I n a recent paper ( 2 5 ) , Bailey and Boulter have presented evidence for a single N-terminal rnethionine residue per (a suggested subunit of) 75,000 daltons. A single C-terminal sequence, -Tyr-Leu-Phe, was found using carboxypeptidases A and B and hydrazinolysis. Asparagine has also been reported as the N-terminal amino acid (59).

D. UREASE DERIVATIVES As noted above, Sumncr and co-workers were unable t o determinc the diffusion coefficient of urease unless thcy added Na2S0, and NaHSO, to the phosphate buffer (40) used. Nichol and Creeth, employing identical concentrations (GO),measured both the sedimentation coefficient and the electrophoretic mobility of sulfite-modified urease. They concluded that sulfite contributed to the formation of -S-SO,- groups attached to the (16n) species. Some of these groups they ascribed to the scission of intermolecular disulfide bonds of aggregated forms; others, they suggested, arose from the 22 reactive sulfhydryl groups that react with 0, (air) to form transitory disulfides that can, in turn, react with sulfite. An unusual type of derivative is the complex that forms between urease and bentonite in acid medium (61). The adsorbed form was found catalytically active. Similarly, urease immobilized in a polyacrylamide gel matrix has been used to prepare a urea-specific enzyme electrode (62). Yet another active water-insoluble derivative has been prepared (63) by allowing p-chloromercuribenzoate-treated urease to react with a diazotized copolymer of p-amino-D,L-Phe and L-Leu. Urease has been found to retain about 20% of its original activity when encapsulated in 100 p microcapsules of benzalkonium-heparin in collodion (64). Both carboxymethyl and aminoethyl urease have been prepared (251, 59. R. L. Blakeley, J. A . Hinds, H . E. Kunze, E. C. Webb. and B. Zerner. Biochemitry 8, 1991 (1969). 60. I,. W. Nichol and J. M. Creeth. BBA 71, 509 (1963). 61. G. Surand, Ann. Znst. Pasteur Suppl. 3. 121 (1965). 62. G . G. Guilbault and J. G. Montalvo, Jr., JACS 91, 2164 (1969). 63. E. Ricsel and E. Katchalski, JBC 239, 1521 (1964). 61. T. M. S. Chang, L. J. Johnson, and 0. J . Ransome, Can. J. Physiol. Plinrmacol. 45, 705 (1967).

1.

UREASES

13

the latter being used to prepare tryptic peptide “maps” of urease. Reduced carboxymethylated urease was reported (65) to have a molecular weight of 16,000 on the basis of tryptophan (Trp) content.

E. IMMUNOLOGICAL BEHAVIOR

It was noted in previous reviews (1,s) that injection of urease into rabbits elicited an antibody and that the precipitate formed between urease and its antibody still possessed catalytic activity. This indicates that the antigenic and catalytic regions are not identical. A more recent study employing horse and rat antisera (66)revealed only one major antigenic component in urease preparations and confirmed the lack of complete enzyme inhibition by the precipitin reaction. Urease-immunized laboratory animals have been observed (67) to suffer a depression in the in vivo hydrolysis of 14C urea. This has received close scrutiny since there is evidence (68) that passively immunized animals (injected with anti-urease) show increased survival after whole-body y radiation. IV. Ureases from Other Sources

Ureases occur in a large number of microorganisms. A listing has appeared in a paper (69) calling attention t o the existence of urease even in organisms that do not hydrolyze urea in the culture medium. Particularly designed for the assay of urease in bacterial extracts is a coupled enzyme assay ( 7 0 ) . This procedure is based on the high K , value of glutamic acid dehydrogenase for ammonia. Thus, in the presence of a-ketoglutarate, the oxidation of NADH was found to be proportional to the ammonia resulting from urease action. It has been noted that the various strains of Enterobacteriaceae contain either urease or P-galactosidase (71). The urease of the genus Proteus has continued to receive attention. 65. K. Sekita and G. Mamiya, Proc. 7 t h Intern. Congr. Biochem., T o k y o , 1967 Abstracts IV, p. 761. Sci. Council Japan, Tokyo, 1968. 66. W. J. Visek, M. E. Iwert, N. S. Nelson, and J. H. Rust, A B B 122,95 (1967). 67. H. C. Dang and W. J. Visek, Proc. SOC.Ezpptl. B i d . M e d . 105, 164 (1960). 68. W. J. Visek and H. C. Dang, Ezcerpta M e d . Monogr. Nucl. Med. B i d . No. 1, 292 (1966). 69. H. Seneca, P. Peer, and R. Nally, Nature 193, 1106 (1962). 70. H. Kaltwasser and H. G. Schlegel, Anal. Biochem. 16, 132 (1966). 71. M. Catsaras, Ann. Znst. Pasteur Lille 16, 175 (1965).

14

F. J. REITHEL

A partially purified enzyme from P . mirabilis (72) was found to have a molecular weight of 151,000. The urease of P . rettgeri is an inducible enzyme that appears only when urea, but not its analogs, are present in the media ( 7 3 ) .Proteus vulgaris urease was found to be inhibited in vitro by thiourea and two derivatives ( 7 4 ) ,and by hydroxamic acids (93). I n Azotobacter vinelandii, urease appears to be synthesized only when urea or thiourea is present ( 7 5 ) . A study of the urease constitutive in Corynebacterium renale (76) did not reveal features remarkably different from the plant enzyme. A similar conclusion was reached in the characterization of a highly purified enzyme from B . pasteurii ( 7 7 ) . Stewart (78) has devised a medium for the detection of urease activity in pseudomonads and has resolved uncertainties that have developed in the literature. It has been reported that Sarcina ureae produces urease as an exoenzyme (79). Bacteria-free cultures of blue-green algae yielded extracts in which urease could be demonstrated (80,81). Again, in algae as in bacteria (69) the function of urease is conjectural (82). The addition of urea to tissue cultures of human amnion cells has been found to induce urease formation (83).Urease from the seed of the legume Glyciridia maculata has been purified (84) and found to be advantageous for commercial purposes. This enzyme was reported to have no activity a t 30" and below but to be active at 50"-60"! Ureases from several sources have been examined for enzymically active low molecular weight forms ( 4 7 ) .It was noted that the 12 S forms from jack bean did not hybridize with that from B . pasteurii. It now seems probable that gastric urease is bacterial in origin (85). The OC72. J. A. Andersen, F. Kopko, A. J. Siedler, and E. G. Nohle, Federation Proc. 28, 764 (1969). 73. I. Magana-Plaza and J. Ruiz-Herrera, J . Bacteriol. 93, 1294 (1967). 74. R. S. Pianotti, R. R. Mohan, and B. S. Schwartz, Proc. SOC.Exptl. Biol. Med. 122, 506 (1966). 75. S. I,. Mehta, M. S. Naik, and N. B. Das, Indian J . Biochem. 4, 194 (1967). 76. A. J. Lister, J . Gen. Microbiol. 14, 478 (1956). 77. A. D. Larson and R. E. Kallio, J . Bacteriol. 88, 67 (1954). 78. D. J. Stewart, J . Gen. Microbwl. 41, 169 (1965). 79. A. S. Pel'ttser, Izv. Timiryazev. Selskokhoz. Akad. No. 3, p. 230 (1969). 80. D. S. Berns, P. Holohan, and E. Scott, Science 152, 1077 (1966). 81. D. S. Berns, E. Scott, K. T. O'Reilly, and P. D. Holohan, N . Y . Stale Depl. Health, Ann. Rept. Div. Lab. Res. p. 82 (1964). 82. It should be noted that urease-negative extracts of Candida utilis and Chlorella catalyze an ATPdependent cleavage of urea to CO, and NH3. See R. J. Roon and B. Levenberg, JBC 243, 5213 (1968). 83. F. Lieben and K. Springer, Enzymologia 27, 47 (1964). 84. K. Valmikinathan, V. N. V. Rao, and N. Verghese, Enzymologin 34, 257 (1968). 85. A. M. Delluva, K . Markley, and R. E. Davies, BBA 151, 646 (1968).

1. UREASES

15

currence of urease in the gastric mucosa of man and several animals has been observed repeatedly for 40 years. Attempts to use antibiotics such as oxytetracycline (867,chloramphenicol, penicillin, and neomycin led to results of poor reproducibility. However, it is rather clear that gastric tissue from germfree animals (85) contains no urease. Urease activity in soils has been found to reflect the bacterial count and content of organic matter. The urease isolated from an Australian forest soil (87) was crystallized and found to have a specific activity of 75 Sumner units (S.U.) per mg. The molecular weight species were estimated (sedimentation velocity) to be 42, 131, and 217 x lo3. T h a t urease activity persists in soils is shown by the finding that enzymic activities, including urease, could be demonstrated in soil samples over 8000 years old (88).

V. Catalytic Properties

The Enzyme Commission catalog (EC 3.5.1.5) lists the urease reac2 NH,. Since two C-N bonds are broken tion as urea 2 HzO = CO, it is evident that the stoichiometric relation above is the result of two component reactions. Any conjecture concerning the mechanisms of these reactions and the nature of the intermediates must encompass the action of inhibitors and the spectrum of substrates. Some of the organic inhibitors that have been reported are shown in Table I. The substrates that have been shown to be hydrolyzed are listed in Table 11.

+

+

A. MECHANISM The work of Gorin (89) and of Blakeley et al. (59) has provided convincing evidence that carbamate is the intermediate in a two-step reaction.

o=c,,OH

NH,

-H,CO,

+

,ONH O=C\

NH,

+

NH,

]

:

2 NH,

86. M. E. Belding and F. Kern, Jr., J. Lab. C h a . M e d . 61, 560 (1963). 87. M. H. Briggs and L. Segal, Life Sci. 1, 69 (1963). 88. J. J. Skujins and A . D. McLaren, Enzymologia 34, 213 (1968). 89. G. Gorin, BBA 34, 268 (1959).

4-

HOH

16

F. J . REITHEL

Presumably urease forms a carbamoyl complex HtN-C-enz

II

0 as one of the ES complexes and presumably water is the acceptor in a carbamoyl transfer reaction. Carbamate thus becomes the obligatory substrate for thc second step. Since the evidence derives from kinetic data there is, as yet, no direct evidence for the mechanism presented. However, Sumner had long since shown that ammonium carbonate was formed. The present explanation accounts for thc observations and rules out CO, as an initial product. There are two reports that hydroxamic acid inhibition is reversible (59,90)and one that inhibition is irreversible (91). The inhibition appears to be competitive. The rather extensive screening of 36 hydroxamic acids was accomplished with sword bean urease (N),but Proteus urease (92) and jack bean urease (59) also have been found to be inhibited by these specific inhibitors. Using tritium-labeled caprylohydroxamic acid and sword bean urease, Kobashi et al. (94) have shown the formation of an inactive complex containing two moles of inhibitor per mole of enzyme. B. SUBSTRATE SPECIFICITY Hydroxyurea is a substrate ( 9 5 ) , but the rate a t which it is hydrolyzed decreases during the progress of the reaction. The concomitant inhibition disappears as the hydroxyurea is depleted. There is no satisfactory explanation of the events described, and there has been no adequate description of the active site. It has been demonstrated (96) that hydroxyurea added to urease solutions inhibited the hydrolysis of urea and that the extent of inhibition depcnded both on the order of addition and the time of exposure of the enzyme to the inhibitor. Dihydroxyurea is a substrate at high concentration (97) ; K , = 1.25 X 1k2.The products are hydroxylamine and CO,. Like hydroxyurea, hydroxylamine and (in some cases) hydroxamic acids, dihydroxyurea is a 90. 91. 92. 93. 94. 95. 96. 97.

K. Kobashi, J. Hase, and K. Uehara, BBA 65, 380 (1962). W. N. Fishbein and P. P. Carbonc, JBC 240, 2407 (1965). G. R. Gale, J . Bacten’ol. 91, 499 (1966). J. Hnse and K. Kobashi, J . Biochem. ( T o k y o ) 62, 293 (1967). K. Kobaslii, J. Haw, and T. Komai, BBRC 23, 34 (1966). W. N. Fishbein, JBC 240, 2402 (1965). G. R. Galc, Biochem. Phnrmacol. 14, 693 (1965). W. N. Fishbein, JBC 244, 1188 (1969).

1.

17

UREASES

noncompetitive inhibitor. Whether ureasc binds these compounds at, or near, the catalytic site for urea is not known. Unlike hydroxyurea, the extent of inhibition by dihydroxyurea does not increase with time after TABLE I ORGANIC INHIBITORS OF UI{EASI: Compound Hydroxamic acids

Formula

lief.

13-C-NHOH

C hlorttmphenicol

Pheny lurea

I,-Phe Chlormerodrin (neohydrin)

CI-Hg-CH-CH-CH2--S

Hy droxyurea

HgN-C-NHOH

Hydroxylamine Dihydroxyurea

0 HzN-OH HOHN-C-NHOH

OCHs I

H-C-N

K

Hz

I1

Thiourea

Oxytet racycline

Dimethyl sulfoxide

98. E. Mueller, Nnturwissenschaften 54, 226 (1967). 99. E:. Gerhards and H. Gibian, Ann. 'V. Y . Acntl. Sci. 141, 65 (1967). 100. K. Kleczkowski and M. Dabrowska, Bull. Acad. Polon. Sci., Ser. Sci. Biol. 16, 267 (1968).

18

F. J. REITHEL

TABLE I1 UREASE SCJBSTR.4TES OTHER

Compound

Forniiila

THANUREA lief.

Hydroxyurea

HZN-C--N

HOH

(95)

Dihydroxyiirea

0 HOHN-C-NHOH

(97)

II

II

0

an equilibrating preincubation. The inhibition is kinetically reversible and less than that of hydroxyurea. The relation between catalytic activity and pH as measured in 0.1 M tris-maleate buffers is substantially the same for each of the thrce substrates (97) when percent of maximal activity is plotted as the ordinate. C. KINETICSTUDIES The earlier documentation of iirease kinetics by Kistiakowaky and co-workers has been extended more recently by Lynn ant1 Yankwich (101,102). Both overall rcaction rates and 13C kinetic istope effects were determined under conditions where the following variables wero altered: temperature, pH, extent of rcaction, ionic strciigth, conceiitration of reactants, type of buffer, arid age of preparation. I t was concluded that each preparatioii of urcasc was unique, respon(letl uniquely to the medium, and changed throughout the course of tlic reaction. In view of the handling of tlic enzyme, its exposure to glycerol, and grcat dilution and aging changes, the variations found in kinetic data also not surprising. Table I11 gives representative literature values of I>NH, >F >OH>>Cl which suggests that electron withdrawing groups decrease the reactivity. Perhaps altered positioning of the various substrate analogs in the catalytic site accounts for the most significant effect on the altered rates of deamination in these cases. In any event the data are consistent with a subtle balance between the bond-breaking and bond-forming steps (118) in the development of the transition state. The isolation of the proposed covalently linked enzyme intermediate would aid materially in the elucidation of this mechanism.

c. CONSIDERATIONS O F PHYSIOLOGICAL

FUNCTION

The suggestion of Berne (118) that adenosine may play an important part in regulating coronary blood flow has provided considerable inieptus to the study of adenosine deaminases. The auriculoventricular block of mammalian heart caused by adenosine is transient, probably because of rapid deamination to the inactive inosine (119). 2-Chloroadenosine1 which has long-lasting effects and 10 times the potency of adenosine in causing heart block in the guinea pig (119), was not deaminated by the ox heart enzyme (86). A number of 2-substituted adenosine analogs with a free 6-amino group which inhibit ox heart adenosine aminohydrolase have also been found to cause prolonged vasodilation in a number of mammalian species ; however, comparison of the relative potencies of these analogs with their inhibitor constants shows no correlation between the inhibitory effects of the analogs on ox heart aden118. R. N . Berne, Physiol. Rev. 44, 1 (1964). 119. R. H. Thorp and L. B. Cobbin, Arch. Interrt. Pharmacodyn. 118, 95 (1959).

64

C. L. ZIJCLKE AND

C. H.

SUELTER

osine aminohydrolase and their effect as vasodilatory agents (86). The inhibition constant for ouabain also shows no correlation with biological effects (86). Thus the role of adenosine in the regulation of coronary blood flow requires additional study. I n addition to the general catabolic role of adenosine aminohydrolase in purine metabolism first suggested by Brady (120) and in the detoxification of the pharmacologically active adenosine, other results (121) imply a role in cellular deoxyribonucleic acid (DNA) metabolism. I n the liver where catabolic enzymes are mostly known to increase with age of the animal, the level of adenosine aminohydrolase does not show any variation. On the other hand, in tissues showing high DNA synthesis and intense mitotic activity, as observed during normal tissue regeneration and normal or neoplastic cell proliferation, the levels of catabolic enzymes are low but adenosine aminohydrolase is surprisingly high.

IV. 5'-Adenylic Acid Aminohydrolase

A. MOLECULAR CHARACTERISTICS 1 . Purification and Homogeneity

Several preparations for 5'-AMP aminohydrolase (EC 3.5.4.6) have been described which yield homogeneous enzyme (68, 122, 123). Of these the method utilizing the direct absorption of AMP aminohydrolase from a crude rabbit muscle extract with cellulose phosphate and its consequent elution provides a one-step purification of ultracentrifugal and electrophoretically homogeneous enzyme in high yields (68). This method has been adapted with slight modifications for preparations from rat ( l 2 4 ) , chicken breast (123), and carp muscle (125).The scheme for purification of enzyme from elasmobranch fish (Raia clavata) involved calcium phosphate gel and DEAE-cellulose chromatography (126). While the enzyme from chicken breast muscle was homogeneous by 120. T . Brady, BJ 36, 478 (1942). 121. M. Galy-Fajou. C . E. Stripati, and Y. Khouvine, Bull. Soc. Claim. Biol. 51, 52 (1969). 122. Y.-P. Lee, JBC 227, 987 (1957). 123. H. Henry and 0. P. Chilson. Co'mp. Biochem. Physiol. 29, 301 (1969). 124. A. W. Murray and M. R. Atkinson, Biochemitry 7 , 4023 (1968). 125. J. Purzycka-Preis and M. Zydowo, Actn Biochim. Polon. 16, 235 (1969). 126. W. Makarewicz. C o m p . Biochem. Physiol. 29, 1 (1969).

3.

PURINE AMINOHYDROLASES

65

clectrophorctic, ultr~iccutrifugal,itiitl im~iii~noclcctrophoreticcriteria, 110 data were given for the rat and fish muscle preparations. A previous preparation of rat and rabbit musclc enzyme (127) and enzyme purified from rat liver (128) was not homogeneous. No criteria were published for the 200-fold purified calf brain cnzymc (129). 2. Chemical and Physical Properties Whereas the rabbit muscle (68)and brain preparations (129) required 1-10mM mercaptoethanol and KCl or LiCl for stability over extended time periods, the preparation described by Lee was not affected by reducing or oxidizing agents (130). Multivalent anions, such as tripolyphosphate, 3-iso-AMP, ATP, and GTP, but not substrate, stabilized the calf brain enzyme against heat inactivation (129, 131). The molecular weight of 320,000 obtained for the muscle enzyme from sedimentation-diffusion data a t 2-6 mg/ml and 8 = 0.75 (132) is to be compared with 270,000 obtained by Wolfenden et al. from s ~ =~11.1, S ~ and D,,,, = 3.75 X lo-‘ cm2 sec-I, and V = 0.731 calculated from the amino acid content (92). The rabbit muscle enzyme has a normal amino acid content, that is, no unusually low or large amount of a particular amino acid was found. Of the 32 cysteine/half-cystine residues per mole based on a molecular weight of 270,000, 6.2 were rapidly titrated with p-mercuribenzoate (92). Typical protein absorption spectra were reported for elasmobranch fish (126), carp (125), rat (127), and rabbit muscle enzyme (68). An Ettm a t 280 nm = 9.13 has been reported for the rabbit muscle enzyme (133). The atypical absorption spectrum with a maximum a t 275-276 nm observed by Lee (132) is indicative of contaminating bound nucleotides. Inhibition of the chicken breast enzyme by rabbit antisera for chicken breast enzyme; the lack of effect on the chicken brain, heart, or erythrocyte enzyme ; and the differences in substrate specificity exhibited by the brain and breast muscle enzyme are consistent with at least two isozymes of chicken 5’-AMP aminohydrolase (123). Isozymic patterns, while perhaps implied by differences in certain kinetic pa127. R. D. Curric and H. L. Webster, BBA 64, 30 (1962) ; G. Nikiforuk and S. P. Colowick, “Methods in Enzymology,” Vol. 2, p. 469, 1955. 128. L. D. Smith and D. E . Kiaer, BBA 191, 415 (1969). 129. B. Setlow and J. M . Lowenstein, JBC 242, 607 (1967). 130. Y.-P. Lee, JBC 227, 999 (1957). 131. B. Setlow and J. M. Lowenstein, JBC 243, 3409 (1968). 132. Y.-P. Lee, JBC 227, 993 (1957). 133. C. L. Zielke and C. H. Suelter, Federation Proc. 28, 728 (1969).

66

C. L. ZIELKE AND C. H. SUELTER

rameters of the calf brain (129) and rabbit muscle enzyme (68),have not been clearly delineated for other preparations.

B. CATALYTIC PROPERTIES 1. Specificity I n general, AMP aminohydrolase specificities have not been thoroughly defined perhaps because of difficulties until recently in obtaining pure enzyme. I n addition to AMP and dAMP, the muscle enzyme catalyzes the deamination of N6-methyl AMP, N”-ethyl AMP, formycin-5’-monophosphate, adenosine-5’-monosulfate, adenosine-!Y-phosphoramidate, adenosine, ADP (133), adenosine-5’-phosphorothioate, and 6-chloropurine 5’-ribonucleotide (124) ; ATP, GMP, CMP, 2’-AMP, 3’-AMP, 3’, 5’-cyclic AMP, 3-iso-AMP, N1-methyl AMP, toyocamycin5’-monophosphate, tubercidin-5’-monophosphate, and 6-mercaptopurine5’-ribonucleotide are not deaminated (133). The elasmobranch fish muscle, carp muscle, and avian brain enzymes appear to be specific for AMP and dAMP (123, 125, 126). Extracts from pea seed and erythrocytes and the purified calf brain enzyme are specific for AMP (48,131, 134). 2. Kine tics

The kinetic parameters of various muscle AMP aminohydrolases presented in Table V (51, 68,12.2, 124-127) are similar except for the lower specific activities exhibited by the fish enzymes for which no criteria of homogeneity are presently available. Specific activities reported for brain enzymes not shown in Table V are 1 5 pmoles/min/mg for calf (129) and 30 pmoles/min/mg for chicken (123).Although the pH optimum for AMP deamination varies depending upon the source, it normally occurs in a range from pH 5.9 to 7.1 (48, 125, 126, 129, 150, 135-137).

a. Activation. Most preparations of AMP aminohydrolase are activated by monovalent cations and nucleoside di- or triphosphates (Table VI) (48,54,68,123, 126, 126, 128, 129,131,137-148). Potassium is gen134. A . Askuri, Science 141, 44 (1963). 135. G. Nikiforuk and S. P. Colowick, JBC 219, 119 (1956). 136. S. Naril, Seikagaku 32, 204 (1960); C A 60, 5815 (1964). 137. A . Askari and J. E. Franklin, Jr., BBA 110, 162 (1965). 138. B. Setlow and J. M. Lowcnstein, JBC 243, 6216 (1968). 139. J. Mendicino and J. A. Munts, JBC 233, 178 (1958). 140. A . Askuri, Nature 202, 185 (1964). 141. B. Setlow, R. Burger, and J. M. Lowenstein, JBC 241, 1244 (1966). 142. S. N. Rao, L. Ham, and A. Askuri, BBA 151, 651 (1968).

3.

67

PURINEI AMINOHYDROLASES

TABLE V KINETICCONSTANTS FOR AMP DEAMINATION BY SEVERAL PREPARATIONS OF AMINOHYDROLASE V mar

Source (Ref .)

K,

(#mole/ min/mg.)

Conditions

Rabbit muscle (199)

1.4

1660

(61, 68)

0.4

12006

Rat muscle (187)

1.4

1140

(184)

0.95

13306

0.1 M Na+ succinate, pH 6.4, 30" 0.05 M Tris-cacodylate, pH 6.3, 0.15 M KCl, 30" 0.1 M succinate, pH 6.4, 0.5 M KCl, 30" 0.02 M K + cacodylate, pH 6.5, 1 mM mercapto-

ethanol Fish-elasmobranch (196)

1.52

350

0.1 M K + succinate, pH 6.6, 30"

Carp muscle (1956)

0.4

180

0.5 M K + succinate, pH 6.4, 30"

Vmaxwas calculated from the Michaelis-Menten equation where K , is the value reported in this table and v the specific activity of the purest fraction a t the concentration of AMP used for the standard assay. Ir,,, values were obtained from double reciprocal plots.

erally th most effective monovalent cation activator although Lit, Na+, Rb', and NH,+ can often substitute. Monovalent cations are not required for the brain, muscle, and rat liver enzyme since the same v,, was observed a t high concentrations of AMP in either the presence or absence of cations. However, the enzyme associated with a brain particulate fraction (54),Ehrlich ascites tumor cells (148), and the human erythrocyte membrane (142) is reported to absolutely require K+ for activity. The soluble human erythrocyte enzyme, which constitutes 85% 143. A. R u i n and J. Mager, Israel J. M e d . Scz. 2, 614 (1966). 144. S. Nara, Seikagaku 34, 654 (1962) ; C A 58, 10456g (1963). 145. K. L. Smiley, Jr. and C. H. Suelter, JBC 242, 1980 (1967). 145a. K. L. Smiley, Jr. and C. I,. Lohrnan, Federation Proc. 26, 560 (1967). 146. M. N. Lyubimova and E. Sh. Matlina, Dokl. Akad. Nauk SSSR 94, 927 (1954) ; C A 48, 7084g (1954). 147. G. Ronca, A. Raggi, and S. Ronca-Testoni, BBA 167, 626 (1968). 148. M. R. Atkinson and A. W. Murray, BJ 104, 1Oc (1967).

TABLE VI MONOVALENT CATIONAND N~JCLEOTIDE ACTIVATORSOF AMP AMINOHYDROLASE Source

Activator

Comments

Ref.

Brain Calf

Chicken Dog Rabbit Particulate Soluble Rat

Li+ > Na+ > K+ > Rb+

> CS+ > NHI+ = (CHI)IN+ ATP > dATP > ITP > CTP = UTP Li+ > Na+ > NH4+ K+, Rb+, Cs+ no effect ATP ATP ATP ATP Na+ Na+ or Kf Na+ or K+

+

(189)

(131)

Inhibitor GTP present

(138) (183s) (139) (140)

Synergistic effect Required Activate, not required

ATP

(64) (64)

(141)

Erythrocyte Human Soluble

K+, NHI+ ATP >> ITP, GTP, UTP Na+, Li+, Rb+

Membrane bound Cat and dog

K+, Na+ ATP no effect ATP

Rabbit

ATP=ADP>GTP

Rat

Liver Na+, K+, Li+, ATP, ADP

Carp

ATP

Required No effect in absence of K+ or Na+ Activation only in presence of ATP Required

(137)

Monovalent cationsno effect

(137)

(148

(143)

Muscle

Elasmobranch fish Rabbit

K+ > Cs+ > Na+ > NH4+ > Rb+ K+ > Rb+ > N&+ Li+ > Na+ >> (CH&N+ ATP > 2'-AMP K+ = Na+ > Li+ > Rb+ NH4+ >> CS ATP, ADP N

N

Rat Chicken

K+, ADP ATP

Ehrlich ascites tumor cells Pea seed

ATP

Other

NH4

> Li' > Na+

(48)

3.

69

PUBINB AMINOHYDWLASES

of the total AMP aminohydrolase content of the cell (I,@), was activated only by K+ and NH4+ (137); activation by Na+, Li+, and Rb+ required the presence of ATP. Adenosine triphosphate alone did not activate the soluble enzyme but did lower the effective concentration for K+ activation. I n contrast the cat and dog erythrocyte enzyme were activated by A T P but not by monovalent cations either in the presence or absence of ATP (137). I n the absence of activators AMP aminohydrolase from brain (149), erythrocytes (143, 1501, muscle (145), and liver (128) gave sigmoid curves for velocity vs. AMP concentration which were hyperbolic after the addition of monovalent cations, adenine nucleotides, or a combination of monovalent cations and adenine nucleotides. For the rabbit muscle enzyme (I&), addition of K', ADP, or ATP produced normal hyperbolic saturation curves for AMP as represented by a change in the Hill slope nH from 2.2 to 1.1; V,,, remained the same. The soluble erythrocyte enzyme and the calf brain enzyme required the presence of both monovalent cations and ATP before saturation curves became hyperbolic. I n contrast, the bound human erythrocyte membrane enzyme did not exhibit sigmoid saturation curves and K activation was not affected by ATP (142).

b . Inhibition. A variety of anions such as inorganic phosphate (123, 126, 130, 143, 151-153)) sulfate (153), nitrate (153)) pyrophosphate (130, 131), tripolyphosphate (131), 2,3-diphosphoglycerate (150), creatine phosphate (153n), as well as several phosphate esters (153b), carboxylate (133, 147), F- (48, 126, 130, 136, 151, 152), #-AMP (126, 131, 1 3 S ) , G T P (128, 141, 145), G D P (145), and 3-iso-AMP (131) have been shown to inhibit AMP aminohydrolase. G T P inhibited the ATP activation of the enzyme from rat brain, heart, and liver, calf brain ( 1 4 1 ) ) and rabbit muscle (145) but had no effect on the elasmobranch fish muscle enzyme (126). G T P inhibition of calf brain enzyme was competitive with respect to ATP, apparent Ki = 10 3-ISO-AMP was not a substrate but was an effective inhibitor for the brain (apparent K i = 60 &) (131) and muscle enzyme (133). A recent study ( 1 5 3 ~ )of the effect of purine nucleotides on the K+-activated enzyme from muscle of several species showed that while

a.

149. B. Cunningham and J . M. Lowenstein, BBA 96, 535 (1965). 150. A. Askari and S. N. Rao, BBA 151, 198 (1968). 151. D.E.Kizer, B. Cox, C. A. Lovig, and S. F. DeEstrugo, JBC 238, 3048 (1963). 152. Y.-P. Lee and M. H. Wang, JBC 243, 2260 (1968). 153. A. Askari, Mol. Pharmncol. 2, 518 (1966). 153a. S. Ronca-Testoni, A. Raggi, and G . Ronca, BBA 198, 101 (1970). 153b. D.W.Sammons, H. Henry, and 0. P. Chilson, JBC 245, 2109 (1970).

70

C.

L. ZIELKE AND

C. H. SUELTER

a t low K+ concentrations ATP was an activator, a t KCl concentrations greater than 100 mM ATP as well as G T P and ITP were inhibitors. The conflict between these data and those previously reported (145,1 4 5 ~ ) may reflect the general sensitivity of this enzyme to differences in assay conditions. Carboxylic acids have been reported either to have no effect (129, 130), to activate (135),or to inhibit (133)AMP aminohydrolase. While the activation of the rabbit muscle enzyme was not thoroughly examined (135),the reported inhibition of this enzyme by citrate, succinate, and maleate was most effective in the absence of activators or in the presence of ADP (153).Enzymic activity in intact myofibrils was activated by ATP, ADP, and ITP in succinate buffer but not in citrate buffer (154). With the rat enzyme citrate, succinate, cacodylate, acetate, and lactate increased both the apparent K , and Hill slope for AMP; V,,, decreased only slightly (14'7). The inhibition by carboxylic acids, o-phenanthroline and dithioerythritol led to the finding of 2.8 g-atoms Zn2+/300,000g of rabbit muscle AMP aminohydrolase (135); 2 g-atoms Zn2+/290,000g of rat muscle The mechanism for AMP aminohydrolase has also been reported (154~). the reported inhibitions by Zn2+ (130,I % ) , Cu2+ (129,130, 155),Fe3+ (130),Ag+ (130,155),Cd2+and Ni2+ (129,155),and Hgz+ (135,136) is not understood but may involve interaction with a sulfhydryl group(s) necessary for catalysis or displacement of the presumably required zinc. Iodoacetate had no effect on the rabbit muscle enzyme (130,135) but did inhibit the carp muscle and pea seed enzyme (48,136).Organic mercurials are also reported to inhibit the enzyme from several sources (48, 125, 126, 130, 156). Except for the preliminary report by Wolfenden et al. (92) that mercurials desensitized the rabbit muscle enzyme to allosteric inhibition by GTP, the role of sulfhydryl residues in AMP aminohydrolase is not understood. 3. Mechanism

A detailed discussion of the mechanism for 5'-AMP deamination is a t present premature. The sigmoid relationship for substrate saturation and activation by monovalent cations and adenine nucleotides is consistent with mechanisms involving active site-effector site interaction. However, the activation brought about by this site-site interaction is a relatively 154. G. Kaldor, Proc. SOC.E z p t l . Biol. Mecl. 110, 21 (1962). 154a. A. Raggi, M. Ranieri, G. Taponeco, S. Ronca-Testoni, G. Ronca, and C. A . Rossi, FEBS Letters 10, 101 (1970). 155. H. Kluge and V. Wieczorek, Acta B i d . M e d . Ger. 22, 205 (1969).

3.

PURINEI AMINOHYDROLASES

71

slow first-order process independent of protein concentration (156)comparable to observations reported for yeast glyceraldehyde-3-phosphate dehydrogenase (157) and homoserine dehydrogenase (158).The activation was discussed in terms of a simple scheme similar to those proposed by Rabin (159) and Weber (160)which provides a plausible explanation for the sigmoid curve for initial velocities vs. substrate concentration without involving additional phenomena such as cooperative interactions between catalytic sites. The hydrolytic deaminstion catalyzed by rabbit and rat muscle AMP aminohydrolase may be facilitated by Zn2+(133,1 5 4 ~ )in contrast to the mediation of a common purinyl enzyme intermediate for adenosine aminohydrolase catalysis (see Section 111). 4. Considerations oj Physiological Function

As with many enzymes the role of AMP aminohydrolase in the hierarchy of metabolic catalysts is not clearly understood. Enzymic activity in muscle is markedly reduced in the dystrophic mouse (161,16?2), in humans suffering from Duchanne type muscular dystrophy (163),in hypokaliemic periodic paralysis (164), and upon denervation of normal and dystrophic mouse gastronemii (165).Activity is reported to increase in both transplanted and primary hepatomas (151) and in precancerous livers prior to the onset of neoplasia induced by feeding or by intraabdominal injections of the potent carcinogen 3’-methyl-4-dimethylaminoazobenzene (166). The weak carcinogen, 4’-methyl-4-dimethylaminoazobenzene was not effective (166).Increases in enzyme activity concomitant with altered nuclear-nucleolar morphology, nuclear RNA content, and nuclear RNA biosynthesis were also observed after injections of thioacetamide, a hepatocarcinogen (167,168). 156. C. H. Suelter, A. L. Kovacs, and E. Antonini, FEBS Letters 2, 65 (1968). 157. K. Kirschner, M. Eigen, R. Bittman, and B. Voigt, Proc. Natl. Acad. Sci. U . S. 56, 1661, (1S66). 158. E. D. Barber and H. J. Bright, Proc. Natl. Acacl. Sci. U . S. 80, 63 (1968). 159. B. R. Rabin, BJ 102, 22c (1967). 160. G. Weber, i71 “Molecular Biophysics” (B. Pullman and M. Weissbluth, eds.), p. 369. Academic Press, New York, 1965. 161. R. J. Pennington, Nature 192, 884 (1961). 162. R. J. Pennington, BJ 88, 64 (1963). 163. R. J. Pennington, Proc. Nutr. Soc. (Engl. Scot.) 21, 206 (1962). 164. A. C. Engel, C. S. Potter, and J. W. Rosevear, Nature 202, 670 (1964). 165. M. W. McCaman and R. E. McCaman, Am. J. P h y s d . 209, 495 (1965). 166. D. E. Kizer, B. A. Howell, B. C. Shirley, J. A. Clouse, and B. Cox, Cancer Res. 2 6 4 822 (1966).

72

C. L. ZIELKE AND C. H. SUELTER

A M P aminohydrolase activity was low but distinguishable in the leg, diaphragm, and heart muscle of a 20-24-day-old rabbit fetus (169). The activity in the heart remained low in both neonatal and adult life, whereas a rapid increase occurred in the activity of the enzyme in the diaphragm during the 4 or 5 days before parturition reaching a maximum activity immediately after birth. In contrast the enzymic activity of the mixed leg muscles remained relatively constant until 8-9 days after birth when it began to rise steadily with increased physical activity reaching an adult value of 7-8 times that of the fetal muscle within 14 days. Similar but qualitatively different effects were observed with guinea pig and rat leg muscle and chicken leg and pectoral muscle. Increases in aldolase, myokinase, and creatine phosphokinase activity were roughly parallel to increases in AMP aminohydrolase activity. Although it has been reported that increased AMP aminohydrolase activity occurred during prolonged stimulation of muscle bundles (170l 7 2 ) , the participation of this enzyme in the contractile process seems unlikely in light of the lack of significant changes in the levels of AMP and I M P during a single contraction of frog abdominal muscle (173). This is corroborated by the absence of AMP aminohydrolase activity in muscle of some invertebrates (174-176) and in human uterine muscle (177). It is tempting to consider regulation of the concentration of AMP, a known effector of several glycolytic enzymes, by the antagonistic action of adenine and guanine nucleotides on AMP deamination as a control factor in glycolysis and gluconeogenesis (178). Setlow et al. ( 1 4 1 ) suggested the participation of AMP aminohydrolase in a self-regulating system for purine nucleotide interconversion as presented in Fig. 1. As the G T P concentration decreases, AMP aminohydrolase inhibition is re167. D. E. Kizer, B. C. Shirley, B. Cox, and B. A. Howell, Cancer Res. 25, 596 (1965). 168. D. E. Kizer, B. A. Howell, J. A. Clouse, and B. C. Shirley, Cancer Res. 26, 1376 (1966). 169. J. Kendrick-Jones and S. V. Perry, BJ 103, 207 (1967). 170. J. Wajzer. R. Weber, J. Lerique, and J. Nekhorocheff. Nature 178, 1287 (1956). 171. E. M. Szentkiralyi. ABB 67, 298 (1957). 172. N. Moldoveanu, R e v . Roiimaine Biochim. 2, 327 (1965). 173. D. F. Cain, M. J. Kushmerick, and R . E. Davies. BBA 74, 735 (1963). 174. S. Kitagawa and Y. Tononiura. J . Biochem. ( T o k y o ) 44, 317 (1957). 175. D. Gilmour and J. H. Calaby. Enzynzologia 16, 23 (1953). 176. D. C. Cochran, BBA 52, 218 (1961). 177. T. T. Hayashi and P. S. Olmstcad, Anal. Biochem. 10, 354 (1965). 178. M. C. Scrutton and M. F. Utter, Ann, R e v . Biochem. 37, 249 (1968).

3.

PURINE AMINOHYDROLASES

1 3

73

Fumarote

ATP-----

I ----

+

Aden ylosuccinate

GDP Aspartate t P, t GTP

I I

I I

I I

I

I

“k ATP t Glutamtne AMP

+ Pyrophosphate + glutamate

I I I I I L------GTP

FIG.1. Purine nucleotide interconversions

leased with a concomitant increase in hypoxanthine and G T P which completes the self-regulating system by inhibiting the AMP aminohydrolase. I n the case of the rat and calf brain enzymes, the ATP activation and G T P inhibition were observed a t the normal i n vivo concentrations of these nucleotides and AMP (131, 155). However, such a control mechanism based upon kinetically observed changes with an in vitro system is subject to presently undefined effects by other factors i n vivo. At present only preliminary data exist as to (1) the effects of divalent metals such as Mg2+and Ca2+upon the activation and inhibition by nucleotides (138, 147, 153a) and (2) the effects of anions other than nucleotides (133, 147, 15%). Consequently, the control and function of AMP aminohydrolase remain interesting questions.

V. Adenine Nucleoside and Nucleotide Aminohydrolase (Nonspecific)

A. ADENOSINE AMINOHYDROLASE (NONSPECIFIC) Of the two homogeneous preparations of a nonspecific adenosine aminohydrolase from Aspergillus oryzae (Takadiastase) (92,179) that described by Wolfenden et al. (92) appears to be more facile and concise. Both procedures yield enzyme with turnover numbers near lo5 moles adenosine deaminated per minute and molecular weights near 215,000. The mo)

74

C. L. ZIFLKE AND C. H. SUFLTEIt

lecular weight in 8 M urea is reduced to 103,000 ( 9 2 ) ; Minato (179) observed additional dissociation in 3 M guanidine-HC1 to a 29,000 molecular weight unit or 7-8 subunits. The two preparations from A . oryzae reportedly differ in amino acid and carbohydrate composition. The enzyme prepared by Minato contained 25% carbohydrate; no cysteine was detected either by titration with p-mercuribenzoate in 6 M urea or by cysteic acid analysis after performic acid oxidation (179). In contrast, Wolfenden et al. ( 9 2 ) reported 14 cysteine residues per mole of enzyme which reacted instantaneously with p-mercuribenzoate in the absence of urea. No explanation is available for this apparent discrepancy. The enzyme catalyzes the deamination of a wide spectrum of naturally occurring adenosine derivatives including, in addition to the majority of compounds noted in Table VII, many other phosphorylated derivatives (75, 179). The I7 DFP

ko (see-l) for good substrates Esters Amides

63* 0.039"

P

OTHER

ACYLASES

Papainb

Carboxypeptidasec

Pepsind

Cysteine, histidine

Aspartic acid 2 4

Heavy metals iodoacetate

Zinc, glutamic acid tyrosine >7 Metal chelating agents

15.7, 8.5,

4669 1829

0.77h 0.29

5-8

Glutaminase ?

3.5-5.5 Heavy metals, DON (iodoacetate, DFP, chelating agents not inhibitory)

645 1265

~~

See Hess (19). b See Smith (20). 6 See Lipscomb and Hartauck (bf ) . d See Fruton (2.2). 0

N-Acetyl-lrphenylalanine derivatives, 25" (23). N-Benaoyl-karginine derivatives, 25" (2.4). Hippuryl derivatives of phenyllacta& and phenylalanine, 25" (26). h CBZ-histidyl-p-nitrophenylaianyl derivatives of phenyllactic methyl ester and phenyldanyl methyl ester, 37" (96).

6

f

3 B%

D

92

STANDISH C. HARTMAN

transferred forms an intermediate compound with a component of the enzyme. I n the case of acylases, the group transferred is the acyl moiety, and i t is well documented that a t least certain members of this class proceed through the intermediate formation of an acyl-enzyme derivative in a two-step acylation-deacylation sequence. Although the importance of this mode of action is established, in principle, an alternative path exists in the direct reaction between the ultimate nucleophilic agent (e.g., water) and the enzyme-bound substrate. There can be very little doubt about the existence of an acyl-enzyme intermediate in cases where the derivative has been isolated or otherwise directly observed. However, the implication of such an intermediate strictly by indirect kinetic means almost invariably is open to uncertainty. As mentioned above, chemically different substrates of glutaminase, including glutamic acid, the amide, the methyl ester, and the thiomethyl ester, all react in the presence of hydroxylamine to produce glutamyl hydroxamic acid. At a fixed concentration of hydroxylamine, all of these substrates show the same ratio of initial reaction rates with the amine and water. The nucleophilic agents must therefore react with a common intermediate, regardless of the substrate, since one would expect the partitioning ratio between hydroxylamine and water to vary over several orders of magnitude if a direct reaction between nucleophilea and substrates governed the kinetics (9). Despite the fact that a distinct intermediate appears to occur in the pathway, i t is unlikely that this component could be observed directly since its expected rate of reaction should exceed its rate of formation (except possibly for the best substrate, glutamic acid). This conclusion follows from the observation that the rate of the slow step in the overall hydrolytic reaction, i.e., that which controls k,, depends upon the nature of the substrate and therefore precedes the formation of the common intermediate. It is reasonable to identify the intermediate indicated by the abovementioned experiments as a 7-glutamyl-enzyme compound, an interpretation not excluded by any of the experimental results. There is, however, another plausible explanation for the observations, which does not necessarily involve a covalent enzyme-substrate compound of this kind. In this alternative proposal the rate determining steps in the catalytic reaction are not involved with the covalent bond processes but are conformational changes in the enzyme-substrate and enzyme-product complexes. If product is not released from the enzyme until a large number of rapid covalent reactions with the available nucleophiles has occurred, then any substrate will be converted to the same equilibrium mixture of bound products (e.g., glutamic acid and glutamyl hydroxamic

4.

GLUTAMINASES AND y-GLUTAMYLTRANSFERASES

93

acid) and the formation of free products observed in the steady state will be independent of substrate structure. The postulation that enzyme isomerization is rate controlling is not inconsistent with the lack of observable effect of p H upon k,, with the small solvent deuterium isotope effects and with the rather low values for enthalpies of activation. A scheme representing this type of mechanism is summarized in Eqs. ( 8 ) (12) :

(8) (9) + + + (10) + (11) (12) where E*S and E"P indicate conformational states different from the Michaelis complexes and N, and N, represent water and some other nucleophilic reactant. The fast covalent steps of Eq. (10) could in principle occur either through nucleophilic participation of a group on the enzyme or through direct displacement by the terminal acyl acceptors N, and N,. It is not possible to distinguish between these paths with present information. It is tempting to consider the (unidentified) residue which reacts with DON as a nucleophilic catalyst in the hydrolytic reactions. On the other hand, its function as an acidic or basic catalyst in a direct displacement reaction is also easily envisaged. E+SGES ES E'S E'S N1 E'P N) E'P EP EPeE+P

(slow, rate determining) (fast) (slow)

111. Other Glutaminases and Glutamyltransferases

A. SURVEY The glutamine transforming enzymes treated in this chapter may be arbitrarily placed into four categories based upon the nature of the acyl group acceptor: (1) Strictly hydrolytic. The anion-stimulated glutaminase I of animal tissues apparently is unable to utilize acceptors other than water and ammonia. Ammonia, the product of the normal hydrolytic reaction, is observed to exchange with the NH, group of glutamine in the absence of a concurrent exchange of the acyl portion of the molecule (27').Glutamyl hydroxamic acid is not a substrate, nor will hydroxylamine serve to accept the glutamyl group of glutamine ( 6 , 28). 27. J. D. Klingman and P. Handler, JBC 232, 369 (1958) 28. C. Lamar, Jr.. BBA 151, 188 (1968).

94

STANDISH C. HARTMAN

(2) Reaction with water preferred. The glutaminase of E. coli preferentially catalyzes hydrolysis of glutamyl derivatives but will also utilize acceptors such as hydroxylamine and methanol. Larger amine acceptors are excluded. The glutaminases from Pseudomonas (29, 30) and from Azotobacter agilis (12) are probably of this sort. The latter enzymes act upon D- as well as L-glutamine, and upon asparagine. (3) Reaction with amine acceptors preferred. Hydrolysis of glutamyl compounds occurs with the transferase from mushrooms (31) and with one type of transpeptidase isolated from kidney ( 3 2 ) . However, when suitable amine acceptors are present the hydrolytic reaction is largely suppressed in favor of transfer to the amine. (4) Totally nonhydrolytic. The glutarnyl transferase from Proteus vulgaris acts upon glutamine as substrate and utilizes hydroxylarnine, hydrazine, and ammonia as acceptors, but it is apparently devoid of any glutaminase activity ( 3 3 ) . Glutamyl units derived from L- or Dglutamine are polymerized into a y-linked polymer by an enzyme from Bacillus subtilis ( S 4 ) , a function which is similar to that of the kidney transpeptidase except that the bacterial enzyme does not appear to catalyze hydrolytic reactions. An enzyme from pig kidney, distinct from the hydrolase-transpeptidase mentioned in category (3) catalyzes the transfer of glutamyl units from glutathione (but not glutamine) to certain peptides such as glycylglycine without formation of glutamic acid under any condition ( 3 5 ) . The “7-glutamyl cyclotransferase” found in animal tissues, including liver and brain, might be considered a special member of this group (36).This enzyme effects an intramolecular acyl transfer by converting certain y-glutamyl peptides to the cyclic pyrrolidone carboxylic acid (see Orlowski and Meister, Chapter 6, this volume). Glutamine is not a substrate, and glutathione is acted upon very slowly. A cyclotransferase with somewhat different properties has been obtained from papaya latex (37). L-Glutamine and N-terminal a-glutaminyl peptides lacking substituents on the y-NHz group are cyclized to the corresponding pyrrolidone carboxylic acid derivatives. Several other enzymes which catalyze y-glutamyl transfer reactions 29. M. A. Ramadan and D. M. Greenberg, Anal. Biochem. 6, 144 (1963). 30. K. Soda, K. Uchiyama, and K. Ogata, Agr. Biol. Chem. (Tokyo) 30, 547 (1966). 31. H. J. Gigliotti and B. Levenberg, JBC 329, 2274 (1964). 32. M. Orlowski and A. Meister, JBC 240, 338 (1965). 33. H. Waclsch, “Methods in Enzymology,” Vol. 2, p. 267, 1955. 34. W. J. Williams and C. B. Thornc, JBC 210, 203 (1954). 35. F. H. Leibach and F. Rinkley, A R B 127, 292 (1968). 36. M. Orlowski, P. G. Richman, and A . Mrister, Biochemislry 8, 1048 (1969). 37. M. Messer and M . Ottesen, BBA 92, 409 (1964); Compt. Rend. Tmv. Lob. Cnrlsbcig 35, 1 (1965).

4.

GLUTAMINASES AND 7-GLUTAMYLTRANSFERASES

95

have been described, but insufficient information is available to allow their classification in the above scheme. These would include the glutaminase from Clostridium welchii (16),an anion-activated enzyme derived from the microsomal fraction of rat kidney which hydrolyzes glutamine and glutamyl hydroxamic acid and which is distinct from and a hydrolase from pig kidney which cleaves “glutaminase I” (,%I), glutamyl naphthylamide but will not transfer the glutamyl unit to peptide acceptors (35). Some of the above-mentioned enzymes about which significant enzymological information is availatde are singled out for further description in the following sections.

B. ~GLUTAMYLTRANSFERASE FROM AGARICACEAE Gigliotti and Levenberg (31) have described an enzyme isolated from certain species of mushrooms of the genus Agaricus which is associated with the occurrence of agaritine (y-L-glutamyl-p-hydroxymethyl-phenylhydrazide) and y-L-glutamyl-p-hydroxyanilide. Partial purification (20fold) yielded a preparation capable of hydrolyzing a number of y-acyl derivatives of L-glutamic acid in a narrow region of pH, with optimum about 7. When rates of hydrolysis a t a fixed concentration of substrate (1 mM) were determined, the phenylhydrazide, p-hydroxyphenylhydraaide, p-hydroxyanilide, cyclohexylamide, ethylamide, and benzyl ester were rapidly cleaved, while glutamine, glutamyl hydrazide, and glutamyl 1-naphthylhydrazide were acted upon somewhat more slowly. Glutathione and other y-glutamyl peptides were not substrates. Apparently, a bulky hydrophobic substituent is preferred in contrast to the situation with the E. coli glutaminase. In the presence of hydroxylamine, ammonia, phenylhydrazine, or p-hydroxyaniline, transfer of the glutamyl unit from substrates to these amines is observed. The rate of hydrolysis and transfer to hydroxylamine (at 0 . 2 5 M ) for a given substrate are approximately equal. I n this respect, the behavior of the Agaricus and E. coli enzymes are rather similar. When an acceptor amine containing an aromatic group is present (e.g., phenylhydrazine) , hydrolysis is almost completely suppressed in favor of transfer to the aromatic amine, and in this sense the mushroom enzyme is best considered a transferase. These results are consistent with the view that the enzyme has a hydrophobic site which can be occupied either by the substituent group of the substrate or by the acceptor amine. Water or other small amine acceptor can react with a bound derivative of glutamic acid if this site is not occupied, but a t a slower rate than that which occurs with the hydrophobic amine. The possibility that the enzyme may depend upon sulfhydryl groups

96

STANDISH C. HARTMAN

for activity is indicated by its sensitivity to inhibition by mercuric, cupric, and zinc ions, mercuribenzoate, and iodoacetate.

C. Y-GLUTAMYL TRANSPEPTIDASES FROM KIDNEY The ability of enzyme preparations from various animal tissues to catalyze transfer of the glutamyl unit from glutamine or glutathione to certain a-amino acids and peptides, first observed by Hanes et al. (38),has been examined in several laboratories (32,35,39-41). Enzymes of this type have been studied recently by Orlowski and Meister (in hog kidney) (32)and by Szewczuk and Baranowski (in beef kidney) (41). Results with the hog kidney enzyme will be reviewed here; the beef enzyme is quite similar in many respects, but it is not identical in physical properties. After solubilization from a particulate fraction of hog kidney cortex, the transpeptidase has been obtained in highly purified, but not yet homogeneous, form. The enzyme is active in the alkaline p H region with a maximum at about p H 8.8. Although appreciable magnesium is present in the enzyme preparation and added Mg2+ partially stimulates the catalytic action, it is not certain whether this metal is essential for activity since EDTA exerts an enhancement rather than an inhibition. The transpeptidase catalyzes y-glutamyl transfer from a number of compounds in addition to glutathione and glutamine, including glutamyl p-nitroanilide, glutamyl naphthylamide, y-benzyl glutamate, y-ethyl glutamate, and y-glutamyl peptides. Derivatives of D-glutamic acid are also substrates, although they react a t only about 10% the rate of the L-isomers. Asparagine and homoglutamine are not utilized. When the enzyme acts upon one of the above substrates some glutamic acid is produced in addition to products formed by transfer of the y-glutamyl moiety from one molecule of substrate either to the a-amino group of the product, glutamic acid, or to that of another molecule of substrate, to form a y-linked peptide. Subsequent addition of glutamyl units to the dipeptides can build up oligopeptides of glutamic acid. Eventually, since the reactions are reversible, the thermodynamically favored hydrolytic process overrides the kinetically favored transpeptidation reaction and glutamate ion results as the predominant product. If one of a large number of a-amino acids or peptides is present, it 38. 39. 40. 41.

C. S. Hanes, F. J. R. Hird, and F. A. Isherwood, BJ 51, 25 (1952) P. J. Fodor, A. Miller, and H. Waelsch, JBC 202, 551 (1953). E. G. Ball, J. P. Revel, and 0. Cooper, JBC 221, 895 (1956). A. Szewczuk and T. Baranowski, Biochem. Z. 338, 317 (1963).

4.

GLUTAMINASES AND y-GLUTAMYLTRANSFERASES

97

will serve as acceptor of the glutamyl unit to form the corresponding y-glutamyl peptide. Hydroxylamine will also accept the acyl group. It is evident that the hog kidney transpeptidase exhibits relatively little specificity with respect to substituent groups on the substrate (or acyl acceptor). I n contrast to the transferase from mushrooms, the transpeptidase is particularly reactive with amines containing polar substituents such as amino acids and peptides. Another type of glutamyl transpeptidase (or transferase) is obtainable from hog kidney, as shown by Leibach and Binkley (35). The enzyme is obtained in soluble form by treating renal microsomes with ficin, after which it has been purified 2000-fold to apparent homogeneity. Differences between this enzyme and the one described by Orlowski and Meister are striking, even though a detailed examination of the specificity of the microsomal enzyme has not been reported. The latter system catalyzes glutamyltransfer from either reduced or oxidized glutathione to certain peptide acceptors such as glycylglycine: Glutathione

+ glycylglycine S cysteinylglycine + -pglutamylglycylglycine

(13)

Neither glutamine nor glutamyl naphthylamide are substrates. The specificity requirements for the acceptor are considerably more demanding in that acyl transfer to water (hydrolysis) or to hydroxylamine is not observed under any conditions. The transferase was separated chromatographically from an enzyme capable of hydrolyzing glutamyl naphthylamide which conceivably is related to the previously described t ranspeptidase.

D. GLUTAMINASE FROM Azotobacter agilis Ehrenfeld et al. have described the purification of a glutaminaseglutamyltransferase from this soil bacterium ( l a ) . Although not completely pure, the enzyme was observed to have a sedimentation coefficient of about 4.4-4.8 S. It not only catalyzes hydrolysis of L-glutamine (relative rate, 1) but also that of D-glutamine (0.57), L-asparagine (0.7), Dasparagine (0.24), succinamic acid (0.15), and glutamyl hydroxamic acid. A number of other amino acid amides, as well as glutamyl y-ethylamide and y-ethyl glutamate, are not substrates. The enzyme exhibits a remarkably broad pH optimum, the V,,, for L-glutamine being constant between pH 4.5 and 9. Glutamyl hydroxamic acid is produced by action of the enzyme on substrates in the presence of hydroxylamine. Hydroxylaminolysis and hydrolysis are about equal in rate a t p H 7.2 when the hydroxylamine concentration is 0.5 M. The rate of formation of hydrox-

98

STANDISH C. HARTMAN

amic acid a t fixed total hydroxylamine concentration is not constant with pH, but rather it increases between pH 4.5 and 6.5 in the manner expected if the basic form of hydroxylamine were the reactant. When expressed in terms of the basic form of the amine, the relative rates of transfer to water and hydroxylamine for this enzyme are almost identical to those observed with E . coli glutaminase. Glutamic acid is also a substrate of the Azotobacter enzyme since it reacts readily with hydroxylamine to form the hydroxamic acid, the reverse of which process is the hydrolysis of the latter substance. The equilibrium constant for the formation of the hydroxamic acid from glutamic acid and hydroxylamine has been determined with the aid of this enzyme. I n the absence of glutamine the diazo analogs, DON and azaserine, strongly and irreversibly inhibit the enzyme. Unlike the E . coli enzyme, this one is insensitive to mercuribenzoate. Ethylenediaminetetraacetate is not inhibitory. Evidence is presented that glutamic acid binds to the enzyme in a form that dissociates to free glutamic acid more slowly than it reacts with hydroxylamine. When the enzyme is briefly exposed to 14C-glutamic acid, then hydroxylamine and a large amount of unlabeled glutamic acid are added, the initial molecules of glutamyl hydroxamic acid formed have an excess of radioactivity over that expected from the average specific activity of the glutamic acid in the mixture. The authors suggest that the formation of a covalent glutamyl-enzyme intermediate could account for these results. The alternative possibility advanced in the discussion of the E . coli glutaminase could be offered here as well: the bound form of substrate is not covalently linked to enzyme but is separated kinetically from free substrate by slow conformatianal changes.

IV. Concluding Remarks

That formal similarities occur among the reactions discussed above raises the question of whether the enzymes responsible might be related structurally, mechanistically, and perhaps, evolutionarily. Insofar as the limited information available allows comparison there would seem to be no obvious basis for supposing that a common derivation or mode of action of these enzymes exists. While data on structure are scanty, variability in functional groups is evidenced by the disparate pH-activity profiles and spectra of sensitivity toward inhibitors, for example. AS a first approximation, such criteria set the limits for the kinds of groups which are essential for catalytic action.

4.

GLUTAMINASES AND 7-GLUTAMYLTRANSFERASES

99

The most evident phenomenological differences among these enzymes reside in their abilities to utilize diverse acceptors for the glutamyl moiety transferred by each. Specificities with respect to acceptor are a consequence of the specificities shown toward substrates, both in principle and experimentally. The acceptor is mechanistically equivalent to the 7-substituent group of a substrate since the process in which the C-N bond is formed is microscopically the reverse of the C-N bond cleavage. Those and only those substituent groups which are tolerated by the enzyme in the direction of bond cleavage will be inserted in the “acyl transfer’:. reaction. Thus, for example, an enzyme capable of transferring th y-glutamyl group to hydroxylamine must necessarily utilize the hydr amic acid as a substrate, and vice versa, otherwise the enzyme would bk a one-way catalyst. It may sometimes be difficult to observe transfer .reactions with certain substrates and acceptors which themselves are normal reaction products (e.g., glutamic acid or ammonia). In these cases the difficulty is undoubtedly an experimental one of not being able to achieve significant concentrations of the proper ionic forms of the reactant in the pH range where the enzyme is active. The observation that acyl groups (including those derived from glutamic acid) are transferred to amines, especially hydroxylamine, is often regarded as evidence for the formation of an “activated” intermediate, in analogy to the reactions by which acyl phosphates or acyl thiols may be trapped. However, the view that such an intermediate is a compound of large free energy of hydrolysis is objectionable on the ground that it would seem to require an unnecessarily large energy barrier in the reaction pathway. Any reasonable mechanism for the hydrolysis of amide or hydroxamic acid, whether or not it involves an intermediate activated in this sense, provides a path, in its reverse, for the synthesis of the amide. In discussing possible mechanisms for the reactions catalyzed by E . coli glutaminase in Section I, it was concluded that either a two-step acylation-deacylation pathway or a one-step route, displacement by the ultimate nucleophile, could be accommodated by the results. It may be noted that any single displacement mechanism for a group transfer reaction requires that both incoming and outgoing substituent groups associate with the enzyme a t the same time

i

0

II

x1-c-xp I R

in contrast to the case with a double displacement mechanism in which only one X need be present (the other one is a component of the enzyme).

100

STANDISH C. HABTMAN

At least thii is the case for enzymes which show a selectivity with respect to the substituent or acceptor, a situation indicating that the agent does not react from the solution phase. The single displacement mechanisms, then, require the existence of two distinct and specific sites for recognition and reaction of XIand X,, capable of carrying out the mechanistically symmetrical events of catalysis. It is not too difficult to imagine a hydrolytic reaction occurring in this way, in which one group is H,O (or OH-). However, in the acyltransferase reactions in which water is not a reactant, two often bulky and structurally complex groups would need to be accommodated at once. If the principle of simplicity in nature carries any force, one would be inclined to favor the two-step displacement pathway to describe these reactions. In this event the enzyme would need to possess only one site for interaction with any acyl donor or acceptor. The simplification would not extend to the reaction mechanism however, since the same type of acyl transfer must be catalyced in either case; uiz., transfer to the dissociable acceptor or transfer to an interim carrier which is part of the enzyme.

JOHN C. WRISTON, JR. I. Introduction . . . . 11. Occurrence . . . . 111. Guinea Pig Serum Asparaginase A. Isolation . . . B. Properties . . . IV. EscheTichia coZi Asparaginase A. Isolation . . . B.Pmperties . . . V. Other Asparaginases . . . VI. Physiological Properties . .

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101

102 105 105 106 107 107 109 116 117

1. Introduction

L-Asparaginase (L-asparagine amidohydrolase, EC 3.5.1.1.) that catalyzes the hydrolysis of L-asparagine to L-aspartic acid and ammonia is widely distributed. In addition, there is at least one other enzyme system capable of catalyzing the hydrolysis of asparagine. Meister has demonstrated (1,2)that rat liver, for example, contains an asparaginase activity which is a-keto acid dependent, and which actually results from the presence of an w-amidase (a-amidodicarboxylate amidohydrolase, EC 3.5.1.3) that catalyzes the hydrolysis of the a-keto succinamate formed from asparagine by transamination. Rat liver extracts also appear to contain a true L-asparaginase, but one which is phosphate dependent 1. A. Meister, H. A. Sober, 8.V. Tice, and P. E. Fraser, J . BWZ. Chem. 197, 319 (1952). 2. A. Meister, L. Levintow, R. E. Greenfield, and P. A. Abendschein, J . Biol. Chem. 215, 441 (1955).

101

102

JOHN C. WRISTON, JR.

and relatively heat labile, properties which tend to differentiate it from other asparaginases that have been more thoroughly studied. Asparagine deamidation can also be accomplished by o-replacement reactions (S) . Little new information has been reported concerning either the a-keto acid-dependent asparaginase or the phosphate-dependent asparaginase, and this article will focus almost entirely on the so-called true asparaginases.

II. Occurrence

Asparagine hydrolyzing activity is widely distributed [for earlier reviews, see references ( 4 4 1 , but in many of the earlier reports describing the formation of ammonia from added asparagine by cell suspensions or crude cell extracts over a period of hours or even days, it is not possible to ascribe the activity to a true asparaginase. For this reason, a historical survey will be avoided here, and the examples that will be noted are those where there is reasonable assurance that the enzyme being considered is kasparagine amidohydrolase. I n 1953, Kidd reported (7‘) that whole guinea pig serum could bring about the regression of certain transplanted lymphosarcomas in inbred mice. His experiments indicated that the substance responsible for the antilymphoma activity was a protein, and in 1961 Broome (8) presented evidence linking this activity to the presence of the enzyme L-asparaginase in the guinea pig serum. These observations, together with the discovery that an E . wli asparaginase also had antilymphoma activity (9),have been reponsible for a marked increase in interest in asparaginase since 1961. The presence of asparaginase in guinea pig serum was first reported by Clementi (10), who also noted the absence of this enzyme in the sera of a number of other common mammals including the rat, cat, dog, monkey, and human. The list of animal sera from which asparaginase H. Waelsch, Aduan. Enzymol. 13, 237 (1952). J. E. Varner, “The Enzymes,” 2nd ed., Vol. 4, p. 243, 1960. C. A. Zittle, “The Enzymes,” 1st ed., Vol. 1, Part 2, p. 922, 1951. A. Meister, “Biochemistry of the Amino Acids,’’2nd ed., Vol. 2, p. 606, Academic Press, New York, 1965. 7. J. G. Kidd, J . Exptl. Med. 98, 565 and 582 (1953). 8. J. D. Broome, Nature 191, 1114 (1961). 9. L. T. Mashburn and J. C. Wriston, Jr., Arch. Biochem. Biophys. 105,451 (19&1). 10. A. Clementi, Arch. Intern. Physiol. 19, 369 (1922). 3. 4. 5. 6.

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103

is absent has subsequently been extended to include the chicken, pig, sheep, cow, horse, and nutria (11-13), although rabbit serum has been reported to contain traces of activity (11).Clementi also reported that asparaginase was present in the liver and kidney of certain birds. The enzyme has also been found in fish liver, but not in the liver of frogs ( 1 4 ) , in guinea pig liver (16-I?),in chicken liver (18), and in the serum of several animals closely related to the guinea pig (12, 19). Serum from all members of the superfamily Cavioidea, to which the guinea pig belongs, contain asparaginase. Agouti serum, in particular, was found to contain several times as many units of activity per milliliter as guinea pig serum, and this fact, coupled with the larger size of the animal, would appear to make it a more practical source of mammalian enzyme than the guinea pig. Members of three other hystricomorph families related to the Cavioidea do not contain serum asparaginase. It has also been found that the serum of newborn guinea pigs contains very little asperaginase (8,11,20) and that the levels do not appear to be affected by strain, sex, or dietary intake of asparagine ( 1 1 ) . Asparaginase is present in the serum of two species of new world monkeys, but not in three old world species ( 2 1 ) . The guinea pig serum enzyme has been purified to apparent homogeneity and is discussed in more detail in a later section. Several asparaginases from microorganisms were described prior to 1961. Asparaginase has been found in Aspergillus niger (22, 23), and is also present in yeasts (24-27). Extracts of Mycobacterium smegmatis, 11. D. B. Tower, E. L. Peters, and W. C. Curtis, J . Bwl. Chem. 238, 983 (1963). 12. N. D. Holmquist, Proc. SOC.Exptl. BWZ. M e d . 113, 444 (1963). 13. M. B. Lee and J. M. Bridges, Nature 217, 758 (1968). 14. G. Steensholt, Acta Physiol. Scand. 8, 342 (1944). 15. H. A. Krebs, BWchem. J . 47, 605 (1950). 16. N. DeGroot and N . Lichtenstein, Biochim. Biophys. Acta 40, 92 (1960). 17. H. M. Suld and P. A. Herbut, J . Biol. Chem. 240, 2234 (1965). 18. T. Ohnuma, F. Bergel, and R. C. Bray, Biochem. J . 103, 238 (1967). 19. L. J. Old, E. A. Boyse, H. A. Campbell, and G . M. Dana. Nature 198, 801 (1963). 20. H. Ainis, H. M. Kurtr, P. I. Kramer, H. E. Weimer, R. M . Ryan, and E. Jameson, Cancer Res. 18, 1309 (1958). 21. R. Arrison, personal communication, cited in J. D. Broome, Trans. N . Y . Acnd. Sci. [21 30, 690 (1968). 22. D. Bach, Compt. Rend. 187, 955 (1928). 23. K. Schmalfuss and K. Mothes, Biochem. Z . 221, 134 (1930). 24. W. Grassman and 0. Mayr, Z . Physiol. Chem. 214, 185 (1933). 25. G. Gorr and J. Wagner, Biochem. Z . 254, 1 (1932). 26. J. D. Broome, J . NatZ. Cancer Znst. 35, 967 (1965). 27. A. K. Abdumalikov and A. Y. Nikolaev, Bwkhimiya 32, 4 (1967).

104

J O H N C. WRISTON, J R .

M . phlei, and several strains of M . tuberculosis (2830)contain a true asparaginase, as do extracts of Bacillus coagulans and Bacillus stearothemnophilus (31).Brucella abortus contains two asparaginases, one specific for L-asparagine and the other for the opposite enantiomorph (3.2). An asparaginase is also present in Pseudomonas fluorescens ( 3 3 ) .Also, Tsuji, in 1957, had reported the presence of asparaginase in extracts of acetone powders of E . coli, Staphylococcus, M . avium, and Aspergillus oryzae (34). Since 1961, several of the previously reported asparaginases have been reexamined for antilymphoma activity, and a number of additional asparaginases have been found. Mashburn and Wriston (9) reported that an L-asparaginase from E . coli B had antilymphoma activity but that the B . coagulans enzyme did not. The discovery of a bacterial source of the enzyme with antilymphoma activity made it possible to prepare large amounts of enzyme and carry out extensive clinical studies, which are still under way. Several purification schemes have been reported, and the E . coli asparaginase is also discussed in detail in a later section. Asparaginases with antilymphoma activity have also been found in extracts o f Serratia marcescens (36-57), Erwinia carotovora (58), and Proteus vulgaris ( 3 6 ) . Escherichia w l i extracts contain a second asparaginase that is devoid of antilymphoma activity (39),and a similarly inactive enzyme has been found in a fungus, Fusarium tricinctum (40). An enzyme from Streptomyces griseus has L-asparaginase activity when assayed in tris-Cl (pH 8.6) but little or no activity in sodium borate (pH 28. W. F. Kirchheimer and C. K . Whittaker, Am. Rev. Tuberc. 70, 920 (1954). 29. Y. S. Halpern and N. Grossowicz, Biochem. J . 65, 716 (1957). 30. H. N. Jayaram, T. Ramakrishnan, and C. S. Vaidyanathan, Arch. Biochem. Biophys. 126, 165 (1968). 31. G. B . Manning and L. L. Campbell, Jr., Can. J . Mimobiol. 3, 1001 (1957). 32. R. A. Altenbern and R. D. Housewright, Arch. Biochem. Biophys. 49, 130 (1954). 33. N. DeGroot and N. Lichtenstein, Biochim. Biophys. Acta 40, 99 (1980). 34. Y. Tsuji, Naika Hokan 4, 222 (1957); CA 53, 10350 (1957). 35. B. Rowley and J. C. Wriston, Jr., Bwchem. Biophys. Res. Comm. 28, 160 (1967). 36. J. W. Boyd, M.S. Thesis, University of Delaware, 1965. 37. R . E. Peterson and A. Ciegler, Appl. Microbiol. 18, 64 (1969).

38. H. E . Wade, R. Elsworth, D. Herbert, J. Keppie, and K . Sargeant, Lancet i, 776 (1968). 39. H. A. Campbell, L. T. Mashburn, E . A. Boyse, and I,. J. Old, Biochemistry 6, 721 (1967). 40. R . W. Scheetz, Ph.D. Thesis, University of Delaware, 1969.

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105

8.5) (41). Peterson and Ciegler have recently surveyed 123 species of bacteria for L-asparaginase activity and found the highest yields in an Erwinia aroideae strain (42). It is interesting to note that some of the E . coli strains they examined contain no asparaginase. Roberts et al. (43) have listed several additional asparaginase-containing microorganisms but found no asparaginase activity in several molds or yeast that they examined. An amidase, apparently a single enzyme, that catalyzes the hydrolysis of both asparagine and glutamine and that appears to have only limited antilymphoma activity, has been reported in extracts of Pseudomonas (44-46). Lee and Bridges (13) have confirmed the absence of asparaginase in the serum of a variety of animal species and in 25 human sera but have observed that human and sheep sera, themselves devoid of detectable asparaginase activity, enhanced the activity of guinea pig serum (GPS). Other sera did not show this effect with the GPS enzyme, but all the sera enhanced E . coli asparaginase activity. Further work by these authors (47) and by Ho and Jones (48) indicates that the effect results from a nonspecific stabilization by plasma proteins. The presence of asparaginase has also been reported in certain plant tissues (49, 5 0 ) , but a recent study by Lees et al. ( 5 1 ) , in which they used U-14C-asparagine to provide a sensitive assay system for asparagine conversion to aspartic acid, failed to show any asparaginase activity in extracts of wheat or lupin seedlings. 111. Guinea Pig Serum Asparaginase

A. ISOLATION Several partial purifications of guinea pig serum asparaginase have been reported, beginning with a sevenfold purification by Meister in H. A. Campbell and L. T. Mashburn, Biochemistry 8, 3768 (1969). R. E. Peterson and A. Ciegler, A p p l . Microbiol. 17, 929 (1969). J. Roberts, G. Burson, and J. M. Hill, J. Bacteriol. 95, 2117 (1968). M. E. A. Ramadan, F. El Asmar, and D. M. Greenberg, Arch. Biochem. Biophys. 108, 143 and 150 (1964). 45. F. A. El Asmar, and D. M. Greenberg, Cancer Res. 26, 116 (1966). 46. L. P. Evseev, A. Y. Nikolaev, V. V. Eremenko, and S. R. Mardashev, Biokhimiya 32, 873 (1967). 47. M. B. Lee and J. M. Bridges, personal communication (1969). 48. P. P. K. Ho and L. Jones, Bwchim. Biophys. Acta 177, 172 (1969). 49. C. E. Grover and A. C. Chibnall, Biochem. J. 21, 857 (1927). 41. 42. 43. 44.

106


1955 (11, 17, 69-55). Purification to apparent homogeneity was reported by Yellin and Wriston in 1966 ( 5 6 ) . Their procedure, described in detail elsewhere ( 5 7 ) , involves salt fractionation with Na2S0,, gel filtration on G-200 Sephadex, chromatography on DEAE-cellulose, followed by chromatography on calcium hydroxylapatite. A 900-fold purification was achieved with an overall recovery of about 10%. Asparaginase accounts for about 0.1% of the total protein of guinea pig serum, and the amount recovered corresponds to less than 1 mg of homogeneous enzyme from 100 ml of GPS, with a specific activity of about 470 I U mg (57a). The enzyme was judged to be homogeneous on the basis of immunoelectrophoresis, analytical ultracentrifugation studies, and polyacrylamide gel electrophoresis. The homogeneous enzyme was found (58) to have antilymphoma activity toward the Gardner lymphosarcoma in C3H mice comparable to that of the guinea pig serum itself, thus confirming Broome’s original suggestion.

B. PROPEF~TIES Guinea pig serum asparaginase has a molecular weight of approximately 138,000 (equilibrium sedimentation) and- an s;,~ of 6.55 S ( 5 6 ) . The purified enzyme is stable for at least 6 months at -20”, to repeated freezing and thawing, and to heating at 55” for 10 min (11,56).It is also stable to dialysis in the cold but is labile under conditions which promote surface denaturation. Whole guinea pig serum can be stored for several weeks a t 4” without appreciable loss of asparaginase activity. It has also been reported (15) that a dry powder obtained from whole guinea pig serum by precipitation with two volumes of cold ethanol can be stored in a desiccator for a year or more with no appreciable loss of 50. W. L. Kretovich, Advan. Enzymol. 20, 319 (1958). 51. E. M. Lees, K. J. F. Farnden, and W. H. Elliott, Arch. Biochem. Biophys. 126, 539 (1988). 52. A. Meister, “Methods in Enzymology,” Vol. 2, p. 383, 1955. 53. S. R. Mardashev and V. Shao-Khua, Dokl. Akad. Nauk SSSR 142, 709 (1962). 54. L. T. Mashburn and J. C. Wriston, Jr., Biochem. Biophys. Reg. Commun. 12, 50 (1963). 55. J. D. Broome, J . Exptl. Med. 118, 99 and 121 (1963). 56. T. 0. Yellin and J. C. Wriston, Jr., Biochemitry 5, 1605 (1966). 57. J. C. Wriston, Jr., “Methods in Enzymology,” 1971 (in press). 57a. The international unit (IU) is defined as that amount of enzyme which will catalyze the release of 1 pmole of ammonia per minute under standard assay

conditions. 58. T. 0. Yellin and J. C. Wriston, Jr., Science 151, 998 (1966).

5. L-ASPAEAGINASE

107

activity. The enzyme has the electrophoretic mobility in starch of an a-2-globulin ( 8 ) . The optimal pH range (in 0.1 M sodium borate buffer) is from 7.5 to 8.5 (52),although Tower et al. reported (11) a pH optimum of 9.6 for 100-fold purified enzyme in buffers of lower ionic strength than those used by Meister (r/2 = 0.2 instead of 0.4). The K , value has been reported as 2.2 X lo-’ M (11) and 7.2 X M (18). Broome has also shown (59)that the apparent K , for agouti serum asparaginase is 4.1 X M. The enzyme is inhibited by p-mercuribenzoate and HgCl, (40 and 83% inhibition, respectively, at 0.1 mM) but not by L-glutamic acid or Lglutamine, 10 mM; L-aspartic acid, 20 mM; NH,CI, 50 mM; alanine, oxaloacetate, a-ketoglutarate and pyruvate, 5 mM; FOB,-, 50 mM; SO:-, 5 mM; MgZ+,Ca2+,and MnZ+,5 mM; iodoacetate, fluoride, and cyanide, 1OmM; and N-ethylmaleimide and pyridoxal phosphate, 1 mM (11). Of a large number of amides examined, only L-asparagine (loo%),D-asparagine (3%), L-leucine amide (5.3%), L-phenylalanine amide, (8.2%), and L-tyrosine amide (8.5%) were hydrolyzed (2, 11). L-Glutamine is among those amides not hydrolyzed. The enzyme will not hydrolyze N-acetyl-Lasparagine or N-L-glutaminyl-L-asparagine (11). Guinea pig serum asparaginase also catalyzes hydrolysis of L-p-aspartyl hydroxylamine and synthesis of the hydroxamate from asparagine and hydroxylamine, but these reactions proceed more slowly than the hydrolysis of L-asparagine ( 2 ) . The amino acid composition has been determined (56).Guinea pig serum asparaginase has been used occasionally for the quantitative determination of asparagine (11, 16, 60).

IV. Escherichia coli Asparaginase

A. ISOLATION Tsuji (34) was the first to report the presence of L-asparaginase in

E . coli extracts. In 1964, Mashburn and Wriston (9)described a partial purification of the enzyme from E . coli B, and found that this bacterial asparaginase possessed antilymphoma activity. Purification to apparent homogeneity was reported in 1968 (61). Roberts et al. (43) in that same year also described an extensive purification in high yield of E . coli 59. J. D. Broome, Bn’t. J . Cancer 22, 595 (1968). 60. B. F. Sansom and J. M. Barry, Biochem. J . 68, 487 (1958). 61. H. A. Whelan and J. C . Wriston, Jr., Biochemistly 8, 2386 (1969); Federation Proc. 27, 586 (1968).

108


L-asparaginase. Several pharmaceutical firms have developed greatly simplified purification schemes making it feasible to produce large amounts of highly purified enzyme for clinical trials (62-65). The steps used in the first purification to homogeneity are shown in Table I [for details, see Wriston (57)l. In the procedure developed by Roberts et al. (43) there is an alcohol fractionation, DEAE-cellulose chromatography, an isoelectric precipitation with alcohol, chromatography on carboxymethyl Sephadex, and a final prep disc electrophoresis step. Ho et al. (6%) obtained crystalline L-asparaginase from E. coli B by a procedure involving essentially ammonium sulfate and ethanol fractionation, and they reported an h’ig of 7.1 for the dry crystalline enzyme. The Bayer group (63-65) have also crystallized E . coli asparaginase recently. I n one case this was done by using 50% aqueous polyethylene glycol (average molecular weight 1500) with resuspended acetone powders of E. coli cells; and in another it was done by a combination of heat denaturation, acetone and 2-methylpentane-2,4-diol fractionation, and Sephadex G-200 gel chromatography. TABLE I PURIFICATION OF E. coli B ASPARAGINASE~

Step

Sample vol. (ml)

Total protein (mg)

Total units

Specific activity (units/mg)

Crude extract MnCls, heat P-150 Bio-Gel D EAE-cellulose Calcium hydroxylapatite Prep disc electrophoresis

140 90-100 8-10 40-50 25 4

12-14 1800 1200 500-800 3-6 1.5-2

3500 2400 2200 1600 800 600

0.2-0.25 1.3 1.8 20-32 150-250 300-400

Stepwise recovery

(%) 65 90 70 50 90

From 20 g of lyophilized cells. Experimental details are given in the text. Recoveries in ultrafiltration and dialysis were essentially quantitative, and these steps are omitted in the table. Overall purification of approximately 2000-fold with 15% recovery. 62. P. P. K. Ho, B. H. Frank, and P. J. Burch, Science 165, 510 (1969); P. P. K. Ho, E. B. Milikin, J. L. Bobbitt, E. L. Grinnan, P. J. Burck, B. H. Frank, L. D. Hoeck, and R. W. Squirrs, J . Bid. Chem. 245, 3708 (1970). 63. K. Bauer, A. Arens, E. Rauenbusch. E. Irion, 0. Wagner. W. Kaufmann, W. Scholtan, and S. Y. Lie, Abslr. 6th. FEBS Meeting, Madrid, 1969 p. 93. Academic Press, New York, 1970. 64. 0. Wagner, K. Bauer, E. Irion, E. Rauenbusch. W. Kaufmann, and A. Awns Angew. Chem. Intern. Ed. 8, 885 (1969). 65. A. Arens, E. Rauenbusch, E. Irion, 0. Wagner, K. Bauer, and W. Kaufrnuiiii. 2.Phy8M1. Chem. 351, 197 (1970).

5.

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109

B. PROPERTIES 1. General

The content of asparaginase varies widely in different strains of E . coli (39,4.2, 4.9) and there are certain differences in properties as well. There are two quite different asparaginases in E. coli B cells, designated EC-1 and EC-2 by Campbell et al. (391,which can be separated by DEAEchromatography or by differential heat inactivation. Only EC-2, the relatively heat-stable asparaginase, has antilymphoma activity, and only this enzyme has been purified and studied in detail. The two enzymes differ in several other respects and can be distinguished by taking advantage of marked differences in their pH-activity profiles. A broad pHactivity profile is shown by EC-2 with a maximum a t about 8, while EC-1 shows a rapid decrease in activity below pH 8.4. This has served as the basis for a differential assay a t pH 5 and 8.5 (39).Two asparaginases have also been detected in E. coli K-12 (66) and here a differential assay has been based on the fact that asparaginase 11, the enzyme with antilymphoma activity, has a higher substrate affinity than asparaginase I. Both enzymes are saturated at 10 mM asparagine; but a t 0.1 mM asparagine, asparaginase I1 is almost entirely responsible for the observed hydrolysis. The EC-2 asparaginase of E. coli B also has a higher substrate affinity than EC-1 and presumably corresponds to asparaginase I1 of E. coli K-12. In E. coli K-12, asparaginase I1 appears to be located in the periplasmic space between the bacterial plasma membrane and cell envelope (67). Little information is available as yet on optimal conditions for enzyme production. Roberts et a2. (43) compared several media but did not determine how much of the total activity resulted from the EC-2 asparaginase. Schwartz et al. (66) reported that maximal yields of asparaginase I1 are produced under “highly anaerobic” conditions. I n the most thorough study to date of growth conditions, Bilimoria (68) found that gentle aeration (shake cultures) are most suitable for good cell growth and high enzyme content. The situation with respect to the number of different E. coli asparaginases is complicated in one sense, but clarified in another, by the recent report of the Bayer group (65) that homogeneous crystalline 66. J. H. Schwartz, J. Y. Reeves, and J. D. Broome, Proc. Natl. Acad. Sci. 56, 1516 (1966). 67. H. Cedar and J. H. Schwarts. J. Bid. Chem. 242, 3753 (1967). 68. M. H. Bilirnoria. Appl. Microbial. 18, 1025 (1969).

U. S.

110

JOHN C. WRISTON,

JR.

asparaginases from two different strains of E . coli, while being apparently identical in a number of ways, differ with respect to isoelectric point and with respect to the half-life in the circulation. Asparaginase A, in the terminology of the Bayer group, isolated from E . coli ATCC 9637, has an isoelectric point of 5.0, while asparaginase B, from E . coli ATCC 11303, has a value of 4.8. This observation, attributed to different proportions of isoenzymes in the two asparaginases, helps to explain certain apparent discrepancies in reports of the properties of asparaginase from different laboratories. The subunit structure of asparaginase is discussed in a later section. 2. Substrate Specificity and Effect of Inhibitors

Escherichia coli B asparaginase consistently shows about 2-4% of the activity toward L-glutamine that it does toward L-asparagine (39,69) and has about 5% as much activity toward D-asparagine ( 4 1 ) . This is in contrast to GPS asparaginase, which has no effect on L-glutamine but does hydrolyze D-asparagine to a limited extent ( 2 ) .Campbell and Mashburn (41) have presented evidence strongly suggesting that the hydrolysis of all three compounds occurs a t the same active site in E . coli asparaginase. Product inhibition occurs with ammonia a t pH 8.5, although not a t pH 7.4 or 5.0, but neither hydrolysis nor inhibition is found with L-aspartic acid, D-aspartic acid, L-glutamic acid, or D-glutamic acid ( 4 1 ) .Escherichia coli asparaginase will not catalyze the cleavage of p-aspartyl glucosylamine compounds and has not been observed to release ammonia from several proteins known to contain asparaginyl residues (70). Escherichia coli B asparaginase has no free sulfhydryl groups and is not sensitive to such reagents as p-mercuribenzoate, N-ethylmaleimide, M (71) 1. It is inhibited specifically by 5and iodoacetic acid [ 1 x diazo-4-oxo-~-norvaline (DONV) (41, 72, 73) as is guinea pig serum asparaginase, but not by the corresponding glutamine analog, DON, which inhibits glutaminase ( 7 4 ) . Handschumacher has shown (72,73) that DONV acts both as an alternate substrate and as an inhibitor. 69. J. H. Kim, E. A . Boyse, L. J. Old, and H . A . Campbell, Biochim. B i o p h y ~ . Acta 158, 476 (1968). 70. L. T. Mashburn, personal communication (1969). 71. H. A . Whelan and J. C. Wriston, Jr., unpublished results (1969). 72. R. E. Handschumacher, C. J. Bates, P. K. Chang, A . T. Andrew. and (;. A . Fischer, Science 161, 62 (1968). 73. R. C. Jackson, D. A. Cooney, and R. E. Handschumacher, Fedemtion P w c . . 28, 601 (1969); R. C. Jackson and R. E. Handschumacher, Biochemistry 9, 3585 (1970). 74. B. Levenberg, I . Melnick. and J. M. Buchanan,

J. Biol. Chem. 225,

163 (1957).

5.

L-ASPARAGINASE

111

In experiments with a relatively high ratio of asparaginase to substrate, DONV is catalytically decomposed, with the evolution of nitrogen and the formation of 5-hydroxy-4-keto-~-norvaline. This reaction can be blocked by L-asparagine but not by L-glutamine. I n experiments with saturating amounts of DONV, however, a slower, irreversible reaction also occurs involving the binding of the analog to the enzyme. Covalent binding has been demonstrated by using 5-14C-DONV, and since the modified enzyme is no longer active toward asparagine, binding may well have occurred at the active site of asparaginase. The hydrolysis of asparagine is strongly inhibited by dimethylsulfoxide (DMSO) in concentrations higher than 5 M , but activity can be regained by rapid dilution. Catalytic decomposition of DONV is also inhibited in 5 M DMSO, but covalent binding appears to proceed at an undiminished rate. The D-isomer of DONV is inactive both as substrate and inactivator of the enzyme. Asparaginase also catalyzes the slow hydrolysis of another asparagine analog, P-cyanoalanine, to L-aspartic acid, and since pcyanoalanine can competitively inhibit the decomposition of DONV it would appear that the nitrilase activity occurs at the same active site. 3. Amino Acid Composition

The amino acid compositions of several E. coli asparaginase preparations are presented in Table 11. The first such report (61) assumed a subunit molecular weight of approximately 22,000, based partly on the apparent molecular weight of subunits in dissociating solvents, and partly on the necessity of achieving integral values for the number of residues of certain amino acids such as cystine, present to only a limited extent in asparaginase. The amino acid composition data in Table 11, however, are presented as residues per subunit of molecular weight of approximately 33,000 because of the growing body of evidence indicating that there are 4 subunits, not 6, per mole of enzyme. Four subunits a t 33,000 each are as -compatible with the generally agreed upon monomer molecular weight of about 130,000 as are 6 a t 22,000. The evidence for the $-subunit model is discussed in a later section. I t may be seen from Table I1 that there is good agreement in general between the reports from the various laboratories. It is worth noting, as Arens et al. (65) have done, that about 50% of the molecule is composed of only five amino acids: aspartic acid, threonine, alanine, valine, and glycine. Whether the differences that are reported result from experimental variation in the analyses, real differences between strains, or different proportions of isoenzymes in the enzymes examined remains to be seen. As mentioned in another section, the Bayer group (65) found

112

J O H N C. WRISTON, J R .

TABLE I1 AMINO ACID COMPOSIT~ON OF E . coli ASPARAGINASES~ AmLiiio acid

Squibb [Ref. (GI)]

ASP Thr Ser Glu Pro GlY Ala Val CYS Met Ile Leu TYr Phe LYS His -4% T rP NH,

45 30 15 20 11 27 29 30 2 6 11 22 10 8 20 3 S 1 -

50 31 13 19 11 29 34 33 1 3 12 23 10 8 22 3 S 1 50

46 30 15 19 12 26 30 31 2 5 12 21 10 S 20 3 7 2 27

29s

311

299

Total residues a

Lilly [Ref. (SO)]

Bayer [Ref. (SS)]

As residues, per 33,000 molecular-weight subunit.

no differences in amino acid composition between their asparaginases A and B. One explanation that has been offered for the fact that the Bayer asparaginases A and B differ with respect to isoelectric point without showing any differences in amino acid composition is that there are differences in the number of amide groups in the two enzymes. Arens et al. (65) have reported an experimental value of approximately 27 moles of ammonia per 33,000 molecular weight subunit for asparaginase A but have not reported the corresponding figure for asparaginase B. This is in good agreement with a value obtained by Greenquist and Wriston (75) for the amide content of asparaginase from E. coli B by the carboxyl modification method of Hoare and Koshland (76). It is difficult to draw an accurate comparison with GPS asparaginase, the only other asparaginase whose amino acid composition has been 75. A. C. Greenquist and J. C. Wriston, Jr., Federation Proc. 29, 882 (1970). 76. D. G. Hoare and D. E. Koshland, Jr., J . Biol. Chem. 242, 2447 (1967).

5.

L-ASPARAGINASE

113

reported, because its molecular weight appears to be a little higher than the E. coli enzyme and because there is as yet no evidence on subunit structure. On the assumption that the 138,000 molecular weight GPS asparaginase molecule consists of 4 subunits of about 35,000 molecular weight each, however, there appear to be inajor differences in composition between the two enzymes. The GPS enzyme has only about half as much aspartic acid, for example, but twice as much proline, and it is also strikingly low in tyrosine. 4. Structure

Analytical ultracentrifugation at 0.1% enzyme concentration in 0.2 M potassium phosphate and 1% sucrose at pH 7.25 gave an apparent molecular weight of approximately 139,000. I n a slightly different solvent system, 0.02 M sodium phosphate and 0.2 M NaCl at pH 6.85, a value of 125,000 was obtained (61, 77). At 1% asparaginase, however, a value about twice as high was obtained, and a t low protein concentrations (0.02%) the apparent molecular weight was 64,000. Frank and Veros (78) also obtained a molecular weight value of 130,000 by sedimentation equilibrium, but they did not confirm the apparent dissociation upon simple dilution seen by Kirschbaum et al. (77). The Bayer group (65) obtained a value of 127,000 for the molecular weight of the monomer. They also note a second species with a molecular weight of about 240,000 in higher protein concentrations and observed partial dissociation in aqueous solution by dilution to below 0.1%. Evidence for the existence of subunits has also been obtained by dedetermining the Jdimeritation coefficiciits i n several solvent systems (77). The s , O , , ~ value at 10 mg/ml is 8 . 6 s (molecular weight 255,000). At a concentration of 1 mg/ml, however, a second species of sedimenting material appears with an s&,, value of 5 . 6 s (molecular weight 125,000). When the enzyme concentration is lowered to about 0.4 mg/ml, only a trace of the 8 . 6 s peak remains, and a third species appears with an s,",,, value of 4.0s (molecular weight of 65,000). Increasing the NaCl concentration from 0.1 M to 3 M with protein concentrations in the ra!igc of 5-10 mg/ml also causes similar changes in sedimentation coefficients, and although the Sedimentation coefficient of E. coli asparaginase remains constant at 8.6 S from pH 4 to 10.6, below pH 4 the 77. J . Kirschbaum. J. C. Wriston, Jr.. and 0. T. Ratych, Bzochim. Biophys. Actn 194, 161 (1969). 78. B. H. Frank and A. J . Veros, Federntion Proc. 28, 728 (1969); B. H. Frank, A . H. Pekar. A. J. Veros. :inti P. P. K. Ho. J. Biol. Chem. 245, 3716 (1970).

114


enzyme again begins to dissociate. I n all these cases, only the 8.6s form of asparaginase is observed several hours later. Rich e t al. (79) have carried out a temperature jump relaxation study of E . coli asparaginase and reported that the finding of multiple relaxation times is qualitatively consistent with the suggested subunit structure. The apparent molecular weight of asparaginase in 8 M urea, 5 ill guanidinium chloride, or 9 M formamide was first reported as being 19,000-24,OOO, with an s , ~ ,of~ 1.5 to 1.8 S (77), demonstrating the presence of subunits. Arens et al. (65) and Frank and Veros (78) have confirmed the value of the sedimentation coefficient of the subunits obtainctl in 8 M urea, but the latter authors reported an apparent molecular weight of approximately 32,500 instead of 19,000-24,000, and their value is almost certainly the correct one. Asparaginase preparations thought to be homogeneous or nearly so usually show one or more faint bands in addition to the principal one when subjected to disc electrophoresis. Gel slicing experiments show asparaginase activity in both the major and minor peaks in such experiments, and it appears that the minor peaks result from aggregates of the 130,000 molecular weight monomer species. Ho and Milikin (80) have shown that the multiple forms seen on disc electrophoresis represent a size isomer family of proteins by plotting the log of protein mobility vs. acrylamide gel concentration. Size isomeric proteins give a family of nonparallel lines extrapolating to a common point in the vicinity of 2% gel concentration, whereas charge isomers would give a family of parallel lines (81). The former kind of plot was obtained with the crystalline Lilly E . coli asparaginase. Whelan and Wriston (61) reported a value of 4.85 for the isoelectric point of their preparation of E . coli B, and a value of 4.95 was reported by Campbell and Mashburn (41). Roberts et al. (43) found a completely different value of 4.35. Arens et al. (65) noted that the electrophoretic mobilities of asparaginases A and B (from E. coli strains ATCC 9637 and 11303) were different, the isolectric points being 5.0 and 4.8 for A and B, respectively. They noted in addition, however, that the single proteill band obtained in these cellulose acetate electrophoresis experiments was asymmetric when examined in a densitometer. When they carried out prolonged electrophoresis, the apparently single band separated into several equally spaced bands in each case. The principal zones of the asparaginases were labelcd A, and B,, with the remaining bands being numbered consecutively in order of increasing electronegativity. A r m 79. L. D. Rich, E. M. Eyring, and J. Roberts, personal communication (1969). 80. P. P. K. Ho :ind E. 13. Milikin. B i t ~ l i i mR. i o p k y s . Acln 206, 196 (1970). 81. J. L. Hedrick and A. Smith, Arch. Biochem. Biophys. 126, 155 (1968).

5.

L-ASPARAGINASE

115

et al. showed that the zones A, and B, carry the same net charge by performing electrophoresis with a mixture of asparaginases A and B. However, if the mixture was incubated briefly in the presence of 6 M urea and then subjected t o electrophoresis, a completely different pattern was obtained with a Gaussian distribution of isoenzymes. Both asparaginases A and B have the same specific activity, the same broad pH-activity profile with a maximum a t pH 7.2, the same tendency to hydrolyze L-glutamine, and the same apparent amino acid composition. They have the same apparent molecular weight, and the N-terminal amino acid sequences of asparaginases A and B are identical for the first 14 residues. Aside from the differences i n electrophoretic mobility, the only other difference yet established is the longer half-life time of asparaginase B, the more acidic of the two enzymes. Arens et al. (65) have concluded that both enzyme preparations consist of a series of isoenzymes with different isoelectric points though with the same specific activities, with asparaginase B containing more of the acidic isoenzymes than asparaginase A. Although not yet established, it would appear reasonable to assume that other E . coli asparaginases will be constructed on the same principle. It was first suggested tentatively (61) that there were 6 subunits in asparaginase from E . coli B, primarily because of the apparent molecular weight values (19,00&24,000) of the species obtained in 8 M urea or 5 M guanidine. However, it is now becoming reasonably certain, as a result of work from several laboratories, that there are in fact 4 subunits per mole of asparaginase. The preliminary results obtained by Greenquist and Wriston (75) (working with E . coli B asparaginase provided by Squibb) provide support for the 4-subunit model and also suggest that the subunits are identical; but one cannot, of course, have isoenzymes unless there are a t least two kinds of subunits. If the finding of isoeiizymes by Arens et al. (65) in asparaginases A and B proves to be characteristic of all the E . coli asparaginases, only further work will be able to resolve the question of the apparent similarity of the E . coli B subunits. The structure work of the Delaware group (75) may be summarized as follows. Treatment with Sanger’s reagent yields only DNP-Leu, approximately 4 moles per mole of enzyme. Tyrosine is the only free aniino acid liberated in significant quantity upon hydrazinolysis, to the extent of 3.3 moles per mole of enzyme. In studies with carboxypeptidase A , Tyr is released very rapidly from S-aminoethylated enzyme (approximately 4 moles per unit of enzyme) followed by Gln, Ile, Phe, and Asn. The order of these 4 aniino acids is not yet established unequivocally, but there are approximately 4 moles of each per mole of asparaginase. Finally, fingerprints of tryptic digests of S-aminoethylated asparaginase

116

JOHN C. WRISTON, J R .

show 32-36 ninhydrin-positive spots, 10 of which are also Sakaguchi positive. On the basis of 20 Lys, 8 Arg, and 2 Cys residues per minimum molecular weight unit of about 33,000, these results are about what would be expected if the subunits were identical. It is true, however, that there appear to be a few more pepticle spots on the fingerprint map than would theoretically be formed, allowing for the possibility of a small number of differences between the subunits. Arens et a2. (65) also reported finding only N-terminal Leu in both asparaginases A and B. In addition, using the Edman procedure, they have established the sequence of the first 14 amino acids from the Nterminal end (shown below) still without encountering a difference between the two H-Leu~Pro~Asn~Ile.Thr~Ile~Leu~Ala~Thr~Gly. (Gly.Val.Ile.Ala)1

2

3

4

5

6

7

8

9

10

11 1 2 1 3 1 4

asparaginases, or without finding any position that appears to be occupied by more than one amino acid, as might he the case with subunits of different structure. Wagner et al. (82) have used nitrous acid to prepare a partially deaminated asparaginase in which the a-amino group of the N-terminal Leu and one of the c-amino groups of Lys are removed. This modified asparaginase can be crystallized and retain full activity. It is more acidic than the native enzyme and has a longer half-life time.

V. Other Asparaginares

The propertieb of several other asparaginases are coniparcd with those of the better known GPS and E . coli cnzymes in Table 111. Three of these enzymes (from Sermtia mnrcesceiis, Erwinin cnrotovorn, and Erwinia aroideae) have antilymphoma activity, while the labt two ~~1~ do not. Thc Ser(from Bacillus coagulans and F i ~ t l i . 1 tricinctzm) ratia enzyme has bceii purified extensively [ apl)roxiinntcly 600-foltl (35)] but not to homogeneity. Hcinemaiin and Howard (85) have recently studied fermentation conditions for thc production of thc Serratia enzyme. The Eminin ctrrotovora asparagin:i>e has I)eeii crystallized by the Portori group (38,84) and has becn found to have an isoelectric point that is strikingly higher than those of the other known 82. 0. W:igncr, E. Irion. A . ;\rcsns. nntl K. B:iric~.Hioclietn. Hiripliya. 11'c.s. C o n t ~ t r / r . 37, 383 (1969). 83. 13. Heincmann and A . J. H o ~ w r d A , / ~ p lMicrobial. . 18, 550 (106'3). 84. A. C. T. North. H. E. Wxlc. and K . A . Cnmmack. Nolitre 224, 59.4 (I%!)).

5.

L-ASPARAGINASE

117

asparaginases (85). Extracts of Erwinia carotovora also have specific enzymic activities several times higher than either E . coli or Serratia extracts, and the specific activity of the crystalline enzyme appears to be about twice as high as that of E . coli asparaginase (700 IU/mg as compared to 280-400 IU/mg). Partial purification of an asparaginase from Erwinia aroideae (37) has also been described. No reports of clinical trials with any of these three enzymes have yet appeared. The asparaginase from Bacillus coagulnns, first described by Manning and Campbell (311 , has been purified about 300-fold but not to homogeneity (86). It does not have antilymphoma activity and differs from those asparaginases that do in sever:il ways. Its pH optimum is a good deal higher, for example; it has a lower molecular weight, and is very rapidly removed from the circulation of thc mouse, in which respect it resembles yeast aaparaginase. I t is also rapidly and irreversibly denatured by 8 M urea and is inhibited (reversibly) by p-mercuribenzoate ((38% inhibition at 1 X lO-'iM). The enzyme appears to bear a closer resemblance to one of the M . tuberculosis asparaginases described by Jayaram et al. (SO) than it does to any of the asparaginases possessing antilymphoma activity. I t has proved difficult to purify because of the small amounts present in cell extracts. The asparaginase from the mold Fusariuuz tricinctum, also without ailtilymphoma activity, has been purified to apparent homogeneity, although its amino acid composition has not yet been established (40). It is perhaps worth noting that it has been found to contain both galactosamine and glucosamine. VI. Physiological Properties

The mechanism by which L-asparaginase exerts an antitumor activity is not yet clear. The cells of susceptible tumors require asparagine, while normal cells and the cells of resistant tumors appear to be independent of an external source of this amino acid (18, 39, 56, 87-90). Scveral groups of workers have established a correlation between S5. 1.. T. M : i s I i l ~ i i ~:~i nnc I I.. M. 1,:incIin. i,c "licvvnt Rrsults in C:inc.rr Rrsenrc,li" (;riintlin:inn ; i n ( l H . I;. Orttgrn. rcls.). Vol. 33. 1). 48. Sliringcr-Vrrlag. Hcilin :ind Scs\v Toil;. 1070. 86. A . 8. I,nw, PIi.L). l'lit&. TJnivc,rsityof L)rl:iwiiic~. 1969. S7. 1'. A . Boysc, I,. J . Old, H. -4. C:iniph~ll,and I,. T. M:isliburn. J . E x p t l . A l e ( / . 125, 17 (1967). 88. I,. J. Old. E. A . lloysr. H . A . C~inilibrll. It. S.Ikodey, J . Fidler, and J. D. Ti~llc~i. ('(ltirer 20, 1066 (1967). 59. LV. C. Doloivy. D. Henson. J. Cornet. and H . Sellin, Cn/rcer 19, 1813 (1966). 90. R. A . Keriman and T. A . McCoy. Sczenre 124, 114 (1956). (11;.

118


TABLE I11

Property Molecular wt

Guinea pig serum [Ref. (6611

E. coli B (Squibb) [Ref. (6l)l

138,O0Oa 133, OOob

125,000139, OOW 106, OOob

Nessler GLDHd pH stat Isoelectric point

7 . 2 X 1W6 3.6-4.5e

1.25 X 1 W 5.25 X 6 . 0 X 1W6 4.85,; 4.979

pH optimum

7.5-8.5

No

8 (broad maximum) 4%

+

+

Kill

Hydrolyzes glutamine Antitumor activity Urea denaturation Half-life time

?

11-19 hrh

E. coli B (Lilly) [Ref. (6.211

E. coli (Bayer) [Ref. (66)]

130,000”

127, OOW 120, ooob

-

-

5.39, 5.5n

1%

Bayer A, 5 .Of Bayer B, 4.81 Bayer A, 5 . 3 , 5.59 Bayer X, 4.639 7 . 2 (broad maximum) 34%

+

+

-

Iteversi ble

Reversible

Reversible

4 . 0 hri

2 . 9 hr‘

Bayer A, 2 . 8 hr; Bayer X, 7 . 3 hri

Analytical ultracentrifuge. Gel filtration. c Sucrose density gradient. d Coupled assay with glutamic dehydrogenase, E. D. Mooz and J. C. Wriston, unpublished data (1970).

resistance to asparaginase and the presence of the asparagine synthetase, on the one hand, and asparaginase sensitivity and an absence of the capacity to synthesize asparagine, on the other. The low levels of asparagine synthetase in several normal tissues are puzzling in this connection, however (91-98). 91. 92. 93. 94.

J. D. Uroonie, J . E z p t l . M e d . 127, 1036 (1968). J. D. Broome and J. H. Schwartz, Biochim. Biophys. Acta 138, 637 (1967). M. K.Piitterson and G. Orr, Biochem. Biophys. Res. Cornmiin. 26, 228 (1967). M. D. Prager and N. Bachynsky. Biochem. Biophys. Res. Commun. 31, 43

(196s).

95. M. D. Prager and N. Bachynsky, Arch. Biochem. Biophys. 127, 645 (1968). 96. 13. Horowitz, B. K. Madras, A. Meister, L. J. Old. E. A. Boyse, and E. Stockert, Scieltce 160, 533 (1968). 97. M. D. Prager, P. C. Peters, J. 0. Jones, and I. Derr, Notitre 221, 1064 (1969).

5.

119

L-ASPARAGINASE

PROPERTIES OF SEVERAL ASPARAGINASES

E. coli (Kyowa Hakko)

E. E. S . marcescens carolovora aroidcae [Ref. (%)I [Ref. (S8,84)] [Ref. (4.213

141 ,OOOa

150,OOOE

-

1 x 10-4 1 . 2 x 10-6

-

7.2 X 4.7

5.029

8.0 1

5.11 5.85,

6.9 (broad maximum) 4%

+

+

Reversible

lteversi ble

4 . 2 hr'

?

-

128,000145,000 3

x

10-3

B . coagutans F. lricinctum [Ref. (86)j [Ref. (40,] 85,OOOE

4.7 x 10-8

5.2 x 10-4 4 . 2 x 10-4 5.181

8.79,8.9"

-

-

7.5

8.5-9.5

7.5-8.7

2

No

No

No

No

15%'

-

160,000171 ,OO@

+

+

2

2

Irreversible

Reversible

4.1 hr'

>

p H 9 ) ( 5 , 19). It should be noted that the catalytic reaction by the enzyme cannot be stopped by simply heating the reaction mixture to loo", and even a t 100" an abnormal reaction might occur (K. Satoh, Y. Inoue, and F. Egami, unpublished). Heparin, which inhibits RNase A, does not inhibit RNase TI, nor does a natural RNase A inhibitor from rat liver (do), or Aspergillus oryzae nuclease inhibitor ( 2 1 ) . 16. S. Minato, T . Tagawa, and K. Nakanishi, Ann. Sankyo Res. Lab. 15, 122 (1963). 17. S. Minato. T. Tagawa, and K. Nakanishi, J . Biochem. ( T o k y o ) 59, 443 (1966). 18. S. Irie, T. Itoh, T. Ueda, and F. Egami, J . Biochem. ( T o k y o ) 68, 163 (1970). 19. S.Iida and T. Ooi, Biochemistry 8, 3897 (1969). 20. K. Shortman, BBA 55, 88 (1962). 21. T. Uozumi, G. Tamura, and K. Arima, Agr. Bid. Chem. ( T o k y o ) 33, 636 (1969).

214

T. UCHIDA AND F. EGAMI

TABLE V BASESPECIFICITY OF RNase TI

R

R

SIC( 1 (Refs. I X , 2 7 )

A (-)

(Ref. 23)

H G l y o x a l G (t) (Refs. 26.31

R

N - 1 M e t h y l G (*) (Refs. 2 8 , 2 9 )

I

R

R

CMC-G ( 1 (Ref. 30)

9.

215

MICROBIAL RIBONUCLEASES

3. Specificity and Mode of Action Ribonuclease TImay be regarded as a guanyloribonuclease [EC 2. 7.7.26, ribonucleate guaninenucleotido-2’-transferase (cyclizing) ] (22, 23). It splits the internucleotide bonds between 3’-guanylic acid groups and the 5’-hydroxyl groups of the adjacent nucleotides with the intermediary formation of guanosine 2’,3’-cyclic phosphate. The first step, transphosphorylation, is reversible. The second step, hydrolysis of the cyclic phosphate to produce 3’-guanylate, is practically irreversible. The base specificity has been extensively investigated by the 3‘-terminal analysis of digestion products of RNA and polyribonucleotides and by the studies on the susceptibility of dinucleoside monophosphates or nucleoside 2’,3’-cyclic phosphates to the enzyme. The results are summarized in Table V ( 2 4 3 7 ) . From the base specificity study, the esTABLE V (Continued)

I

I R

N-2 Dimethyl G

(*)

(Refs. ? 8 , 2 3 , 3 3 )

I N P - G (-) (Ref. 3 G )

R

P -Sulfophenylazo G (T. Ito, T. Uchida, and F. Egami, unpublished) ( 6 )

8 - A m G (+ +)

(Ref. 37)

22. K. Sat0 and F. Egami, C o m p t . Rend. Soc. B w l . 151, 1792 (1957). 23. K. Sato-Asano, J. Biochem. (Tokyo) 46, 31 (1959). 24. P. R. Whitfield and H. Witzel, BBA 72, 338 (1963). 25. K. Sato-Asano and Y. Fujii, J. Biochem. (Tokyo) 47, 608 (1960). 26. T. Uchida and F. Egami, J . Biochem. (Tokyo) 57, 742 (1965). 27. S. Irie. T. Uchida, and F. Egami, BBA 209, 289 (1970). 28. M. Staehelin, BBA 87, 493 (1964). 29. R. W. Holley, J. Apgar, G. A. Everett, J. T. Madison, M. Marquisee, S. H. Merrill, J. R. Penswick, and A. Zamir, Science 147, 1462 (1965). 30. T. Uchida, T. Arima, and F. Egami, J . Biochem. (Tokyo) 67, 91 (1970).

216


sential requirements for the preferred substrates of RNase T, are the keto group a t the 6 position and the trivalent nitrogen a t the 7 position of purine base (11). The presence of a proton a t N-1, as in guanine, further increases the susceptibility. Thus, these groups may be regarded as the specific binding sites of the purine base with the enzyme. This consideration may explain the apparent resistance of doublestranded RNA, such as RNA from cytoplasmic polyhedrosis virus (38, S9), and of polyribonucleotides highly rich in guanylyl residues, such as RNase A resistant core of RNA (40),since these sites participate in the formation of double-stranded structure and in the association of guanylyl residues. The optimum pH values of RNase TI for the hydrolysis of guanosine cyclic phosphate and xanthosine cyclic phosphate are pH 7.2 and 4.5, respectively. The xanthosine cyclic phosphate may have the lactam form (-NH-CO-) susceptible to the enzyme only in the acidic medium (18). (1) ( 6 ) The resistance of S-substituted analogs, such as 6-thioguanylylcytidine, may result from the lower electronegativity of sulfur atom than that of oxygen and/or the longer atomic distance of C=S than that of C=O (27). The resistance of N-2 p-sulfophenylazoguanylyl bonds (T. Ito, T. Uchida, and F. Egami, unpublished) may result from the steric hindrance by the large substituent. A considerable effect by the neighboring nucleoside on the splitting rate of 3’-guanylyl phosphodiester bonds was observed by Whitfeld and Witzel (Z4),and Irie ( 4 1 ) , as shown in Table VI and VII. The K , values for GpA, GpC, GpG, and GpU are of similar magnitude, but the V,,, values decrease in the order GpC > GpG > GpA > GpU (15.1 : 3.5:3.3:1) (41). This is the reverse order of the reaction rate catalyzed by OH- and is a promising observation for the analysis of the reaction

31. N. W. Y. Ho and P. T. Gilham, Biochemistw 6, 3632 (1967). 32. M. Hiramaru, J. Sokawa, T. Uchida, and F. Egami, Seikagaku 38, 662 (1966). 33. U. L. RajBhandary, A. Stuart, and S. H. Chang, JBC 243, 584 (1968). 34. C. B. Reese and J. E. Sulston, BBA 149, 293 (1967). 35. J. Hashimoto, T. Uchida. and F. Egami, BBA 199, 535 (1970). 36. M. Azegami and K. Iwai, J . Biochem. ( T o k y o ) 55, 346 (1964). 37. D. H . Levin, BBA 61, 75 (1962). 38. K. Miura, I. Fujii, T. Sakaki, S. Kawase, and J. Fuke, Proc. 7 t h Intern. Congr. Biochem., T o k y o , 1967, Abst. IV, p. 618, B-5.Sci. Council Japan, Tokyo, 1968. 39. M. A. Billeter, C. Weissmann, and R. C. Warner, J M B 17, 145 (1966). 40. K. Itagaki, Y. Kuriyama, Y. Shiobara, H. Hayashi, T. Yamagata, and F. Egami, Seikagaku, 37, 217 (1965). 41. M. Irie, J . Biochem. ( T o k y o ) 63, 649 (1968).

9.

217


TABLE VI RELATIVE RATESOF SPLITTING BY RNase Tp

4

Substrates

Relative rate

GPCP GPC GPA GPG GPU IPC XPC GlyoxalGpC G cyclic-p

1100 800 550 450 250 150 10 5 2

From Whitfeld and Witzel (94).

mechanism catalyzed by the enzyme and by OH- (T. Koike and Y. Inoue, unpublished). As mentioned in the beginning of this chapter, the base specificity of RNases is, in general, not absolute, but relative. Ribonuclease TI is not exceptional in this sense. In a longer incubation with higher concentrations of the enzyme (substrate-enzyme ratio: 40/1 w/w or less) compared with the usual experimental condition (substrate-enzyme ratio : 100/1 or more), nonbase-specific cleavage may occur even with an enzyme preparation of guaranteed purity. The order of the susceptibility to the enzyme is G>> A > U > C (T. Uchida and F. Egami, unpublished). The observation by Irie (42) of cleavage of poly A, poly C, and poly U and by Michelson and Money of poly 7-methylguanosine digestion (43) may be nonbase-specific cleavages, although a slight contamination by TABLE VII Km AND Vmax VALUESAT pH 7.5

OF

RNase TIo VmEZ

Substrate GPC GPG GPA GPU

(mole/min/O.27 unit enzyme)

Km 0.445 X 0.291 X 0.268 x 0.238X

lo-' lo-' lo-' lo-'

6.65 x 1.54x 1.47 x 0.44 x

10-4 10-4

10-4 10-4

From Irie (41).

42. M.h i e , J. Biochem. ( T o k u o ) 58, 599 (1965). 43. A. M. Michelson and C. Money, BBA 166, 294 (1968).

218


other RNases such as RNase T, in their enzyme preparations could not be excluded. The phosphodiester bond has been thought to be specific to the internucleotide 3’,5’-phosphodiester bonds (44-46). However, convincing evidence has been presented quite recently (Y. Inoue, unpublished) that Gp (3‘,5’) -N, but not Gp (2’,5’)-N, is split. A curious observation by Podder and Tinoco (47) that G-(2’,5’)-G bond was synthesized by RNase T, from G-cyclic-p led Egami and Inoue (unpublished) to reinvestigate the phosphotransferase activity a t various temperatures. At loo”, unlike at 36”, G- (2’,5’)-G,,-, is split to produce G,,,, and G-(3’,5’)-G3,-, is attacked in quite a different way a t 100” than it is a t 36”. This may result from the altered action of partly heat-denatured RNase T,. Native RNase TIis specific, however, to the internucleotide 3‘,5‘-phosphodiester bonds a t normal temperature. Unlike guanosine 2’,3’-cyclic phosphate, guanosine 3’,5’-cyclic phosphate is quite resistant to the enzyme (S. Sato, T. Uchida, and F. Egami, unpublished). The sugar specificity of RNase T, appears to require a 2‘-hydroxyl group for the substrate because DNA is not attacked by RNase TI. This is consistent with the intermediary formation of 2‘,3’-cyclic phosphate and also with the finding that a’-O-methylated guanylyl bonds in tRNA is resistant to the enzyme ( 4 8 ) . Holy and Sorm (49) found that RNase T, did not attack L-guanosine 2’,3’-cyclic phosphate and L-inosine 2‘,3‘cyclic phosphate. They found further that RNase T, split 9- (a-L-lyxofuranosyl) -hypoxanthine 2’,3’-cyclic phosphate but not the D-lyxofuranose derivative, and they concluded that the substrate molecule was fixed a t least to three regions of RNase TI( 5 0 ) . 4. Structure and Function

The amino acid sequence of RNase T, has been elucidated by K. Takahashi. It consists of a single polypeptide chain of 104 amino acid residues cross-linked by two disulfide bonds, essential for the maintenance of the enzymically active structure, as shown in Fig. 2 (51). The reductive cleavage of the bonds destroys the enzyme’s activity, but 44. Y. Kuriyama, J. Koyama, and F. Egami, Seikagaku 36, 135 (1964).

45. M. N. Lipsett, JBC 239, 1250 (1964). 46. H. Nishimura, T. Sekiya, and T. Ukita, BBA 174, 653 (1969). 47. S. K. Podder and I. Tinoco, BBRC 34,569 (1969). 48. H. G . Zachau, D. Dutting, and H. Feldmann, 2. Physiol. Chem. 347,212 (1966). 49. A. Holf and F. Sorm, Collection Czech. Chem. Commun. 34, 3383 (1969). 50. A. Holg and F. Sorm, Collection Czech. Chem. Commun. 34, 3523 (1969). 51. K. Takahashi, JBC 240, PC4117 (1965).

9. I

tl

219

MICROBIAL RI BON U CLEASES

Ald

(‘ys I

=

I0 15 3, 25 .I0 CY, Tyr Scr Srr Ser A * p Val Scr Tlir A h (;In A h AIL (;ly Tyr G > C . (3) No small oligonucleotides such as di- or trinucleotides with 3’terminal Ap are produced throughout the process of digestion, but those with 3’-terminal Gp or Up are produced in the course of digestion. (4) Adenosine 2‘,3‘-cyclic phosphate is scarcely accumulated, though other nucleoside 2’,3’-cyclic phosphates are accumulated as intermediates. This result suggests that the action of RNase TIon RNA is owing to the cooperation of an adenylic acid specific endonuclease activity and a nonspecific exonuclease activity releasing mononucleotides from the 3‘ terminal. The mode of action on poly U is similar to that on RNA. A remarkable increase of 3’ terminus a t an initial stage (about 25% of total residues), subsequent accumulation of UpUp or U > p, and rapid appearance of Up after a short lag time are observed. This shows that RNase T, clearly behaves as an endonuclease for poly U as for RNA. Therefore, it should be interpreted that the apparent initial endo85. T. Uchida and F. Egami, J . Bbchem. (Tokyo) 61,44 (1987).

226


nuclease activity on the digestion of RNA with RNase T, is not specific but preferential for adenylic acid. The susceptibility to RNase T, depends on the nature of various RNA's and homopolymers (85).Poly A is fairly resistant to RNase T, in spite of the preferential cleavage of adenylic acid linkages in RNA, whereas, unexpectedly, poly U is the most sensitive to RNase T, of the various homopolymers tested. Poly C is also resistant to digestion by RNase T,. I n the digestion of poly A or poly C, only 3'-mononucleotide is detected as a digestion product and a small increase of 3' terminus in the initial stage (about 4% of total residues) is observed; the initial endonuclease activity appears to be markedly reduced. The reason may be that poly A forms a rigid double-stranded helical structure in acidic medium, because poly U, which has no secondary structure a t 37", is easily digested. I n 40% methanol solution, where a part of its rigid double-stranded structure is destroyed, poly A is more sensitive to RNase T, than it is in water; and the mode of action of RNase T, on poly A in 40% methanol solution is very similar to that on RNA or poly U. Furthermore, the susceptibilities to RNase T, among various RNA's are in the order of commercial yeast RNA (low molecular weight) > high molecular weight yeast RNA > tRNA mixture containing more double-stranded structure than the other RNA's ( 8 5 ) .These observations also confirm the resistance of higher structure in substrate molecule to RNase T,. Ribonuclease T, has no absolute specificity, but 8-methyl ribofuranoside 2',3'-cyclic phosphate is not cleaved by RNase T, ( 8 5 ) , indicating that a t least a part of the base Structure is of significance for the action of RNase T,. The order of preference of RNase T, for various bonds in RNA is ApN > UpN > GpN > CpN ( 8 5 ) , so no general rule can be stated for the preference for purine or pyrimidine, or for keto or amino substituent. Recently, it was found that RNase T, gave complete cleavage of the phosphodiester bonds of 4-thiouridylate in tRNA (86) and of 6-thioguanylyl- (3'-5') -cytidine (27). This shows that the substitution of sulfur a t the 4 position of pyrimidine base or a t the 6 position of purine base does not effect the action of RNase T,. Furthermore, the phosphodiester bonds of the nucleotides with a large substituent a t the 6 position of purine base such as N6-isopentenyladenosine (87), 2-methyl thi0-W- (Az86. M. Saneyoshi, F. Harada, and S. Nishimura, BBA 190, 264 (1969). 87. S. Hashimoto, M. Miyazaki, and S. Takemura, J . Biochem. ( T o k y o ) 65, 659 (1969).

9.


227

isopentenyl) adenosine (88),or N-9- (P-~-ribofuranosyl) purin-6-ylcarbamoyl threonine (89, 90) were also split completely by RNase T, to produce the corresponding 2’,3’-cyclic phosphates. The effect of the substituent a t N-7 in purine base on the susceptibility to RNase T,has been also studied, in comparison with a purine specific ribonuclease, such as RNase TI or U,, for which the importance of N-7 has been recognized. Ribonucleic acid methylated with dimethyl sulfate was completely digested with RNase T, to the acid-soluble products, in which mononucleotide (66%) and dinucleotide were detected on a paper chromatography ( 3 2 ) . From this result, the good susceptibility of 7methyl guanine residue to RNase T,, unlike its susceptibility to RNase TI, was expected and it has been clearly proved by the studies (86,91,92) on the primary structure of tRNA containing 7-methyl guanine residue. Recently, 7-dcazaadenylyl- (3’4’) -uridine (tubercidyl uridine) was found to be easily cleaved by RNase T,. Also, 7-deazaadenosine-2’,3’-cyclic phosphate was completely hydrolyzed by RNase T, as fast as adenosine 2’,3‘-cyclic phosphate, which is the most susceptible substrate to RNase T, (35). Therefore, it is concluded that N-7 in purine base has nothing to do with the susceptibility to RNase T,. The substituents a t the 5 or 6 position of the pyrimidine base have no relation to the susceptibility to RNase T,. Madison and Holley (93) have shown that RNase T, is able to split the phosphodiester bonds of pseudouridylic acid and 5,6-dihydrouridylic acid. Recently, Saneyoshi et al. (86) have found a new minor constituent, Vp, with 5-methyl cytidine 3’-phosphate and other mononucleotides on the two-dimensional paper chromatogram of RNase T, exhaustive digest of E . coli tRNAVa’; later the structure of V was proved to be uridine-5-oxyacetic acid (94). It has been reported that RNase T, is able to split easily the phosphodiester bond of I-methyl adenylic acid in mctliylated RNA ($2) and 1methyl adenylyl- (3’-5’)-uridine (30).However, the Ap-N-oxide phosphodiester bonds in RNA-N-oxide, prepared by the action of perphthalate on RNA, were found to be quite resistant to RNase T a ( 8 5 ) .The phos88. F. Harada, H. J. Gross, F. Kimura, S. H. Chang, S. Nishimura, and U. L. H:ijBhandary, BBRC 33, 299 (1968). 89. H. Ishikura, Y. Yamada, K. Murao, M. Saneyoshi, and S. Nishimura, BBRC 37,990 (1969). 90. S. Takemura, M. Murakami, and M. Miyazaki, J. Biochem. (Tokyo) 65, 553 ( 1969). 91. T. Seno, M. Kobayashi, and S. Nishimura, BBA 169, 80 (1968). 92. S. Cory and K. A. Marcker, European J. Biochem. 12, 177 (1970). 93. J. T. Madison and R. W. Holley, BBRC 18, 153 (1965). 94. K. Murao, M. Saneyoshi, F. Harada, and S. Nishimura, BBRC 38, 657 (1970).

228


phodiester bonds of glyoxal guanylic acid, blocked with glyoxal to link between N-1 and 2-amino group of guanylyl residues, was also found to be fairly resistant to RNase T, and to remain mostly in the 2’,3’-cyclic phosphate form after the digestion of glyoxalated RNA (SO).According to Tada, the phosphodiester bonds of dimethyl guanylic acid and 1-methyl guanylic acid are fairly resistant to RNase T, ( 9 5 ) . Accordingly, these observations show that blocking a t N-1 of the base with a substituent group results in the increase of the resistance to RNase T2, and that N-1 in the base is of significance for the action of RNase T,. Hol? and Sorm (49, 50) observed that RNase Tz, like RNase TI, attacks 9- (/?-D-ribofuranosyl) and 9- (a-L-lyxofuranosyl) derivatives but not 9- (p-L-ribofuranosyl) and 9- (a-D-lyxofuranosyl) derivatives. Also, RNase T, is quite inactive on the phosphodiester bonds like RNase TI, of the nucleotide with 2’-O-methyl ribose, such as 2’-O-methyl guanylic acid (96) or 2’-O-methyl cytidylic acid (86). Thus, the action of RNase T, is in good accord with that of RNase T, and RNase A on sugar specificity, which may be a common property throughout all RNases, producing 3’-phosphate via 2’,3’-cyclic phosphate. Sat0 et al. (84) studied the action of RNase Tz on various nucleoside 2’,3’-cyclic phosphates and determined the K,,, and V,,, a t the optimum pH, as shown in Table IX. Although the K , value for A cyclic-p was TABLE IX MICHAELIS CONSTANTS, INHIBITOR CONSTANTS, AND MAXIMUM VELOCITY OF RNase TP Substrates and inhibitors

Km

A cyclic-p C cyclic-p G cyclic-p CJ cyclic-p

2.3 x 4.0 x 6.5 x 6.1 x

(M)

Ki (MI

V, (pmole/min/ pmole protein)

Cyclic Nucleotides

2’-AMP 2’-CMP 2’-GMP 3’-AMP 0

13.0 X 12.6 X 9.0 x 15.1 X

10-4 10-4 10-4 10-4

10s lo3 10s 1@

2’- or 3‘-Nucleotides 1 . 1 x 10-6 1 . 2 x 10-4 3 . 4 x 10-4 3 . 8 X 10-6

From Sato et al. (84).

95. M. Tada, Seikagaku 38, 662 (1966). 96. U. L. RajBhandary, R. D. Faulkner, and A. Stuart, JBC 243, 575 (1968).

9. MICROBIAL

229

RIBONUCLEASES

a little smaller than that for others, no remarkable difference was observed between the Km and V,,,,, values with adenosine 2',3'-cyclic phosphate and those with others. This result may reflect lack of base specificity of RNase T,. The apparent preference for adenylic acid bonds observed in RNA digestion might result from the base-specific affinity of RNase T, to adenylate. The findings that the Ki values for 2'- or 3'adenylate toward the hydrolysis of uridine 2',3'-cyclic phosphate are much smaller than those for other nucleotides (Table I X ) , and that the adenylates protect most strongly from bromoacetate inactivation of RNase T,, support this consideration. Imaaawa et al. (97) also measured the Km and V,,,,, values a t pH 6.0 with various dinucleoside monophosphates, XpY (Table X) ; K m values increased in the order of A C < G < U for both X and Y, while the V,,,,, values decreased in the order of U > G > C > A for X and of C > G > U > A for Y. When either X or Y in XpY was adenylyl residue, their K, values were smaller than those of others. This finding also supports the consideration mentioned above.

G > I >> X, indicating that the phosphodiester bonds of adenylic acid and inosinic acid without a keto group a t the position of purine base are more sensitive to RNase U, than those of guanylic acid and xanthylic acid. The resistance of TNP-RNA to RNase U, may be also attributed to the steric hindrance by a larger substituent at 2-amino groups of guanylyl residues, as with RNase T, (SO). Accordingly, no blocking a t N-1, an amino group a t the 6 position, and a nitrogen atom a t N-7 of the purine base are considered to play an important role in the affinity of the nucleotidyl residue to RNase Uz. Ribonuclease U, digestion of ApU has revealed reduced hydrolase activity in the second step of RNaee U, action (SO). When 87.4% of ApU was readily degraded to produce uridine and adenosine 2',3'-cyclic phosphate, no 3'-adenylic acid was detected. After exhaustive degradation of ApU, hydrolysis of adenosine 2',3'-cyclic phosphate occurred and 3'-adenylic acid gradually appeared. Double-stranded cytoplasmic-polyhedrosis-virusRNA obtained from silkworm was scarcely split by RNase U, (106). 4. Applications Many applications of RNase U, for sequence analysis have been reported. (1) Sequence analysis of longer oligonucleotides in RNase T, digest (90, 9.2, 107): Oligonucleotides produced by RNase T, digestion have no guanylyl residues in the nucleotide chain except 3'-terminal guanylic acid. Thus, as a limited amount of RNase U, specifically splits the phosphodiester bonds of adenylic acid in the nucleotide chain, information on localization of adenylyl residues can be obtained. The nucleotide sequence of oligonucleotide in RNase T, digest can be generally determined by using RNase U, digestion together with pancreatic RNase A digestion; for example, fragment No. 16 obtained from RNase TI digest of tRNAXIeproduced Ap, U-N"-Ap, and C-Gp by RNase U, digestion and A-Up, N"-A-Cp, and Gp by RNase A digestion. Consequently, a unique sequence of fragment No. 16 could be determined to be A-U-N"-A-C-G (90). However, the nucleotide sequence of longer

238


oligonucleotides involving more adenylyl residues in the chain, such as oligonucleotide A obtained from RNase TI digest of R17 RNA, can be determined by combining the results obtained with RNase U, and RNase A and with a new method developed by Gilham (111). After treatment with CMCT reagents reacting with uridylyl and guanylyl residues in RNA, the modified RNA can be split only after cytidylyl residues are split by RNase A. Although oligonucleotide A consists of 21 nucleotide units and contains 7 residues of adenylic acid, a unique sequence can be constructed for it as follows: AAUUAACUAUUCCAAUUUUCG (107) This experiment indicates that adenylate tracts in a nucleotide chain increase its resistance to attack by RNase U,. (2) Specific cleavage of adenylate linkages i n RNA by RNase U,: It is expected that if all guanylyl residues in RNA could be completely modified to any form resistant to RNase U,, RNase U, would split specifically adenylate linkages i n RNA ( 3 0 ) .Based on this consideration, the behavior of RNase U, to the modified RNA with CMCT reagents was investigated (108). It was found that the presence of a blocking group on either side of the phosphodiester linkage was sufficient to confer resistance to RNase U, action. Thus, the action of RNase U, on CMC-modified RNA was restricted to the cleavages of -ApA- and -ApC-. The phosphodiester bond of glyoxalguanylic acid in glyoxalGpU was more resistant to RNase Uy than other bonds (SO). Therefore, if all guanylyl residues in RNA could be modified completely with glyoxal, the fragments having the adenylic acid in 3’ termini would be obtained specifically by RNase U, digestion. However, glyoxalRNA modified completely has not yet been obtained. At present, RNase U, is a unique tool for the synthesis of oligonucleotides through adenosine 2’,3’-cyclic phobphate. The synthesis of adenylyl- (3’-5’)-nuclcoside and oligoadenylic acid by RNase U, has been investigated (T. Koike, T . Uchida, and F. Egami, unpuhlislicd). When adenosine 2’,3’-cyclic phosphate was incubated with about %fold uridine a t a low temperature in the presence of RNase U,, a synthetic product ApU was obtained with a high yield of 37% in regard to the initial adenosine 2’,3’-cyclic phosphate. Other by-products were scarcely detected and 58% of adenosine 2‘,3‘-cyclic phosphate remained unchanged, showing a low hydrolase activity of RNase U,. An optimum condition for the synthesis of ndcnylyl- (3’-5’)-nucleoside by RNaw ITTL. 111. P. T.Gilham. JACS 84, 687 (1962).

9.

239


is the same as that for the synthesis of guanylyl- (3’-5’) -nucleoside by RNase N, (see Section II,D, 4 ) . A large-scale preparation of ApU with RNase U, was also described, suggesting that enzymic synthesis of ApU by RNase U, is an excellent method because of the specific formation of 3‘,5‘-phosphodiester linkage, good yield, simplicity of technique, and economy of time (SO). When adenosine 2‘,3’-cyclic phosphate was incubated with RNase U, for 10 days a t 4” to produce oligo Ap, small oligomers such as ApA cyclic-p (14.1724, ApAp (2.7%), and trimer (5.4%) were obtained, suggesting that RNase U, is somewhat suitable for the synthesis of relatively small oligomers. Ribonuclease U, may bc used for the synthesis of guanylyl-(3’-5’)nucleoside and for the addition of adenylyl residue to 5‘ terminal of oligonucleotides including no guanylyl residue. By the fractionation of RNase U,digests of RNA, oligonucleotides of defined sequence with 3’terminal adenylic acid can be prepared.

111. Other Microbial RNases of Special Interest

Other microbial RNases of special interest from points of view of enzymology or of nucleic acid chemistry are briefly mentioned below. RNase A. EXTRACELLULAR

OF

B. subtilis STRAIN H

An extracellular RNase of Bacillus subtilis strain H was isolated in crystalline state and its chemical nature studied by Nishimura and coworkers (112).I t has a very complicated base specificity (113, 114). However, with a few exceptions, the phosphodiester bonds of 3’-purine nucleotides are cleaved faster than those of 3’-pyrimidine nucleotides, and those of 3’-nucleotides with 6(4)-keto group are cleaved faster than those of 3‘-nucleotides with a 6(4) -amino group. Thus, when adjacent bonds are the same, the following results: -GpXp-

>

-ApNp- u p s p-

> -CpNp-

112. S. Nishimura, P w c e d u r e s Nitcleic Acid Res. p . 56 (1966). 113. P. R . Whitfeld and H. Witzel, BBA 72, 262 (1963). 114. G. W. Hushizky, A . E. Greco, R . R. Hartley, Jr., and H. A. Sober, Bioclirmistry 2, 787 (1963).

240


Although, because of its complicated specificity, it is not easy to use the enzyme with a probable expectation for the nucleotide sequence analysis of RNA, sometimes it has been used with success (115,116). It may be useful especially to digest the large fragments produced by RNase TI. I n this case, it usually splits 3’-adenylyl and 3’-uridylyl bonds but not 3’-cytidylyl bonds. The enzyme has also been used to produce large fragments from tRNA in the presence of Mg2+ (116, 117).

B. INTRACELLULAR RNase OF R. subtilis Nishimura and Maruo (118) extracted and RNase from cells of Bacillus subtilis strain H, which is quite different from the extracellular RNases of the same strain. It is remarkable that the digestion products of RNA by the enzyme are exclusively four nucleoside 2’,3’-cyclic phosphates. The same enzyme has been highly purified from another strain (strain K) of Bacillus subtilis, and their properties have been fully investigated by Yamasaki and Arima (119, 120).They have confirmed the findings by Nishimura and Maruo and have found, moreover, that ATP and dATP strongly inhibit the enzyme. Yamasaki and Arima suggested that ATP might participate in the regulation of intracellular RNase activity. The observation that the final digestion products of RNA by the enzyme are four nucleoside 2’,3’-cyclic phosphates led Ukita and coworkers (121) to utilize the enzyme for the synthesis of various dinucleoside monophosphates. Indeed they have successfully synthesized UpU, CpU, ApU, GpU, UpC, CpC, ApC, and GpC in good yields (from 20 to 75% based on initial nucleoside 2’,3’-cyclic phosphates) by the reaction Ncyclic-p + IV’ NpN’ Other similar enzymes such as intracellular RNase of Azotobacter agilis (122) (strain C) may be used for the same object. --t

115. G . C. Brownlee. F. Sangcr, and B. G. Bnrrell. J M B 34, 379 (1968). 116, F. Haradn, F. Kimura. and S. Nishimura, BBA 195, 590 (1969). 117. K. Oda. F. Kirnura, F. Harada, and S. Nishimura. BBA 179, 97 (1969). 118. S. Nishirnura and B. Maruo, BBA 40, 355 (1960). 119. M. Yamasaki and K . Arirna, BBA 139, 202 (1967). 120. M. Ynrnasaki and K. Arima, BBRC 37, 430 (1969). 121. M . Saito, Y. Furuichi, K. Takeishi. M. Yoshida, M. Ynrnasaki. K. Arima, H. Hayntsu. and T . Ukita. BBA 195, 299 (1969). 122. I. Shiio. K. Ishii. and S. Shirnizu. J. Biochem. ( T o k y o ) 59, 363 (1966).

9. MICROBIAL

241

RIBONUCLEASES

C. RNase PP,

OF

Physarum polycephalum

Hiramaru et al. (123, 124) isolated four RNases and two nucleases from a slime mold, Physarum polycephalum. One (RNase PP,) of the RNases was found to be a novel enzyme. It hydrolyzes RNA in an endonucleolytic way, producing PA, pG, and oligonucleotides bearing 5'-phosphates group with a high preference for purine with regard to 3'-terminal nucleotides. Although the specific cleavage of RNA has to be further confirmed in different conditions, it is suggested that the enzyme will hydrolyze polyribonucleotides as shown below in appropriate experimental conditions. This degradation has the advantage that 3' and 5' terminals in the digests may be easily distinguished by alkaline hydrolysis as nucleosides and nucleoside diphosphates, respectively. pG pA plT pC pC pG pA pA pG pG pC pU pU pG pA pC pU

I

RNase P A

pG/pA/pU PC PC PG/PA/PA/PG/PG/PC PU PU PG/PA/PCPU

D. RNases 11, 111, IV, AND V

OF

E . coli

Among RNases of E . coli so far reported, RNase I is a typical nonbase-specific RNase and is listed in Table XI1 of Section IV. Ribonucleases 11, 111, IV, and V are very characteristic enzymes. The RNase designated RNase I1 of E . coli by Spahr (125) has been purified and extensively studied by Singer and co-workers (126, 127). Ribonuclease I1 of E. coli requires the presence of both a monovalent cation (K') and a divalent cation (Mg*+) for its activity. It has no base specificity but is specific for single-stranded polyribonucleotides. It is an exonuclease, attacking polyribonucleotides from the end of the chain bearing a 3'-OH group to produce 5'-mononucleotides. It hydrolyzes short chain oligoribonucleotides (chain length less than eight) with difficulty. The enzyme may be useful for the sequence analysis near 3' terminal and for obtaining a cluster near 5' terminal. Ribonuclease I11 (128, 129) of E . coli was discovered in 30s extracts 123. M. Hirnmaiu, T. Uchida, and F. Egnnii, J . Biochem. ( T o k y o ) 65, 697 (1969). 124. M. Hiramnru. T. Uchida. and F. Egami, J . Biochem. ( T o k y o ) 65, 701 (1969). 125. P. F. Spnlir, JBC 239, 3716 (1964). 126. M. F. Singer and G . Tolbert, Biochemistry 4, 1319 (1965). 127. N. G. Xossal and M. F. Singer, JBC 243, 915 (1968). 128. H. D. Robertson, H . E. Webster, and N. D. Zinder, Virology 32, 718 (1967). 129. H. D. Robertson, R . E. Webstcr, and S . D. Zinder, JBC 243, 82 (1968).

242


of E. coli. When RNA from q-/3 phage was added to 30s extracts of E. coli to direct protein synthesis, it was found that only the RNase A resistant double-stranded fraction of the RNA was digested completely. This observation led to the discovery of RNase I11 of E . coli. The enzyme remains within osmotically shocked cells ( E . coli 526) attached to the ribosomes. It sediments with the ribosomes in less than 0.20M NH,Cl but is detached a t higher concentrations; thus, it can be liberated from the ribosome fraction with higher NH,Cl concentrations. The free enzyme shows an absolute specificity for polymers containing double helical polyribonucleotide regions. Other polymers (single- and double-stranded DNA’s, single-stranded RNA’s) are not digested, nor do they inhibit the digestion of double-stranded RNA’s when present in excess. Thus the enzyme digests reovirus RNA, polyAU, (poly G) * (poly C ) , but not f, RNA, poly C, f, DNA, E. coli DNA, poly (A-T). I n the case of RNA from phage q-8, only the double-stranded region can be digested by the enzyme. Ribonuclease I11 shows an absolute requirement for divalent cations (Mg2+and Mn2+), and for monovalent cations (K+, NH,+, and Na”); Ca2+ cannot replace Mg2+ or Mn2+.No definite pH optimum is found; however, since it is much more active in alkaline medium (pH 7.6 ) , it is recommended that experiments be carried out a t pH 7.6. The mode of action of the enzyme has not been fully studied. It appears to be endonucleolytic, and the internucleotide cleavage appears to yield a 3’-phosphate and a 5’-hydroxyl group without intermediary formation of 2’,3’-cyclic phosphates. According to Libonati ( I S U ) , Mg2+is required, but K+ is an activator. The lower activity of the enzyme a t lower salt concentration might be partly attributed to the conformational change of RNA. The enzyme is somewhat heat stable a t acidic medium. As the discoverers of the enzyme suggest (129), RNase I11 of E. coli will be an important tool in studies of the function and structure of RNA : “Particularly relevant are studies on the replication of RNA-containing viruses, all of which have a double-stranded stage in their life cycles. Additionally, it may yield specific limited cleavages of such single-stranded RNA molecules as tRNA, ribosomal RNA, and phage RNA. Finally its ability to digest the RNA of DNA-RNA hybrids should provide a further measure of specificity in DNA-RNA hybridization experiments.” Ribonucleases IV and V of E. coli are extremely interesting enzymes, 130. M. Libonati, Boll. SOC.Ital. Biol. Sper. 44, 786 and 789 (1968).

9.


243

but because this nature is quite different from that of typical RNases, they can not be treated adequately here. Ribonuclease IV purified from an RNase I minus strain of E. coli (MRE-600) specifically cleaves R17 RNA into two large fragments, one sedimenting a t about 15s carrying the 5’ terminal of the original molecule, and the other sedimenting a t about 21 S (131). The nature of the bonds cleaved by the enzyme is not known. A new RNase activity, tentatively named RNase V , was found in cell-free extracts of E . coli. Ribonuclease V is an exoribonuclease attacking mRNA from 5‘ to 3‘ terminal producing 5’-mononucleotides. It is characterized by the requirements of ribosomes, G and T factors, tRNA, K+ or NH,’, Mg2+,GTP and a sulfhydryl compound; by its specificity; and by the fact that it degrades poly U, poly A, T, phage mRNA or E . coli mRNA, but not ribosomal RNA (132). It should be mentioned here that a quite different RNase was designated “RNase V of E. coli” in France (133).It hydrolyzes ribosomal RNA, BSRNA, poly U, poly A, poly C , and R17 RNA, but not tRNA.

IV. List of Microbial RNases

Studies on microbial RNases began in 1924 when Noguchi found ribonucleic acid degrading enzymes in Takadiastase (134). Since then extensive studies have been carried out on RNA degrading enzymes. It is rather surprising that guanyloribonuclease so widely distributed in microorganisms was found only in 1957. This is because earlier studies did not consider base specificity. Even quite recently studies on nucleases or ribonucleases do not consider base specificity or do not separate nuclease mixtures from each other; thus, information available on microbial RNases is still scant. Microbial RNases with known substrate specificity are listed in Table XII. Nucleases with DNase activity are not included. It should be noted here that the lists of RNases in both animal and plant kingdoms are presented in the monograph by Privat de Garilhe ( 4 ) and in a chapter by E. A. Barnard in Annual Review of Biochemistry (1969) (135). 131. P.F.Spahr and R. F. Gesteland, Proc. N n l l . Acad. Sci. U . S. 59, 876 (1968). 132. M. Kuwano, C. Ning Kwan, D. Apirion,D. Schlessinger, Proc. Natl. Acad. Sci. U.S.64,693 (1968). 133. F. Ben Hamida and F. R. William, Bull. SOC.Chim. Biol. 51, 1545 (1969). 134. J. Noguchi, Biochem. 2. 147, 255 (1924). 135. E.A. Barnard. Ann. R e v . Biochem. 38, 677 (1969).

TABLE XI1 LIST OF MICROBIAL RNasesa Individual name or source

PH optima

Heat stability

Awtobacter agilis

7.5

Yes in acid

Bacillus cereus

7.9

30,000-40,OOO +Np Non-s

Bacillus pumilus

7.9

10,OOO-15,000

Bacillus subtilis H ( B . amyloliquejacias)

7.5-8.5 Yes

B. subtilis Marburg

5.0

B . subtilis (intracellular)

5.8

Clostridium acetobutylicum RNase I1 E. coli RNase I

4.5

RNase I1

8.1

7-8

MW

-+

N

Other information

>p

- + N p G-s Tra, endo, -+N Np, Non-s

>p+

Intracellular RNase: solnbilization and purification ; effectors; activation by Ca*+ Purification; inactivation by EDTA Purification; no inactivation by EDTA Crystallization; relative specificity, amino acid composition; see text Partial purification

Tra, endo, --* N > p Np, Non-s, hydrolysis very slow Tra, endo, -+N > p Inhibition by EDTA; Non-s effects of nucleotides; see text Endo, -+Np Enzyme formation; purification; effectors

No

-+

Tra, endo, -+N Np, Non-s

Yes in acid

NO

Specificityb Bacteria Tra, endo, Non-s

10,700

E

65,000

>p

-+

Hyd, exo, +pN, Non-s

Purification; basic protein; relative specificity; intracellular localization; activation by Mgz+; compare with related enzymes See text

Ref.

(122)

(140) (140)

(112-114)

(1.41) (118-121)

Y (142)

d

d

X

E

(143-146)

&!

9 q H

2

(125-127)

5

RNase I11 RNase IV and V Lactobacillus casei

7.6-9.7 8.1-8.2 (7.4)

L. plantarum RNase I1

8.6

YeS

M ymbaeterium avium

7.5

Proteus mirabilis RNase I1

7.0

Salmonella typhimurium

7.0

Not so stable

Thwbacillua thwpam IIA IIB IA-2

7.0 9.5 5.5

YeS Yes Yes

Actinomyces

Endo, doublmtrsnded Requirement for M%+and (128-1SO) K+ or N&+; see text specific +Np and pN see text (131-133) Endo, exo, +pN Purification; activation (146) by K+ Hyd, exo, - + p N Purification; monovalent (147) and divalent cations required; single-stranded preferred Purine specific or Purification (148) preferential Tra, +N > p, Non-s Purification (14.99) CpN bonds are fairly resistant (150) Tra, endo, +N > p -+ Intracellular enzyme; Np, Non-s, hydrolysis purification; Na+, K+, and Mgz+ have no effect; very slow. poly I resistant tRNA fairly resistant Endo, +Np, pyr-s

aureoventieillatus

s. albogriseolus

7 . W 3 . 5 Yes in acid

S. erythreus

7.3-7.4

Yes

(151, 162)

Purification; specificity for methylated guanylates; splitting rate of GpNp: GPUP > GPCP Purification; effectors

(137, 138)

8

La 0

E! P F La

CI

8

Z

2

ED

m

Endo, +Np, pyr-s Streptomyces 13,000 Tra, endo, +N NP, G-s

7.6-7.8

Purification

9

> p -+

Tra, endo, -+N > p ---t NP, G-s Tra, endo, + N > p -+ Purification; relative specificity for methylated NP,G-s guanylates

(153)

(136)

2

01

2

TABLE XI1 (Continued) Individual name or source

PH optima

Heat stability

MW

Specificity6

Other information

Ref.

Yeast Endomyces Rhdorula glutinis I

I1

4.5

-+NP

7 4

+pN, Non-s +Np, Non-s

7 7.5

No

Acrocylindrium sp.

8.0

YeS

Aspergillus niger Aspergillus oryzae RNase TI

3.0-3.5

I11 Saccharomyes cerevisiae (Bakers' yeast)

RNase Tt Aspergillus saitoi RNase M

Chalaropsis sp. RNase Ch

+Np, Non-s Hyd, exo, -Np,

Fungi (including a slime mold) Tra, endo, -+N G-s s ~ = .2.59 ~ S +Np, Non-s

7.4-7.5 Yes

11,000

4.5

YeS

36,000

4.0

YeS

30,000

Tra, endo, -+N Np, G-s Tra, endo, +N Np, Non-s Tra, endo, -+N Np, Non-s

(164)

Relation between enzyme formation and culture conditions Non-s

(166-167)

Intracellular enzyme; purification; effectors; activated by phosphate and inhibited by Zn'+

> p, Purification; inhibitors

>p+

See text

(160-169)

See text

> p + See text

>p+

Tra, endo, - + N> p NP, G-s

-+

Purification; inhibition by Zn'+, Cuf+; other effeo tors; kinetic data; photooxidation; inhibition by nucleotides

166)

tiU 1 H

(167)

cl 2-

5

Lenzites tenuis Monasczls pilotus Mucor genevensis

7.9 4.5 7.9

Neurospora craasa RNase NI

7.0

30,000-40,OOO +Np, Non-s 30,000-40,OOO -Np, Non-s 12,000 +Np, G-s Yes

11,000

RNase NZ

8.0

36,000

R N m NI

6-7

A little larger than RNase

Inhibited by EDTA Not inhibited by EDTA Scarcely inhibited by EDTA

Tra, endo, +N > p -+ Formation, see text NP, G-s Tra, endo, I ~ N> p + Formation Np, Non-s Tra, endo, + N > p + Intracellular enzyme; formation Np, G-s

N1 Genetics; inhibited by EDTA

Neurospora crassa

7.5

YeS

Physarum polycepholum RNase PPl

6.7

Yes

40,000

RNase PPt

4.5

YeS

40,000

RNase PPa

5.5

YeS

10,000

RNase PPI

4.0

No

Plmspora

7.5

Rhizopzls RNase

5.0

T r i e o d e m koningi I, I1

4.5

Tra, endo, - + N > p + Np, G-s Tra, endo, -+N > p + Np, Non-s Tka, endo, -+N > p -+ Np, Non-s Hyd, endo, +pN Tra, endo, +N Np, Pur-s

Fairly stable

am,w = 2.42 S

25,000

I

>p+

Endo, -+Np, Non-s, purine preferential

Tra, endo, +N Np, Non-s

>p +

(140) (140) (140)

lo

@,8, 102, 106) (8,102)

r

(8,102)

(loo, 101)

Inhibitors; purification

(1.w

Inhibitors; purification, see text Induced formation; inhibition by phosphate; Michaelis constant Crystallization; inhibited by Znz+,Cu2+;no inhibition by EDTA

(124)

Similar to RNase TZ

(168-170)

(171)

(179)

H

BP

? i c:

TABLE XI1 (Continued) c4

Individual name or source 111

Ustilug0 sphaerogena RNase UI

A

Heat stability

PH optima

00

MW

4.5

Specificityb

Tra, endo, -+N Np, purine preferential

Tra, endo, -+N > p + Np, C-s Tra, endo, +N > p -+ Np, Pw-s Tra, endo, +N > p + Np, Pur-s Hyd, exo, +Xp, Non-s

8.0-8..5

Yes

11 ,OOO

RNase UZ

4.5

Yes

10,oOO

RNase US

4.5

Yes

10,OOO

RNase Ud

8.0-8.5

No

Much larger than U,, Uz

Ustilago me

8.9

No

Euglena gracis Paramecium aurelio

4.5

No

6.5

Yes (I, I1 mixture)

I I1 Teirahymena pyriformis I, 11, I11

>p +

10,oOO

Tra, endo, -tN Np, G-s

> p -+

Other information

Ref.

Activated by Mg"

Induced formation; see text See text

Induced formation Induced formation

Protozoa Non-s Non-s

Inhibitors; inhibited by H$+; resistant to other metal ions

Tra, endo, +N > p -+ Np, I, 11, 111, relative specificity different

4 M urea increases the

5.5 5.0

~~~~

activity

~

With the collaboration of Dr. E. Ohmura. * Abbreviations: tra, transphosphorylation; hyd, direct hydrolysis; >p, 2',3'-cyclic phosphate; endo, endonucleolytic; exo, exonucleolytic; Non-s, nonspecific; G-s, guanine specific; Pur-s, purine specific; Pyr-s, pyrimidine specific. 0

1

s

E

9.


249

Since the information is still scant, it is probably premature t o draw general conclusions on the distribution of different RNases in animals, higher plants, and microorganisms. However, it may be pointed out that to date, RNases found in higher plants are nonbase-specific, that both nonbase-specific and pyrimidine-specific RNases are found in animals, and that guanine-specific and purine-specific RNases are found only in microorganisms. Guanyloribonueleases are, as shown in Table XII, widely distributed in microorganisms, especially in fungi and streptomyces. These guanyloribonucleases may exhibit different fine relative specificity. Indeed the action of RNase TI and guanyloribonucleases of streptomyces or actinomyces on minor components in tRNA such as methylated guanylyl bonds or on chemically modified guanylyl bonds has been reported to be different (136‘-139).Further studies on such fine relative specificity are required. 136. K. Tanaka, Procedures Nucleic Aczd Res. p. 14 (1966). 137. N. H. Abrosimova-Amelyanchik, R. I. Tatarskaya, T. V. Venkstern, V. D. hksel’rod, and A. A. Bayev, Biokhimiyu 30, 1269 (1965); Biochemistry (USSR) (English Transl.) 30, 1 W (1965). 138. It. I. Tatarskaya, N. H. Abrosimova-Amelyanchik, and V. D. Aksel’rod, Biokhimiyu 31, 1017 (1966); Biochemistry (USSR) (English Trunsl.) 31, 882 (1966). 139. N. K. Konchetkov, E. I. Budowsky, N. E. Broude, and L. M. Klebanova, BBA 134, 492 (1967). 140. G. W. Rushizky, A. E. Greco, R. W. Hartley, Jr., and H. A. Sober, JBC 239, 2165 (1964). 141. M. Nikai, I. Minami, T. Yamasaki, and A. Tsugita, JBC 57, 96 (1965). 142. M. Tomoyeda, H. Horitsu, and K. Kumagai, Res. Bull. Fac. Agr., Gifu Univ. 28, 153 (1969). 143. P. F. Spahr and B. R. Hollingworth, JBC 236, 823 (1961). 144. Y.Anraku and D. Mizuno, BBRC 18, 462 (1965). 145. Y. Anraku and D. Mizuno, J . Biochem. (Tokyo) 61, 70 (1967). 146. H.M. Keir, R. H. Mathog, and C. E. Carter, Biochemistry 3, 1188 (1964). 147. D.M. Logan and M. F. Singer, JBC 243, 6161 (1968). 148. A. Tsugita and K. Matsui, Seikagaku 41, 588 (1969). 149. M. S. Center and F. J. Behal, BBA 151, 698 (1968). 150. K. Chakraburtty and D. I?. Burma, JBC 243, 1133 (1968). 151. W. Ostrowski and Z. Walczak, Actu Biochim. Polon. 8, 345 (1961). 152. Z. Walczak and W. Ostrowski, Actu Biochim. Polon. 11, 241 (1964). 153. M. Yoneda, J. Biochem. (Tokyo) 55, 469 (1964). 154. T. Hattori and S. Nakamura, Seikaguku ( T o k y o ) 38, 563 (1966). 155. Y.Nakao and K. Ogata, Agr. B i d . Chem. (Toyko) 27, 116 (1963). 156. Y. Nakao and K. Ogata, Agr. Biol. Chem. (Toyko) 27, 499 (1963). 157. Y. Nakao and K. Ogata, Agr. B i d . Chem. (Tokyo) 27, 507 (1963). 158. Y.Ohtaka, K.Uchida, and T. Sakai, J . Biochem. ( T o k y o ) 54, 322 (1963). 159. I. Suhara, F. Kusaka, and E. Ohmura, Koso Kagaku Shimpoziumu 16, 115 (1964).

250


160. Y. Eto, Y. Goto, and M. Tomoyeda, Nippon Nogei Kagaku Taikai Abstr., p. 30 (1969).

161. Y. Azuma, H. Horitsu, and M. Tomoyeda, Nippon Nogei Kagaku Taikai Abstr., p. 30 (1969). 162. H. Horitsu, K. Okamoto, Y. Azuma, and M. Tomoyeda, Nippon Nogei Kagaku Taikai Abstr., p. 30 (1969). 163. M. Irie, J. Biochem. ( T o k y o ) 62, 509 (1967). 164. M. h i e , J. Biochem. ( T o k y o ) 65, 133 (1969). 165. M. Irie, J. Biochem. ( T o k y o ) 66, 569 (1969). 166. M. Irie, J. Biochem. ( T o k y o ) 66, 907 (1969). 167. J. H. Hash and E. Elsevier, Science 162, 681 (1968). 168. C. M. Cuchillo, J. M. Ventura, E. Concustell, and V. Villar-Palasi, R e v . Espan. Fisiol. 23, 81 (1967). 169. C. M. Cuchillo, J. M. Ventura, E. Concustell, and V. Villar-Palasi, R e v . Espan. Fkiol. 23, 87 (1967). 170. C. M. Cuchillo, J. M. Ventura, E. Concustell, and V. Villar-Palasi. R e v . Espan. Fkiol. 23, 93 (1967). 171. M. Tomoyeda, Y. Eto, and T. Yoshino, ABB 131, 191 (1969). 172. M. Hamada and M. I r k , Seikagaku 41, 587 (1969). 173. M. Yanagida, T . Uchida, and F. Egami, Nippon Nogei Kagaku Kaishi 38, 531 (1964). 174. J. Fellig and C. E. Wiley, Science 132, 1835 (1960). 175. G. Gross, B. Skoczylas, and W. Tunski, Acta Protozoal. 4, 59 (1966). 176. L. H. Lazarus and 0. H. Scherbaum, BBA 142, 368 (1967).

Bacterial Deoxvribonucleases I. R. LEHMAN I. Introduction

.

.

.

.

.

.

11. Exonucleases . . . . . . -1.E . coli Exonucleases I :uid 111

.

. . B. Phage A-Induced Exonuclcmc~ . C. E . coli Exonurleasr IV . . .

.

. . . .

.

. . . . D. Pltitge T2- and T4-Intluced ISxonuc1r:iscs .

.

. . . . .

.

. . . . .

.

. . . . .

E. Exonucleases Associated with E . coli DNA Polymerase F. B . subtilis Phage SP-3 Induced Exonuclease . . . G . E . coli ATP-Dependent DNase . . . . . . 111. Endonucleases . . . . . . . . . . . . A. Nonspecific Endonucleases . . . . . . . B. Specific Endonucleases . . . . . . . .

.

251

.

255 258 259 259 259 262

. 252 . 253 . 253 . 254 . 255 .

. . . .

I. Introduction

Deoxyribonucleases, enzymes which hydrolyze specifically the internucleotide bonds of polydeoxynucleotides, have been described in many bacteria. A useful general classification first introduced by Laskowski ( 1 ) divides these enzymes into exonucleases and endonucleases depending upon their mode of attack. Endonucleases attack polynucleotides at many points within the chain, generally producing only a small proportion of mononucleotides ; cxo~iucleases catalyze a stepwise attack a t either the 3’ or 5’ terminus of a polynucleotide, producing predominantly mononucleotides. Recent detailed investigation of several of the purified bacterial deoxyribonucleases has indicated that this categorization is overly 1. M . Laskowski, Ann. N . Y . Acad. Scz. 81, 776 (1959). 251

252

I. R. LEHMAN

simplified and in fact a single physically homogeneous protein may have the capacity to catalyze both the exo- and endonucleolytic cleavage of diester bonds depending upon the structure of its macromolecular substrate. All of the bacterial deoxyribonucleases that have been examined in detail possess a specificity directed in varying degrees toward the secondary structure of the polydeoxynucleotide. With one recent exception, none of the deoxyriboriucleases shows a simple base specificity whereby they attack phosphodiester bonds adjacent to a single base. However, it is now clear that several of the endonucleases may possess an extremely high order of specificity and have the capacity to recognize and attack one or a few phosphodiester bonds in polydeoxynucleotide chains composed of many thousands of internucleotide linkages. In this review, the bacterial deoxyribonucleases to be considered will be divided into exonucleases and endonucleases. The latter category will be further subdivided into “nonspecific” endonucleases ; that is, enzymes which do not show a high level of specificity for the internucleotide bond split and are, as a result, capable of degrading polynucleotides to a mixture of relatively small oligonucleotides and “specific” endonucleases, which display a high level of specificity for a given sequence within the polynucleotide chain, and as a consequence, introduce only a very few cleavages into a high molecular weight polynucleotide. As noted above, the enzymes to be discussed are those which attack polydeoxynucleotides exclusively. Thus, nucleases from, for example, Bacillus subtilis (2-4), Serratia marcescens ( 5 ) , and Staphylococcus aureus (6),which attack both RNA and D N A will not be considered. Such an enzyme from S. aureus is, however, the subject of Chapter 7 by Cotton and Hazen and Chapter 8 by Anfinsen et al. in this volume.

It. Exonucleases

The bacterial D N A exonucleases which have been purified and examined in detail are with two exceptions derived from E . coli and E . coli infected with bacteriophages; all show a strong specificity for the secondary structure of their polydeoxynucleotide substrate.

2. 3. 4. 5. 6.

I. M. Berr. J. R. Chien. iintl I. R. Lehniun, JBC 242, 2700 (1967). R. Okazaki, T. Okazaki, and K. Sakabe, BBRC 22, 611 (1966). H. C. Birnboim, J . Bacterial. 91, 1004 (1966). M. Nestle and W. K. Roberts, JBC 244, 5213 and 5219 (1969). L. Cunningham. B. W. Catlin. and M. Privut De Garilhe. JACS 78, 4642 (1956).

10.

BACTERIAL DEOXYRIBONUCLEASES

253

A. E. coli EXONUCLEASES I AND IIL Both E. coli exonucleases I and I11 have been considered in a recent review (7) and will not be discussed in detail here. It is, however, worth noting again that they are prototypes of nucleases with an extreme specificity for the secondary structure of their substrate. Thus, exonuclease I attacks only single-stranded polynucleotides and exonuclease I11 has an equally strong preference for double-stranded structures. In both instances, once the structural requirements have been met, the enzymes initiate their attack a t the 3'-hydroxyl termini sequentially liberating 5'-mononucleotides. In the case of exonuclease 111, hydrolysis ceases once the bihelical structure is lost as a result of sustained exonucleolytic attack. Exonuclease I is unable to cleave dinucleotides ; hence, the terminal and penultimate residues situated at the 5' end of the polynucleotide substrate remain as a residual dinucleotide. Recent experiments by Masamune and Richardson (8) have demonstrated that exonuclease I11 is able to initiate its attack a t an internally located 3'hydroxyl group in a DNA duplex (i.e., a t a single-stranded interruption) as well as a t the 3'-hydroxyl group at the end of the chain. Finally, it should be noted that exonuclease I11 possesses an intrinsic DNA-phosphatase activity which specifically removes 3'-phosphoryl groups from double-stranded polynucleotides. This enzyme when acting on a 3'-phosphoryl-terminated DNA, first removes the 3'-phosphoryl group as inorganic phosphate, then proceeds as an exonuclease with the stepwise release of deoxynucleoside 5'-monophosphates (9). Lacks and Greenberg (10) have purified an exonuclease from Diplococcus pneumoniae with properties very similar to E. coli exonuclease 111. The pneumococcal enzyme acts preferentially on native DNA producing 5'-mononucleotides and single strands which are not susceptible to further attack. Like exonuclease I11 it shows an intrinsic 3'-phosphoryl-DNA phosphatase activity. A DNA phosphatase-exonuclease activity has also been reported in B. subtilis (11).

B. PHAGEA-INDUCEDE'XONUCLEASE The exonuclease synthesized after induction of h lysogens or after infection with virulent mutants of this phage has received a great deal 7. I . R . Leliman, Ann. Rev. Biochem. 36, 645 (1967). 8. Y. Masamune and C. C. Hichardson, Biophys. Soc. Abstr. p. 18a (1970) 9. C . C. Richardson and A . Kornberg, JBC 239, 242 (1964). 10. S. Lacks and B. Greenberg, JBC 242, 3108 (1967). 11. T. Okazaki and A . Kornberg, JBC 239, 259 (1964).

254

I. R. LEHMAN

of attention because of its involvement in h-recombination. The h exonuclease has been obtained in physically homogeneous, crystalline form and is the only one of the bactcrial DNases available a t this level of purity (12). Phage h possesses its own recombination system, the “red” system (13, 14) which permits its DNA to undergo recombination in recombination deficient (rec-) hosts ( 1 5 ) . Thc rctl systcm is composed of three complementation groups, two of which define the structural gene for the A exonuclease (16). The h exonuclease is antipodal to exonuclease I11 in its polarity of attack ; thus, it specifically degrades double-stranded DNA, starting a t the 5’-phosphoryl terminus, sequentially liberating 5’-mononucleotides. As in the case of exonuclease 111, single strands are generated which are then not susceptible to further hydrolysis (17). The h-exonuclease shows a very strong preference for termini bearing 5’-phosphoryl groups. Unlike exonuclease 111, the h-induced nuclease is unable t o attack a t single-stranded breaks within a DNA duplex (8). Clearly, both exonuclease I11 and h exonuclease could be responsible for the production of the single-stranded regions in recombining DNA molecules postulated in most current models of genetic recombination (18).

C. E. coli EXONUCLEASE IV Exonuclease IV is distinguished from the other DNA exonucleases of E. coli by its strong preference for relatively short chain oligonucleotide substrates (19). Native and denatured DNA are degraded a t rates less than one-twentieth those observed with oligonucleotide mixtures derived from pancreatic DNase digests of DNA. At present it is not clear whether this reflects an inability of the enzyme to bind to macromolecular DNA or the relatively low concentration of DNA molecules compared to oligonucleotide chains present in the usual assay mixtures. Exonuclease IV requires magnesium ion and is optimally active a t alkaline p H (pH 8-9). It produces 5’-mononucleotides exclusively, presumably by an exonucleolytic attack starting a t the 3’terminus, although this latter point 12. J. W. Little. I. R. Lehmnn. and A. D. Kaiser, JBC 242, 672 (1967). 13. H. Echols, R. Gingery, and L. Moore, J M B 34, 251 (1968). 14. E. R. Signer and J. Weil, J M B 34, 261 (1968). 15. A. J. Clark, J . Cellular Physiol. 70, Part 11, Suppl. 1, 165 (1967). 16. C. M. Radding, J. Szpirer, and R. Thomas, Proc. N a l l . Acad. Sci. U . S. 57, 277 (1967). 17. J. W. Little, JBC 242, 679 (1967). 18. C. C. Richardson, Ann. Rev. Biorhem. 38, 795 (1969). 19. A. E. Oleson and J. F. Koerner, JBC 239, 2935 (1964).

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has not been established definitively. Koerner and his colleagues have found that exonuclease I V separates into two subfractions, termed IVa and IVb, upon gradient chromatography on columns of DEAE-cellulose (20). Though chromatographically distinct, exonucleases IVa and IVb have identical pH optima, divalent cation requirement, substrate specificity, and thermolability. The origin and significance of the two fractions are unknown.

D.

PHAGE

T2-

AND

T4-INDUCED EXONUCLEASES

An exonuclease catalytically very similar to exonuclease IV appears after infection of E. coli with phages T2 and T4 (19).Yet another catalytically indistinguibhable cxonuclease activity is synthesized after T2 and T 4 phage infection and is closely associated with the T2- and T4induced DNA polymerases (21, 2 2 ) . An amber mutant which maps in the structural gene for the T4 DNA polymerase has been found by Nossal to induce the polymerase-associated exonuclease but not the polymerase ( 2 3 ) .Both of the phage-induced exonucleases act optimally on mixtures of small oligonucleotides producing 5’-mononucleotides. Recent experiments by Richardson et al. (8) and by Kornberg and his colleagues (24) have demonstrated a 3’+5‘ polarity of attack by the polymerase-associated exonuclease. E. EXONUCLEASES ASSOCIATED WITH E . coli DNA POLYMERASE The hydrolytic activity of E . coli DNA polymerase has until recently been termed exonticlease 11 primarily because of the uncertainty as to whether the polymerizing and hydrolytic functions of the purified enzyme were part of a single unit ( 2 5 ) .I t is now apparent as a result of studies in Kornberg’s laboratory that DNA polymerase is a homogeneous protein composed of a single polypeptide chain with a single binding site for a DNA terminus and that the polymerizing and hydrolytic activities both involve a common active site ( 2 6 ) . A detailed investigation of the 20. S. E. Jorgensen and J. F. Koerner. JBC 241, 3090 (1966). 21. E. C. Short and J. F. Koerner. Proc. Natl. Acad. Sci. U.S. 54, 595 (1965). 22. M. Goulian, Z. J. Lucas, and A. Kornberg. JBC 243, 627 (1968). 23. N . G. Nossal, JBC 244, 218 (1969). 24. N. R. Cozzarelli, R . B. Kelly, and A. Komberg, J M B 45, 513 (1969). 25. I. R. Lehman and C . C.Richardson. JBC 239, 233 (1964). 26. A . Kornberg, Science 163, 1410 (1969).

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exonucleolytic activity by Deutscher and Kornberg (27) and independently by Klett et al. (28) has in fact shown that there are two exonucleolytic activities of opposite polarity intrinsic to the polymerase. One catalyzes an attack starting at 3’-hydroxyl terminus (3’ + 5’) ; the second attacks a t the 5’-phosphoryl terminus (5’- 3’). 1. 3‘ + 5’ Exonuclease

Early studies of the nuclease activity associated with purified preparations of E . coli DNA polymerase indicated that it catalyzed an exonucleolytic attack on both double- and single-stranded polydeoxynucleotides beginning a t the 3’-hydroxyl end, liberating deoxynucleoside 5’-monophosphates. Upon prolonged incubation, hydrolysis to mononucleotides was complete ( 2 5 ) . It is now clear that hydrolysis of single-stranded polymers results exclusively from the 3‘ + 5’ exonuclease component of the polymerase. This activity is optimal in tris buffer a t p H 8.6 and in glycine buffer a t p H 9.2. It has an absolute requirement for a free 3‘hydroxyl group ; 3’-phosphoryl-terminated polynucleotides are resistant ( 2 7 ) .It therefore resembles very closely the exonuclease activity associated with the T4-induced D N A polymerase ( 2 2 ) .Like the phage-induced enzyme, the 3’ + 5‘ exonuclease activity of the E . coli DNA polymerase is almost completely abolished under conditions of DNA synthesis (22, 2 9 ) . Presumably a single 3‘-hydroxyl terminus site serves for either polymerization or 3’ + 5’ hydrolysis. Since the 3’ 3 5’ exonuclease activity removes nucleotide residues a t the same site in the polynucleotide chain (3’-hydroxyl) a t which the polymerizing activity adds nucleotides it has been suggested that this exonuclease activity represents an errorcorrecting mechanism whereby improperly paired nucleotides are removed in the course of synthesis (26). 2. 5’ + 3’ Exonuclease

Studies by Klett et al. (28) with a synthetic d (A-T) copolymer in which diamino purine replaced adenine a t the 3’ end and by Deutscher and Kornberg (27) with 3’-phosphoryl-terminated polynucleotides led to the finding that purified preparations of E . coli D N A polymerase possess a 5‘ += 3‘ exonucleolytic activity. In the latter studies DNA preparations with 3‘-phosphoryl termini introduced by the action of staphylococcal nuclease (6) and therefore inscnsitiw to 3’+ 5’ exonuclease action were 27. M. P. Deutscher and A. Kornberg, IBC 244, 3029 (1969). 28. R. P. Klett, A. Ccrarni, and E. Reich, Proc. Natl. Acnrl. Sci. ”. 8. 60, 943 (1968). 29. D. Brutlag, M. R. Atkinson, P. Setlow. and .4.Kornberg, BBRC 37, 982 (1969).

10.

BACTERIAL DEOXY RIBONUCLEASES

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found to be extensively degraded by purified DNA polymerase preparations yielding 5’-mononucleotides and oligonucleotides terminating in a 3’-phosphoryl group. Detailed investigation of this phenomenon showed that the 5’+ 3’ exonuclease activity occupies a site on the enzyme distinct from the 3’+ 5’ exonuclease. Unlike the latter i t has a strong preference for double-stranded polynucleotide substrates and displays a broad pH optimum ranging from p H 7.4 to 9.2 (a7). The products of 5‘+ 3‘ exonuclease action consist of dinucleotides in addition to 5‘mononucleotides. This was demonstrated in an experiment in which hydrolysis of d (pT) 300 was compared with d (pT)3oo annealed to d (PA) 4&oo and d(pT)joo blocked at the 3’ terminus by a dideoxythymidine residue also annealed to d (PA) 4ooo. The dinucleotide d (pT), was observed among the products of hydrolysis only in the latter two cases; it was not found when single-stranded d (pT)3oo was degraded, i.e., in the absence of 5’- 3‘ exonuclease action. Kinetic studies showed that the rate of appearance of dinucleotides relative to mononucleotides during 5’ 3’ hydrolysis was not altered as a function of time ( 2 4 ) . In further contrast to the 3’ 4 5’ exonuclease, the 5’ + 3’ exonuclease activity was strongly stimulated (as much as 40-fold) by simultaneous synthesis ; moreover, an increased frequency of oligonucleotides was observed among the products of 5 ‘ 4 3’ hydrolysis during synthesis, and oligonucleotides in lengths ranging up to the hexanucleotide appeared (SO). The increase in 5‘ 3’ exonuclease activity accompanying polymerization of nucleotides a t the 3’ termini of the primer strands has been explained by postulating that the DNA polymerase binds a nicked region of a DNA duplex in its active center, thus bringing the 3‘-hydroxyland 5‘-phosphoryl-terminated chains in close alignment. The synthetic activity, by advancing the 3‘-hydroxyl terminus keeps the latter adjacent to or near the 5’ end and increases the rate of release of 5’-mononucleotides at this terminus. The release of oligonucleotides may be the result of the failure of the DNA polymerase to cleave terminal bonds, thereby resulting in exposure of the second or subsequent phosphodiester linkages to hydrolysis a t the 5’+ 3‘ exonuclease site (24).In the case of d(pT)300 with a tetradeoxycytidylate region a t the 5’ terminus annealed to d the deoxycytidylate residues which do not hydrogen bond to the d (PA)4ooo were found to be excised as oligonucleotides ( 3 1 ) .Hence, the 5’- 3‘ exonuclease is able to hydrolyze double-stranded DNA in the duplex region beyond a set of non-hydrogen-bonded residues, removing

-

-

30. R. B. Kelly, N . R. Cozzarelli. M. P. Deutscher. I. R. Lehman, and A. Kornberg, JBC 245, 39 (1970). 31. R. B. Kelly, M. R. Atkinson, J. A . Huberman, and A. Kornberg, Nature 224,

495 (1969).

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oligonucleotides containing these residues. This property of the 5' + 3' exonuclease may play a general role in the removal of mismatched regions, for example, pyrimidine dimers. Kelly et al. (31) have, in fact, found that the 5' + 3' exonuclease can excise oligonucleotides containing thymine dimers from ultraviolet-irradiated d (pT) annealed to d(pA),," as well as from irradiated DNA. I n contrast, the 3'+ 5' exonuclease like E . coli exonuclease I and snake venom phosphodiesterase (31, 3 2 ) is blocked at the point in the (pT):joo a t which it encounters a thymine dimer. Studies by Jovin et al. (33) had shown that acylation of the enzyme with N-carboxymethylisatoic anhydride resulted in a total destruction of polymerase activity with a concomitant 9-fold enhancement of exonuclease activity as measured a t pH 7.4. More recently, Brutlag e t al. (29) and Klenow and Henningsen (34) independently discovered that the 5'+ 3' exonuclease activity could be dissociated from the remainder of the DNA polymerase molecule by limited proteolysis with subtilisin or trypsin. Thus, treatnicnt of the polymerase, a single polypeptide chain of molecular weight 109,000, with these proteases led to the generation of two fragments of 76,000 and 34,000 molecular weight. The larger fragment retained the polymerizing activity and the 3' + 5' exonuclease activity ; the small fragment contained the 5' + 3' exonuclease. The 76,000 molecular weight fragment derived from the E . coli DNA polymerase is therefore similar to the T4-induced polymerase in that both enzymes, contain only the 3'+ 5' exonuclease in addition to the polymerizing activity. The phage-induced enzyme, despite the lack of 5' 3 3'-exonuclease, has a molecular weight of approximately 110,000 (22).

F. B. subtilis PHAGE SP-3-INDUCED EXONUCLEASE Trilling and Aposhian have partially purified a DNase from extracts of B. subtilis infected with phage SP-3 ( 3 5 ) .This enzyme requires magnesium ion and shows optimal activity between pH 7.8 and 8.9 in Tris buffers. It is highly specific for denatured DNA and appears to catalyze a unique type of exonucleolytic attack beginning a t the 5' end of the chain which sequentially releases dinucleotides. Neither mono32. L. Grossman, J. C . Kaplan, s. R. Kushner. and I. Mahler, Cold Spriltg Harbor Symp. Quant. Biol. 38, 229 (1968). 33. T. M. Jovin, P. T. Englund, and A . Kornberg, JBC 244, 3009 (1969). 34. H. Klenow and I. Henningsen, Proc. Nutl. Acad. Sci. U.S. 65, 168 (1970). 35. D. M. Trilling and H. V. Aposhian, Proc. Natl. Acad. Sci U. S . 60, 214 (1968).

10. BACTERIAL

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259

nucleotides nor oligonucleotides of intermediate size between dinucleotides and the polymeric DNA substrate (with the exception of a few percent trinucleotides) is observed at any time during hydrolysis.

G. E. coli ATP-DEPENDENT DNASE An ATP-dependent DNase has been partially purified from extracts of E . coli by Oishi (36) and Barbour and Clark (37). It shows an absolute requirement for magnesium or manganese ion and has a broad pH optimum ranging from pH 7.5 to 9.5. The partially purified enzyme preferentially degrades native DNA (including glucosylated T4 DNA) and has an almost absolute requirement for ATP or dATP. Current preparations of the enzyme are also active on denatured DNA; however, there is only a slight stimulation of hydrolysis by added ATP. This latter activity may therefore represent some contamination with exonuclease I. The mode of attack is stated to be exonucleolytic. A particularly interesting feature of the ATP-dependent DNase is its possible involvement in genetic recombination. Thus, certain recombination deficient strains of E . coli with the recB- and r e d - phenotypes (16) appear to lack this enzyme.

111. Endonucleases

A. NONSPECIFIC ENDONUCLEASES

1. E . coli Endonuclease I The existence of a deoxyribonuclease in E . coli bound to an inhibitory RNA was first suggested by Kozloff (38) who found that the DNase activity of freshly prepared extracts could be markedly enhanced by pretreatment with ribonuclease. The enzyme was subsequently purified and freed of inhibitor (39). The purified enzyme termed endonuclease I could, in turn, be competitively inhibited by a variety of RNA's inM (nucleotide) cluding transfer RXA, and K , values as low as have been observed (40). Examination of various purified RNA species and synthetic polyribonucleotides for their inhibitory activity has led M. Oislii, Proc. N a t l . Acud. Sci. U . S. 64, 1292 (1969). S. D. Barbour and A . J. Clark, I'roc. N o t / . Acad. Sci. U . S. 65, 955 (1970). L. M . Kozloff. Cold Spritig Harbor Sump. Qiront. Bid. 18, 209 (1953). I. H . Lehman, G. G . Roussos, und E. A. Pratt. JBC 237, 819 (1962). 40. I . R. Lehman, G. G. Roussos, and E. A . Pratt, JBC 237, 829 (1962).

36. 37. 38. 39.

260

I. R. LEHMAN

to the suggestion that some ordered conformation is required for a polynucleotide to be an effective inhibitor ( 7 ) . Endonuclease I has a pH optimum ranging from pH 7.5 to 8.5 and absolutely requires magnesium or manganese ion for activity. It degrades native DNA a t a rate sevenfold greater than denatured DNA. Attack is purely endonucleolytic yielding oligonucleotides terminated by 5’-phosphoryl groups. The enzyme shows little, if any, base specificity and limit digests produced after exhaustive digestion of DNA contain a mixture of oligonucleotides with an average chain length of 7 residucs. Analysis of the oligonucleotides has shown an almost random distribution of nucleotides a t the 3’ and 5‘ termini. Studier (41) and independently Bernardi and Cordonnier ( 4 2 ) have found that endonuclease I cleaves both strands of a DNA double helix a t or near the same level. Paoletti et al. (43) have examined this question in greater detail employing a fluorometric method based on the increase in the amount of ethidium bromide bound to closed circular duplex DNA upon the introduction of a single-stranded break ( 4 4 ) . In contrast to pancreatic DNase in which there was a linear increase in fluorescence, nonlinear kinetics were observed with endonuclease I. Further investigation of this phenomenon led to the suggestion that endonuclease I possesses a significant exonucleolytic component with the result that approximately 400 nucleotides are removed at each endonucleolytic scisson introduced by the enzyme. Consequently, endonuclease I does not promote a clean double-stranded break but rather causes a “shattering” a t the point of cleavage, leading to the liberation of small oligonucleotides, presumably by an exonucleolytic mode of attack. 2. Streptococcal and Pneumococcal Endonucleases

A deoxyribonuclease termed streptodornase, optimally active a t pH 7.0 in the presence of magnesium ion, has been partially purified from culture fluids of Streptococcus pyogenes (45). This enzyme yields a distribution of products from DNA very similar to that seen with E . coli endonuclease I: Only traces of mono- and dinucleotides are found, the majority of products being rather large oligonucleotides terminated by 41. F. W . Studier, J M B 11, 373 (1965). 42. G. Bernardi and C. Cordonnier, J M B 11, 141 (1965). 43. C. Paoletti. J. B. LePccq. and I. R. Leliman. J M B (1971) (in press). 44. R. Radloff. W. Baucr, and J. Vinograd, Proc. Natl. Acad. Sci. U . S . 57, 1514 (1967). 45. M. Laskowski, “The Enzymes,” 2nd ed.. Vol. 5. p. 123. 1961.

10.


26 1

5‘-phosphate groups. Like E. coli endonuclease I, streptococcal endonuclease is inhibited by RNA. A careful study of various strains of Streptococcus by Wannamaker has demonstrated that group A streptococci produce three different DNases (A, B, and C) which can be distinguished by their electrophoretic and immunological properties and by their pH optima ( 4 6 ) . The relative amount of each enzyme produced appears to be a function of strain variation. Streptodornase has been identified with DNase A. Lacks and Greenberg havc partially purified an endonuclease from Diplococcus pneumoniae in conjunction with the exonuclease cited in Section II,A (10). This enzyme is active on both native and denatured DNA and produces 5’-phosphoryl-terminated olingonucleotides. 3. Phage TS-Induced DNase

A deoxyribonuclease not normally present in E . coli is rapidly synthesized after infection with phage T5 (4‘7‘). This DNase appears at approximately the same time as the other “early” phage-specific enzymes (DNA polymerase and deoxynucleotide kinase, etc.) induced following infection with this bacteriophage. The T5-induced DNase has been purified nearly to the point of physical homogeneity. It has a pH optimum of pH 9.3 in glycine buffer and absolutely requires a divalent cation (magnesium or manganese) for activity. The mode of attack by the enzyme appears to be both endonucleolytic and exonucleolytic, yielding ultimately a mixture of 5’phosphoryl-terminated mononucleotides and oligonucleotides of average chain length 6 5 . Although the enzyme attacks both native and denatured DNA a t approximately the same rate, the proportion of mononucleotides produced from denatured DNA at the limit of digestion is only about one-fourth that found with native DNA. 4. M . luteus ATP-Dependent Endonuclease

In 1964, Tsuda and Strauss discovered a DNase activity in crude extracts of Micrococcus lysodeikticus (later renamed Micrococcus luteus) which required a nucleoside di- or triphosphate for activity ( 4 8 ) . This enzyme has recently been purified extensively (2400-fold) and examined in detail by Takagi and his colleagues (49). It has an alkaline pH L. W. Wannamaker, J . Exptl. M e d . 107, 797 (1958). A. V . Paul and I. R. Lehman. JBC 241, 3441 (1966). Y. Tsuda and B. S. Strauss, Biocl~emistry3, 1678 (1964). M . Anai. T. Hirahashi, and Y. Tukagi, JBC 245, 767 (1970); M. Anai, T. Hirahashi, M. Yamanaka, and Y. Tnkagi, ibid. p. 775. 46. 47. 48. 49.

262

I. R. LEHMAN

optimum (pH 9.4) and requires a divalent cation, preferably magnesium ion for activity. Double-stranded DNA is degraded a t a rate 40-fold greater than denatured DNA. The mode of attack is endonucleolytic yielding 5’-phosphoryl-terminated oligonucleotides with an average chain length of 5.5 residues a t the limit of digestion. A kinetic analysis of DNA degradation by the M . luteus endonuclease by means of sucrose density gradient centrifugation of the products as hydrolysis proceeded showed that the DNA added initially disappeared and was replaced by very slowly sedimenting material. Products of intermediate size were not detectable. Takagi and his colleagues interpreted these findings in terms of a one-by-one type of degradation in which the products of the initial hydrolysis possess a higher affinity for the enzyme than the undegraded DNA substrate, with the result that a DNA molecule initially attacked is degraded to completion before attack of a second molecule is begun. The role of the nucleoside triphosphate in the hydrolysis of DNA has not yet been clarified. ATP and dATP are the most effective nucleotides and only slight activity (10% or less) is observed with the other triphosphates ; nucleoside diphosphates are inactive. The rate of DNA hydrolysis is proportional to the ATP concentration and the ATP is converted to ADP and inorganic phosphate in the course of the reaction. Three moles of ATP are consumed for each phosphodiester bond cleaved, indicating a complex mechanism of participation of ATP in the endonucleolytic reaction. Preliminary experiments by Takagi and his colleagues indicate that the purified enzyme catalyzes an exchange of ADP with ATP in the absence of DNA, suggesting that a phospho enzyme may be an intermediate.

B. SPECIFIC ENDONUCLEASES During the past three years endonucleases have been discovered which possess a specificity which is considerably more refined than that shown by the nucleases considered thus far. Typically, these enzymes catalyze the cleavage of one or, a t most, a few phosphodiester bonds in a DNA molecule composed of many thousands of nucleotide residues. In no instance has the basis for this remarkable specificity been established. However, in the case of the E . coli “restriction” enzymes the presence or absence of a methyl group on a specific deoxyadenylate or deoxycytidylate residue may be involved. The limited attack catalyzed by the specific endonucleases obviously poses formidable assay problems which have in several cases been re-

10. BACTERIAL

DEOXYRIBONUCLEASES

263

solved by novel and ingenious methods. These are indicated where applicable. 1. E . coli Restriction Endonucleases

The E . coli restriction endonucleases are involved in the phenomenon of host-controlled modification and restriction whereby bacterial cells of one strain are able to destroy DNA from cells of foreign strains (50). As noted above, the current working hypothesis is that resistance to a given restriction endonuclease is conferred by specific methylation a t nucleotide sequences that would otherwise be vulnerable to that enzyme. The restriction enzymes have been assayed either by measuring the decrease in sedimentation coefficient of a homogeneous DNA preparation (usually phage A) (51) or more simply by measuring the inactivation of infectious DNA (phage h or fd) ( 5 2 ) ,both of which are sensitive to the introduction of one or a few phosphodiester bond scissions. One of the restriction endonucleases, called endonuclease R -K, has been purified approximately 5000-fold from E . coli strain K by Yuan and Meselson (61). This enzyme cleaves double-stranded DNA synthesized in other E . coli strains but is totally without effect on DNA synthesized in strain K itself. I n addition to magnesium ion the enzyme specifically requires ATP and S-adenosylmethionine; its pH optimum lies in the range pH 7.5-8.0. Several very similar nucleases with different specificities but with the same unusual cofactor requirements have also been described. An enzyme partially purified from E . coli lysogenic for phage P1 attacks DNA from E . coli lacking P1 ( 5 1 ) , and another enzyme from E . coli strain B attacks DNA from strains other than B ( 5 2 ) . There has also been a brief report of a restriction endonuclease in Hemophilus influenzae (53). Meselson and Yuan have carried out a detailed examination of the mode of attack of X.C DNA (i.e., DNA from phage h grown on E . coli strain C and therefore lacking the K enzyme modification) by purified endonuclease R - K ( 5 1 ) . Sucrose density gradient analysis of the products formed after treatment of h. C DNA with the endonuclease has shown that they consist of duplexes containing little or no single-stranded DNA and no single chain breaks. The products sediment in sucrose gradients a t or near the position at which quarter molecules of A-DNA would sediment. Thus, the h .C DNA appears to undergo double-stranded 50. W. Arber and S. Linn, Ann. Rev. Biochem. 38, 467 (1969). 51. R . Yuan and M. Meselson, Nature 217, 1110 (1968). 52. S. Linn and W. Arber, Proc. Natl. Acad. Sci. U . S. 59, 1300 (1968). 53. T. J. Kelly, Jr. and H. 0. Smith, Federation Proc. 29, 405 (1970).

264

I. R. LEHMAN

cleavage at fixed sites along its length. By examining the action of the enzyme on the twisted circular form of X.C DNA (54),Meselson and Yuan have also been able to demonstrate that single-stranded scission precedes cleavage of the duplex. The occurrence of single chain scissions early in the reaction taken together with the paucity of single chain breaks in the limit product indicate that the enzyme first cleaves only one chain and then a few seconds later breaks the complementary chain a t a point directly or nearly opposite to the initial break. A similar result was obtained by Linn and Arbor with the restriction endonucleases partially purified from E . coli and E . coli lysogenic for P1 (62).It has not been determined whether a given enzyme molecule remains bound to the DNA, catalyzing breaks in both chains, or whether the two chains are attacked independently. I n the case of endonuclease R - K acting on phage X DNA it has been established that duplexes in which only one of the two chains is modified (presumably by the appropriate methylation) are not attacked at all. Thus, heteroduplexes are resistant even to single chain scissions and are therefore modified a t every site of potential attack. The role of ATP and S-adenosylmethionine in the reaction remains an intriguing but as yet unresolved question. Recently Yuan and Meselson have reported that in the presence of magnesium ion, ATP and S-adenosylmethionine the Re K endonuclease forms a specific complex with its DNA substrate ( 5 5 ) . Complex formation is, however, observed M ) a t which nucleolytic activity is a t ATP concentrations (4 x not detectable. This result suggests that ATP may be involved in a t least two steps: (1) formation of a nonhydrolytic complex a t low ATP levels and (2) formation of more stable (or more numerous) complexes and nucleolytic action a t higher concentrations of ATP. The S-adenosylmethionine requirement for complex formation is in the same concentration range as observed for restriction. 2. E . coli Endonuclease I1

Endonuclease I1 of E . coli was first recognized by Friedberg and Goldthwait as an activity in extracts of E . coli mutants lacking endonuclease I, that specifically attacked double-stranded DNA alkylated with the monofunctional alkylating agent methyl methane sulfonate ( 5 6 ) . It was subsequently found that the partially purified enzyme could in fact 54. V. C. Bode and A . D. Kaiser, J M B 14, 399 (1965). 55. R. Yuan and M. Meselson, Proc. Natl. Acad. Sci. U . S. 65, 357 (1970). 56. E. C. Friedberg and D. A. Goldthwait, Proc. Natl. Acad. Sci. I J . S . 62, 934

(1969); E. C. Friedberg. S. M. Hadi, and D. A . Goldthwait, JBC 244, 5879 (1969).

10.


265

introduce a limited number of single-stranded breaks in nonalkylated duplex DNA, approximately 3-4 per single strand. The activities on the alkylated and nonalkylated DNA appear to be associated with the same protein; however, this point has not yet been definitely settled. Endonuclease I1 has a broad pH optimum ranging from pH 8.0 to 9.0. It has no absolute requirement for a divalent cation but is stimulated by added magnesium or manganese ion. Unlike endonuclease I, i t is not inhibited by RNA. Goldthwait and his co-workers have assayed endonuclease I1 by measuring the release of 3H-thymidine-labeled fragments from a suspension of polyacrylamide gel containing alkylated T4 DNA ( 5 7 ) . Most (>90%) of the radioactivity released from the gel is acidprecipitable, hence, still macromolecular. With lightly alkylated DNA, endonuclease I1 makes predominantly single-stranded breaks suggesting that the enzyme can hydrolyze a phosphodiester bond a t or near an alkylated base in a native DNA molecule with no single-stranded breaks in this region. With more extensively alkylated DNA, double-stranded breaks predominate. Friedberg et al. (56) pointed out that since alkylation of DNA occurs principally a t the N-7 position of guanine and the N-3 position of adenine (58) these chemical modifications might be expected to result in electron rearrangements in the purine rings that may affect hydrogen bonding, base-stacking, or both. Thus, alkylation might produce a distortion in the secondary structure of DNA, and it is this distorted structure which is the substrate for endonuclease 11. Since the enzyme makes a limited number of single-stranded breaks in native DNA, it is possible that a similar type of conformational distortion exists in nonalkylated DNA. The role of endonuclease I1 in vivo is not known. Assay of recombination defective mutants of E . coli and mutants abnormally sensitive to ultraviolet irradiation and to treatment with methyl methane sulfonate showed them all to possess normal levels of the enzyme (56).

3. Phage T7-Induced Endonuclease Center e t al. (59) have purified extensively (1000-fold) an endonuclease induced after infection of E. coli with phage T7. This enzyme attacks native DNA to yield products with a molecular weight of approximately 2 X lo4 and denatured DNA to produce fragments having 57. E. Melgur and D. A. Goldthwait, JBC 243, 4401 (1968). 58. P. D. Lawley, Prog. Nucleic Acid Res. Mol. Biol. 5, 89 (1966). 59. M. S. Center, F. W. Studier, and C. C. Richardson., Proc. Natl. Acad. Sci. U.S. 65, 242 (1970).

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a molecular weight of lo4 or less. The fragments have not been further characterized. The endonuclease activity on native DNA was assayed by measuring the conversion of 3H-labeled 4X-174 RF I (60)to a form which could be trapped on nitrocellulose membrane filters after heating at 100". Thus, covalently closed duplex DNA is not retained on these filters, whereas the single strands produced by heating the circular duplexes that have suffered a single-stranded break are retained. The action of the enzyme on single-stranded DNA was followed by measuring the conversion of single-stranded circular +X 174 DNA (61) to a form susceptible to E. coli exonuclease I (62). Studies with conditional lethal mutants of phage T 7 have shown that gene 3 is the structural gene for the DNase (T7 contains at least 19 distinct complementation groups) (63).In the restrictive host, gene 3 mutants synthesize only limited amounts of phage DNA ( 6 4 ) . Inasmuch as they are also defective in carrying out the degradation of host DNA, the gene 3 endonuclease may be the enzyme required for this function. The purified endonuclease produces little acid-soluble material ; thus, an additional activity is presumably required for the complete degradation of host DNA. Center et al. suggested that gene 6 specifies an exonuclease which acts a t the breaks produced by the gene 3 endonuclease (69). As noted above amber mutants in gene 3 produce only limited amounts of DNA under restrictive conditions. It is therefore clear that degradation of host DNA is required for normal T7 DNA synthesis and phage production, a result which is to be anticipated in light of the finding that T7 derives most of its nucleotides from host nucleic acids ( 6 5 ) . Center et al. have observed that the purified T 7 endonuclease can hydrolyze T 7 DNA in in vitro (59). Hence, it is not known how T 7 infection results in the selective degradation of E . coli DNA in vivo. 4. Phage T4-Induced Endonucleases

II and IV

Sadowski and Hurwitz have described two endonucleases synthesized in T 4 phage-infected E . coli which they have named T4 endonucleases II and ZV (66). 60. A. Burton and R. L. Sinsheimer, JMB 11, 327 (1965). 61. W. Fiers and R. L. Sinsheimer, J M B 5, 424 (1962). 62. I. R. Lehman and A. L. Nussbaum, JBC 239, 2628 (1964). 63. F. W. Studier and J. V. Maisel, Virology 39, 575 (1969). 64. F. W. Studier, Virology 39, 562 (1969). 65. F. W. Putnam, D. Miller, L. Palm, and E. A. Evans, Jr., JBC 199, 177 (1952). 66. P. D. Sadowski and J. Hurwitz, JBC 244, 6182 and 6192 (1969).

10.


267

a. T4 Endonuclease II. T4 endonuclease I1 has been purified approximately 300-fold (6‘6‘). The partially purified enzyme has a broad alkaline p H optimum (pH 8.4-10.1) and shows an absolute requirement for magnesium ion. Two methods have been used for the assay of the enzyme. One measures the discharge of 3H-AMP from the E . coli DNA ligase-AMP complex on reaction of the complex with DNA pretreated with endonuclease 11. Since ligase-AMP repairs single-stranded breaks in duplex DNA that have apposing 3’-hydroxyl and 5’-phosphoryl termini (67), the discharge of AMP is a direct measure of the number of breaks introduced into the DNA by T4 endonuclease 11. The second assay estimates the number of 3’-hydroxyl termini formed by measuring the increase in I4C-labeled h DNA which beconies susceptible to E . coli exonuclease I after thermal denaturation of the nicked DNA. Since exonuclease I acts exonucleolytically from the 3’-hydroxyl terminus of denatured DNA ( 6 2 ) ,the amount of acid-soluble 14C formed is an estimate of the number of 3’-hydroxyl termini generated by T4 endonuclease I1 action. The enzyme introduces predominantly single-stranded breaks into native DNA, although at high enzyme concentrations some doublestranded breakage occurs. Purified preparations of the endonuclease do, however, show some activity on denatured DNA, possibly resulting from contamination with T4 endonuclease I V (see below). As noted above, the single-stranded breaks produced by T4 endonuclease 11 bear 3’-hydroxyl and 5’-phosphoryl termini. The enzyme makes a limited number of breaks in duplex DNA and the average length of the limit product from phage h DNA, as determined by sucrose density gradient centrifugation, is about 1000 residues. As in the case of E. coli endonuclease 11, the basis for the very limited extent of hydrolysis of DNA is not known. Sadowski and Hurwitz have examined the nucleotide residues a t the 5’-phosphoryl termini by means of the polynucleotide kinase reaction and found all four deoxynucleotides to be present. There did, however, appear to be a significantly higher proportion of deoxyguanylate and deoxycytidylate residues a t these termini. A limited base specificity of this kind cannot, however, account for the apparent high degree of specificity actually observed. T4 endonuclease I1 appears to differ from E . coli endonuclease I1 in several respects, the most striking difference being the inability of the phage-induced enzyme to attack either glucosylated or nonglucosylated T4 DNA. Friedberg et al. have also found that whereas the activity of 67. J. R. Little, S. B. Zimmerman, C. K. Osliinsky, and M. Gellert, Proc. Natl. Acad. Sci. U . S. 58, 2004 (1967).

268

I. R. LEHMAN

extracts of infected and uninfected cells showed similar activities when tested with alkylated DNA, there was a marked increase in relative endonucleolytic activity on unalkylated DNA in the infected cell extract, suggesting that the phage-induced enzyme either has no activity or very little activity on alkylated DNA (56). Recently, two groups have independently isolated non-lethal mutants of T4 that are incapable of degrading the host E. coli DNA and may therefore be defective in T4 endonuclease I1 (68, 69).

b. T4 Endonuclease I V . T4 endonuclease IV has been purified approximately 150-fold (66). It has a p H optimum which ranges from pH 8.4 to 9.2 and an absolute requirement for magnesium or cobalt ion. Endonuclease IV was assayed by measuring the conversion of singlestranded circular fd DNA ( 7 0 ) , labelcd with 14C to an exonuclease Isensitive form. The partially purified enzyme has a strong preference for denatured DNA; however, it does attack native X DNA a t a significant rate (one-tenth that of fd). Like T4 endonuclease 11, endonuclease IV shows only limited activity, producing oligonucleotides 150 residues long; it does not form measurable amounts of acid-soluble material. The oligonucleotide products contain 3’-hydroxyl and 5’-phosphoryl termini; the latter bear deoxycytidylate residues exclusively. T 4 endonuclease IV therefore has an absolute base specificity and is unique among the DNA endonucleases which have thus far been described. Some other, as yet undetermined feature of its mechanism must, however, be responsible for the limited extent of attack observed. Again, like T4 endonuclease I1 it does not hydrolyze T 4 DNA, whether glucosylated or not. The combined action of T 4 endonucleases I1 and IV and an exonuclease which Sadowski and Hurwitz have identified in extracts of T4-infected cells can result in the double-stranded breakage of duplex DNA. Presumably, endonuclease I1 introduces a single-stranded break in native DNA and the exonuclease removes mononucleotides from the internal 3’-hydroxyl termini exposing a region of single-stranded DNA on the opposite strand. This region is then cleaved by endonuclease IV to yield double-stranded fragments of DNA. Because of the inability of T4 endonucleases I1 and IV to attack T 4 DNA, it is possible that these enzymes together with the “T4 exonuclease” promote the initial stages 68. K . Hercules, J. L. Munro, S. Mendelsohn, and J. S. Wiberg, Federation Proc. 29, 465 (1970). 69. S . E. Jorgensen, J. F. Koerner, D. P. Snustad, and H. R. Warner, Federalion Proc. 29, 465 (1970). 70. H. Schaller, JMB 28, 435 (1969).

10.


269

of breakdown of host DNA following T4 infection. Warren and Bose have, in fact, reported that in the initial phase of host DNA breakdown, E . coli DNA is degraded to fragments of molecular weight approximately 1 X lo6 in a process in which single-stranded breaks occur first, followed by double-stranded cleavage ( 7 1 ) . The double-stranded DNA fragments are then presumably degraded to acid-soluble oligonucleotides and ultimately mononucleotides, which can then be used in the synthesis of T4 DNA. 5. M . luteus “UV Repair Enzymes” I n 1962, Strauss described an activity in extracts of Micrococcus lysodeikticus ( M . luteus) which preferentially inactivated UV-irradiated B . subtilis transforming DNA ( 7 2 ) .It was subsequently shown by Carrier and Setlow that such extracts were able specifically to excise thyminethymine and thymine-cytosine dimers from the irradiated DNA ( 7 3 ) . Fractionation of the M . luteus extracts by Nakayama et al. demonstrated that two chromatographically separable protein fractions were required for pyrimidine dimer excision, one of which they suggested was an endonuclease that introduced a phosphodiester bond cleavage a t or near the pyrimidine dimer and the second of which released the dimer as part of an acid-soluble oligonucleotide ( 7 4 ) . The two activities have been purified and examined in detail by Takagi et al. (75) and independently by Grossman and his colleagues (32, 7 6 ) .The purified enzymes acting together do in fact have the capacity to excise quantitatively thymine dimers from UV-irradiated DNA. The first enzyme in the sequence has been purified approximately 5000-fold; it is of relatively low molecular weight (14,0W15,OOo based on Sephadex gel filtration). The pH optimum of the enzyme is pH 6.5-7.5 and it is stimulated by, but it is not dependent upon, added magnesium ion. The purified enzyme is entirely specific for UV-irradiated, double-stranded DNA and is free of the ATP-dependent endonuclease of M . luteus. The UV endonuclease introduces a single-stranded break in close proximity t o a 71. R. J. Warren and S. K. Bose, J . Viral. 2, 327 (1968). 72. B. S. Strauss, Proc. Natl. Acad. Sci. U . S. 48, 1670 (1962). 73. W. L. Carrier and R. B. Setlow, B B A 129, 318 (1966). 74. H. S. Nakayama, M. Okubo, M. Sekiguchi, and Y. Takagi, BBRC 27, 217 (1967). 75. Y. Takagi, M. Sekiguchi, S. Okubo, H. Nakayama, K. Shimada, S. Yasuda, T. Nishimoto, and H. Yoshihara, Cold Sprlng Harbor Symp. Quant. Biol. 38, 219 (1968). 76. J . C. Kaplan, S. R. Kushner, and L. Grossman, Proc. Natl. Acad. Sci. U . S. 63, 144 (1969).

270

I. B. LEHMAN

thymine dimer leaving a 3’-phosphoryl terminus. Nucleotides are not released during this incision step. The subsequent thymine dimer excision is carried out by the second enzyme, an exonuclease also purified extensively (15OO-fold) by Grossman et a2. This enzyme, whose activity is dependent upon added magnesium ion, acts on unirradiated denatured DNA releasing 5’-mononucleotides by an exonucleolytic mechanism starting a t either the 3’ or 5’ terminus. Native DNA is resistant to the action of the exonuclease; however, the enzyme will attack irradiated native DNA that has been pretreated with the endonuclease in the 5’+ 3’ direction liberating an average of 6 nucleotides per endonucleolytic break. The digestion products consist of mono-, di-, and trinucleotides ; and the thymine dimers are contained in the trinucleotide fragments. Purified preparations of the exonuclease are devoid of DNA polymerase activity, suggesting that the UV exonuclease is not a 5’ + 3’ exonuclease component of the M . luteus DNA polymerase. It would appear from these studies that the excision of thymine, and more generally pyrimidine dimers may be a two-step process. The initial single-stranded incision is probably dependent upon the presence of a distorted area in the DNA duplex resulting from the formation of a thymine dimer. Once the initial break has been introduced, a short single-stranded region containing the photoproduct results which is then susceptible to the action of the exonuclease. Grossman et al. have isolated a mutant of M . luteus by nitrosoguanidine mutagenesis which is abnormally sensitive to UV and X-irradiation and also shows a reduced capacity to support the replication of UVirradiated phages (the hcr- phenotype). Extracts of this mutant have a correspondingly low level of UV endonuclease activity (76).Takagi et al. have transformed the mutant with DNA derived from UV-resistant cells and found that the UV-resistant transformants displayed the same level of sensitivity to UV irradiation as the wild type; however, the W endonuclease activity in the extract remained a t the same low level observed in the original mutant strain ( 7 5 ) . All attempts to isolate a revertant have thus far been unsuccessful, suggesting that the mutant bacterium may harbor a double mutation. Thus, while it appears that the UV endonuclease may be involved in the repair of UV damage in V ~ V O , this point has not been firmly established.

Sfi1een A cid Deoxyribonuclease GIORGIO BERNARD1 I. Introduction . . . . . . . . . . . . 11. Physical and Chemical Properties . . . . . . . A. Isolation . . . . . . . . . . . B. Physical and Chemical Properties . . . . . . C. A and B Components . . . . . . . . D. Dimcric Structure . . . . . . . . . 111. Catalytic Properties . . . . . . . . , . . A. General Features of DNA Dcgradation . . . . B. Methods of Investigation and Activity Units . . . C. Mechanism of the Initial Degradation of Native D S A . D. General Catalytic Properties . . . . . . . E. Specificity . . . . . . . . . . . 1V. Distribution, Intracellular Localization, and Biological Hole .

.

. . .

. . . . . . . . .

271 272 272 273 275 275 276 276 278 278 280 283 285

1. Introduction

Acid deoxyribonuclease (DNase) is an enzyme which splits the phosphodiester bonds of native DNA by both a diplotomic and a haplotomic mechanism (see Section II1,C) leaving the terminal phosphate in a 3’ position. The enzyme is very widely distributed in animal cells and appears to be localized in the lysosomes. The best known acid DNase is that from hog spleen; this explains why most of the data presented here refer to this enzyme. I t should be stressed, however, that the properties of acid DNases obtained from the tissues of other vertebrates appear to be extremely similar to those of the hog spleen enzyme; 271

272

GIORGIO BERNARD1

therefore, the results obtained with hog spleen acid DNase may be of a more general validity. The present article will review the progress made in our knowledge of acid DNase during the past 10 years, therefore after the second edition of “The Enzymes” (1). No attempt has been made to cover exhaustively the literature on this subject since several reviews dealing with acid DNase have been published in the meantime ( 2 4 ) . The DNase activity having a pH optimum comprised between 4.5 and 5.5, first observed in animal tissues in the late 1940’s, was referred to by the early investigators as acid DNase. Later the name DNase 11, intended to mean the second type of animal DNase was suggested for the acid DNase activity (7) to contrast it with pancreatic DNase, which was called DNase I. Subsequently, DNases have been classified as 5’-monoester formers and 3’-monoester formers (I), a division which is not identical with that of DNases I and 11, even if frequently it is assumed to be so. More recently, it has been suggested to abandon the distinction between DNase I and DNase I1 altogether ( 6 ) . Here the original terminology will be used since it is the least inconsistent among those proposed so far.

II. Physical and Chemical Properties

A. ISOLATION Methods leading to homogeneous acid DNase preparations from hog spleen have been described (8-11). The following is a very brief outline of the method of Bernardi et al. (10) as presently used in the author’s laboratory. Hog spleens are trimmed, ground, and homogenized with 0.05M H,SO,; the homogenate is acidified to pH 2.5 with 0.1M H,SO, and centrifuged; the supernatant so obtained is fractionated be1. M. Laskowski, “The Enzymes,” 2nd ed., Vol. 5, p. 123, 1961. 2. M. Privat de Garilhe, “Les nucl6ases.” Hermann, Paris, 1964. 3. I. R. Lehman, Ann. R e v . Biochem. 36, 645 (1967). 4. W. E. Razzell, Experientia 23, 321 (1967). 5. M. Laskowski, Advan. Enzymol. 29, 165 (1967). 6. G. Bernardi, Advan. EnzymoZ. 31, 1 (1968). 7. L. Cunningham and M. Laskowski, BBA 11, 590 (1953). 8. G. Bernardi, M. Griff6, and E. Appella, Nature 198, 186 (1963). 9. G. Bernardi, and M. Griff6, Biochemistry 3, 1419 (1964). 10. G. Bernardi, A. Bernardi, and A. Chersi, BBA 129, 1 (1966). 11. G . Bernardi, Procedures Nucleic Acid Res. p. 102 (1966).

11.

273

SPLEEN ACID DEOXYRIBONUCLEASE

tween 40 and 80% saturation of (NH,),SO,; the final precipitate is dialyzed against 0.05M phosphate buffer, p H 6.8, and clarified by centrifugation. The crude enzyme solution so obtained is purified using three chromatographic steps involving DEAE-Sephadex, hydroxyapatite, and CM-Sephadex, respectively. The enzyme is eluted from the last column in two activity peaks, the first component representing less than 20% of the total activity. The two components, called A and B, respectively, are rechromatographed separately on CM-Sephadex columns, dialyzed against 0.001 M acetate buffer, pH 5.0, concentrated by freezedrying to about 1% concentration, then frozen and stored a t -15". Hog spleen acid DNase, as obtained by the above procedure, is completely free of contaminating phosphatase, exonuclease, and adenosine deaminase activities. The enzyme has a weak intrinsic hydrolytic activity on bis (p-nitrophenyl) phosphate and the p-nitrophenyl derivatives of deoxyribonucleoside 3'-phosphates (see Section III,D,3).

B. PHYSICAL AND CHEMICAL PROPERTIES The physical properties and the amino acid analysis of hog-spleen acid DNase B (main component) are given in Tables I and 11, respectively. The high ammonia level of the acid hydrolyzate and the high amide level seen in Pronase digests suggest that a very large percentage of t,he dicarboxylic acids may be present in the protein as the corresponding amides. Glucosamine ( l a ) and mannose (R. G . Winzler, personal TABLE I ACID DEOXYRIBONUCLEASE" PHYSICAL PROPERTIES OF HOQ SPLEEN (Svedbergs) (analytical centrifugation) (sucrose gradient centrifugation) D20,w(lo-' cmz/sec)b P (ml/g)c Molecular weight (from s and D ) (from sedimentation equi1ibrium)d ~ 2 0 , ~

fEiZ if0 ip

3.4 3.3 0.1 7.8

*

0.72 3.8 x 104 4.1 X 10' 1.34 12.1

From Bernardi el al. (12).

* This value was obtained a t concentrations of 0.5Yo and about 0.1%. c

d

Value calculated from amino acid composition. Townend and Bernardi (2s).

12. G. Bernardi, E. Appella, and R. Zito, Biochemistry 4, 1725 (1965). 13. R. Townend and G. Bernardi, ABB, submitted for publication.

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GIORGIO BERNARD1

TABLE I1 AMINOACID ANALYSISOF HOG SPLEENDEOXYRIBONUCLEASE~ Grams of amino acid residues per 100 g of protein* in hydrolysis time of Amino acid

22 hr

48 hr

72 hr

Coriected values"

LYS His (NHs) -4% ASP Thr Ser Glu PI.0 GlY Ma +-cys Val Met Ileu Leu TYr Phe Try Glucosamine

6.94 2.13 (23.05) 5.04 10.19 5.44 8.30 11.03 7.09 3.88 4.84 1.77 2.62 1.63 2.08 10.47 5.23 6.30 6.30 3.07

6.80 2.17 (23.65) 5.55 10.14 5.20 8.10 11.09 7.14 3.97 4.72 1.79 3.33 1.65 2.49 11.13 5.17 6.56

6.95 2.31 (24.06) 5.37 9.36 4.80 7.18 10.66 6.51 3.87 4.33 1.79 3.35 1.30 2.52 10.49 4.83 6.07

6.89 2.20 (22.5) 5.46 9.89 5.63 8.46 10.93 6.91 3.90 4.63 1.79 3.35 1.53 2.52 10.67 5.28 6.31

3.32

3.06

3.15

98.05 98.9

100.32 98.7

94.75 97.4

99.50

Total

N recovery (%)

Moles of amino acid/ mole of protein MW 38,000

Nearest integral No. of residues/ mole of proteind

20.44 6.08 (49.4) 13.30 32.68 21.16 36.93 32.19 27.05 25.99 24.77 6.65" 12.84 4.45 8.47 35.87 12.31 16.30 6.3' 7.45

20 6 (49) 13 33 21 37 32 27 26 25 8 13 4 8 36 12 16 6 (8) 343

From Bernardi et al. (fa). Total N is 17.2%; total S is 1.0%. c In calculating the corrected values, the criteria given by Tristram and Smith (14) have been followed. dThe selection of the integral numbers of residues has been done also taking into account results from other analyses. After performic acid oxidation, cysteic acid 8.2 residues. From N-bromosuccinimide titration.

communication, 1967) are present in the protein; in the tryptic digests, the glucosamine and neutral sugar residues are found on a single peptide spot. No free sulfhydryl groups can be detected in acid DNase, both native and denatured (G. Bernardi, unpublished observations) ; therefore, the eight half-cystine residues must form four disulfide bridges. Optical rotatory dispersion, circular dichroism, and infrared spectra of

11.


275

acid DNase have shown that the enzyme contains little a-helix; antiparallel pleated sheet p-structure is probably present in the molecule (13a). C. A

AND

B COMPONENTS

A comparison of the properties of the two acid DNase components A and B showed no differences in the sedimentation velocities, elution volumes from Sephadex G-100 columns, ultraviolet spectra, orcinol reaction, and enzymological properties (DNase and phosphodiesterase activities). Two differences (besides their different behavior on CMSephadex columns, which indicates that component A is likely to be slightly less basic than component B) have been found so far between the two components: (1) the level of both glucosamine and mannose are definitely lower in component A compared to component B (R. G. Winzler, personal communication, 1967) ; (2) one particular single tryptic peptide spot of component A is resolved into two spots in the otherwise identical peptide map of component B. Since the amount of component A may be reduced to zero by avoiding acidification, the latter finding may tentatively be explained (10) by assuming that the difference between A and B results from the deamidation of an asparagine (or a glutamine) residue adjacent to a lysine (or an arginine) by the acid treatment involved during the preparation of the enzyme, leading to a peptide bond resistant to trypsin. Alternatively, it may be thought that during acidification a peptide bond has been split by cathepsins present in the homogenate. The loss of sugars might also be due to acid hydrolysis or to an enzyme attack. It is possible that the two chromatographic components of spleen acid DNase seen by Koszalka et al. (16) on Amberlite IRC-W had an origin similar to components A and B, since 5 N H,SO, was used by these authors to adjust the tissue extract to p H 4.0.

D. DIMERIC STRUCTURE Some chemical and physical results seem to suggest that acid DNase may have a dimeric structure (6, 16). When the enzyme is reduced, carboxymethylated, digested with crystalline trypsin (treated with 1,l13a. S. N . ‘rirnasheff and G. Bernardi. ABB 141, 53 (1970).

14. G. R. Tristram and R. H. Smith, Advan. Protein Chem. 18, 227 (1967). 15. T . R. Kosaalka, R. Falkenheim, and K. I. Altman, BBA 33, 647 (1957) 16. G. Bernardi. J M B 13, 603 (1965).

276

GIORGIO BERNARD1

tosylamido-2-phenethyl-chloromethylketone to inactivate contaminating chymotrypsin), and mapped, 17-19 peptides are found as opposed to 32-34 arginine lysine residues present in each enzyme molecule of MW = 38,000. Arginine, tryptophan, and histidine peptides are found in half, or less than half, the number of the respective amino acids in the supposedly dimeric protein. Other results also suggesting a dimeric Structure come from sedimentation studies. At a 1% concentration, the sedimentation coefficient of the enzyme, which is 2 . 8 s in acetate or phosphate buffers, drops to 1.75 and 2.1 S in 6 M guanidine a t pH 5.6 and 8.6, respectively; in the presence of P-mercaptoethanol, the sedimentation coefficient is still lower, as expected: 1.5 S in 6 M guanidine and only 0.8s in 8 M urea; in 4 M u r e a 4 0 5 M P-mercaptoethanol, pH 5.0, two boundaries, having sedimentation coefficients equal to 2.6 and 0.8 S, respectively, can be seen. Cooperative binding of the synthetic substrate bis (p-nitrophenyl) phosphate and of a protein inhibitor (see Sections III,D,2 and 3) might also be considered as an indirect indication of a dimeric structure. Recent equilibrium sedimentation studies (13) show, however, that the molecular weight of acid DNase in 6 M guanidine (with or without 0.1 M P-mercaptoethanol) is close to 40,000. Clearly, further investigations on acid DNase are needed for a better understanding of the relationships between structure and mechanism of action of this enzyme.

+

111. Catalytic Properties

A. GENERAL FEATURES OF DNA DEGRADATION Three different phases can be distinguished in the degradation of native DNA by acid DNase as follows (see Fig. 1) : (1) The initial phase, in which the macromolecular and biological properties of DNA are dramatically modified, whereas no change can be detected in its spectral properties and no acid-soluble fragments are formed. I n terms of molecular weight this phase extends from the initial molecular weight to about 1 0 daltons [weight average molecular weight, M,, of double-stranded fragments (17,18)1. (2) The middle phase, which is characterized by a hyperchromic shift and the formation of acid-soluble oligonucleotides ; monoesterified phosphate can be detected. In this phase, the increase of ultraviolet absorption and of acid-soluble oligonucleotides is linear with the reciprocal 17. G. Bernard, Nature 206, 779 (1965). 18. G. Bernardi, BBA 174, 423 (1969).

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SPLEEN ACID DWXYRIBONUCLEASE

i

1.35

-0.1I

10-0.10

-0.09 -0.08 -0.07

Time (minutes)

FIG.1. Hyperchromic shifts (0; left-hand outer scale), acid solubility ( A ; lefthand inner scale; values corrected for dilution), reciprocal average size (0 ; righthand outer scale) of oligonucleotides from calf thymus DNA during acid DNase digestion. The DNA sample ( A m 0 = 8.0) was digested at 23" in 0.05 M ammonium acetate401 M EDTA, pH 5.5. The horizontal broken line indicates the limit between the middle phase and the terminal phase of the digestion. I n this experiment the initial lag time of both hyperchromic shift and acid solubility is not apparent because of the high enzyme concentration used ( 1 9 ) .

number average molecular weight of oligonucleotides, l/Mn, that is the relative number of terminal nucleotides. Under the experimental conditions of Fig. 1, the average chain length in this phase is comprised between 100 and 14 nucleotides (19). It is important to stress that the limits of the middle phase may be shifted by changes in the experimental conditions of the digestion (temperature, pH, and ionic strength of the solvent). (3) The terminal phase, which shows an increasingly slower, further increase in the hyperchromic shift and acid-soluble oligonucleotide formation. Both phenomena, however, are no more linear with number of 19. C . Soave, J. P. Thiery, S. D. Ehrlich, and G . Bernardi, in preparation.

278

GIORGIO BERNARD1

end groups liberated. Under the experimental conditions of Fig. 1 this phase extends from a chain length of 14 to a chain length of 6 .

B. METHODS OF INVESTIGATION AND ACTIVITY UNITS The degradation of native DNA by acid DNase may be investigated, by physical, biological, and chemical methods. It should be pointed out that none of these methods can be used to follow the entire course of the enzymic degradation and also that all methods, with the only exception of the terminal phosphate determination, are indirect methods. The routine use of indirect assay procedures (most commonly the formation of acid-soluble oligonucleotides, the hyperchromic shift, or the viscosity drop) has made it very difficult to define a satisfactory activity unit. Since a linear relationship exists between the formation of terminal phosphate groups and both hyperchromic shift and the liberation of acid-soluble oligonucleotides (Fig. l ) , the indirect methods can be standardized against the direct method. It is possible, therefore, to define acid DNase activity according to the recommendations of the Commission on Enzymes of the International Union of Biochemistry: One unit of DNase is defined as the amount of enzyme which catalyzes the formation of 1 pmole of terminal phosphate per minute a t 25” under optimal conditions. One such unit is equivalent to about 325 units defined ( S l l ) as the amount of enzyme catalyzing the liberation of oligonucleotides having a corrected A,,, equal to 1, a t 37”, in 0.15 M acetate buffer, 0.01 M ethylenediaminetetraacetate (EDTA), p H 5.0, the DNA concentration being 400 pg/ ml.

C. MECHANISM OF

THE

INITIAL DEGRADATION OF NATIVEDNA

Acid DNase initially degrades native, double-stranded DNA according to two mechanisms (see Fig. 2) : (1) a diplotomic mechanism (8, 20)-

FIG.2. Scheme of the mixed haplotomic and diplotomic mechanism of degradation of native DNA by acid DNase. 20. The introduction of this terminology (6) is justified by the fact that a mechanism of DNA degradation involving the simultaneous breakage of both strands a t the same level shows a “single hit” kinetics whereas that caused by single breaks shows a “double hit” kinetics; this is, of course, a source of ambiguity.

11. SPLEEN

ACID DEDXYRIBONUCLEASE

279

from the Greek diplo's, double, and tome', break-by which both strands are simultaneously split at the same level, and (2) a haplotomic mechanism (6)-from the Greek hapZo's, single-causing scissions on one or another of the two strands. Both mechanisms are operational from the very beginning of the digestion. The diplotomic mechanism, first suggested on the basis of qualitative evidence ( d l ) ,has been rigorously established by kinetic work (22-25) and later confirmed using different experimental approaches (26, 2 7 ) . This mechanism is responsible for the absence of a time lag in the molecular weight decrease and for the linearity of plots of l J M , vs. digestion time, which characterize the acid DNase digestion. A diplotomic degradation of native DNA has subsequently been demonstrated to take place also with other DNases, e.g., E . coli endonuclease I (28, 29) and D.pneumoniae DNase ( S O ) . In spite of the fact that kinetically these enzymes degrade native DNA like acid DNase, i t is conceivable that their mechanism of action at the molecular level is different. Among other differences, the two bacterial enzymes have molecular weights close to half the molecular weight of acid DNase and form 3'-OH-ended oligonucleotides. The haplotomic mechanism is similar to that already known to occur with pancreatic DNase (31, 32) and contributes to the molecular weight decrease only after a time lag, during which single breaks accumulate on the DNA strands. The ratio of total bonds broken to bonds broken by the diplotomic mechanism has been estimated, in different ways, to lie a t least initially between 1.5 and 3 ( 3 3 ) . As can be expected, the initial degradation of DNA by acid DNase not only causes a drastic change in the macromolecular properties of the substrate, but also strongly affects its biological activity. Young and Sinsheimer (26') have been able to show that close to one diplotomic 21. A. Oth, E. Fredericq, and R. Hacha, B B A 29, 287 (1958). 22. G. Bernardi and C. Sadron, Nature 191, 809 (1961). 23. G. Bernardi and C . Sadron, A . Baselli Conference on Nucleic Acids and Their Role in Biol., Milan, 1964 p. 62. 24. G. Bernardi and C . Sadron, Biochemistry 3, 1411 (1964). 25. L. A. MacHattie, G. Bernardi, and C . A. Thomas, Jr., Science 141, 59 (1963). 26. E. T. Young, I1 and R. L. Sinsheimer, JBC 240, (1965). 27. E. Melgar and D. A. Goldthwait, JBC 243, 4401 (1968). 28. G. Bernardi and C. Cordonnicr, J M B 11, 141 (1965). 29. F. W. Studier, JBM 11, 373 (1965). 30. H. Kopecka and G. Bernardi, 6th Meeting Fed. European Biochem. Sac., Mndrid, 1969 (abstracts). 31. C. A. Thomas, JAPS 78, 1861 (1956). 32. V. N . Schumaker, E. G. Richards, and H. K. Schachnian, JACS 78, 4320 (1960). 33. G. Bernardi and M. L. Bach, J M B 37, 87 (1968).

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break per A-DNA molecule is sufficient to destroy its infectivity, whereas, on the average, four phosphodiester bonds can be hydrolyzed by pancreatic DNase in a A-DNA molecule before its infectivity is lost. Bernardi and Bach (33) have found no feature in the inactivation of transforming H. influenzae DNA that could specifically be related to the diplotomic mechanism of action, in agreement with the fact that transformation occurs by integration of single-stranded and not of doublestranded DNA into the host genome. An interesting finding was that a t comparable levels of bond breakage acid DNase is much more inactivating than pancreatic DNase, E . coli, endonuclease I, or sonication; for example, a Poisson average of one inactivating event per cathomycin marker (37% survival) requires more than 50 breaks per molecule of 12 X lo6 daltons by pancreatic DNase but less than 2-4 breaks by acid DNase.

CATALYTIC PROPERTIES D. GENERAL These have generally been investigated by hyperchromic shift or acid solubility assays and therefore bear on the middle phase of the DNA degradation. 1. Effect of Substrate Concentration, p H , and Ions

When acid DNase activity is assayed by the acid-solubility method the optimal DNA concentration is 0.4 mg/ml (9) and higher substrate concentrations appear to be inhibitory (16, 21, 3 4 ) . It has been shown, however, that this inhibition is because increasing substrate concentration decreases the efficiency of acid-soluble oligonucleotide release since the number of breaks per unit length of DNA is lower. If a direct method of estimating enzymic activity is used, such as the determination of phosphatase-sensitive phosphate, it can be shown that the inhibition by high substrate seen by the acid solubility method is only apparent (34). The effect of pH and ions on acid DNase activity has been investigated in several laboratories, and rather different results have been reported. It appears now that many discrepancies result from a rather poor understanding of the complexity of pH and ion effects. In fact, it has been shown (34) that electrolytes and pH modify the acid DNase activity not only by affecting the enzyme itself but also by stabilizing or destabilizing the secondary structure of native DNA. Since the enzyme has a quite different affinity for the native vs. the denatured structure 34. R. Rosenbluth and S.-C. Sung, C u n . J . Biochem. 47, 1081 (1969)

11. SPLEEN

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28 1

of DNA (9),any change in the secondary structure of the substrate will indirectly affect the enzymic activity. Another complicating factor is the presence or absence of contaminating proteins. The effects of Mg2+ and SO:- are quite different a t different levels of enzyme purity (36). Using very highly purified hog spleen acid DNase a t p = 0.15 (9) the pH optimum is close to 4.8. At a 0.01 M level, Mg2+is slightly inhibitory above pH 4.5, whereas EDTA is an activator. Above pH 5.0, HPOf is slightly inhibitory and 5024- is very strongly inhibitory, particularly above pH 4.5. In succinate buffer, p = 0.15, pH 6.7, the activity is less than 3% of that in acetate buffer, pH 5.0, p = 0.15. At low ionic strength, acid DNase is active a t neutral pH (24,33, 36). Activation of acid DNase by cysteine was reported by Maver and Greco (37'), but was not found by Brown et al. (38).Bernardi and Griff6 (9) found an activating effect only on rather highly purified enzyme preparations. This effect was no longer apparent when protecting proteins were added to the enzyme solutions. Since acid DNase has no free sulfhydryl groups (see Section II,B), it is possible that cysteine protects the enzyme against traces of heavy metals. It should be mentioned that cytochrome c is particularly effective as a protecting protein. 2. Inhibitors

The occurrence of a dialyzable, heat-stable inhibitor in human urine has been reported (39, 40). Inhibition results from urinary sulfate (41) as well as from other salts ( 4 2 ) . Iodoacetic acid, N-bromosuccinimide, and H20, were found to be strongly inhibitory, whereas iodoacetamide was only slightly inhibitory and diisopropylfluorophosphate was not inhibitory. These results suggest that tryptophan, methionine, and/or histidine, but not serine, are involved in the enzymic activity ( 4 3 ) . Acid DNase is strongly inhibited by actinomycin D. In contrast with the claim (44) that actinomycin causes the same extent of inhibition of 35. C. Cordonnier and G. Bernardi, Can. J . Biochem. 46, 989 (1968). 36. J. Shack, JBC 226, 573 (1957). 37. M. E. Maver and A. Greco, JBC 181, 853 (1949). 38. K. D. Brorm, G. Jacobs, and M. Laskowski, JBC 194, 445 (1952). 39. T. R. Koszalka, K. Schreier, and K. I. Altman, BBA 15, 194 (1954). 40. 0. D. Kowlessar, S. Okada, J. 1,. Potter, and K. I. Altman, ABB 68, 231 (1957). 41. E. C. Rauenbusch and K. I. Altman, Proc. Soc. Exptl. Biol. M e d . 104, 385 (1960). 42. P. O w , 0. E. Brown, and J. Laszlo, ABB 131, 652 (1969). 43. M. S. Melzer, Can. J . Biochem. 47, 987 (1969). 44. N. R. Sarkar, BBA 145, 174 (1967).

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both pancreatic and acid DNase, the inhibition on acid DNase is much stronger than that on either pancreatic or E. coli DNase (6). The type of inhibition of actinomycin upon acid DNase is that expected for the case of inhibition by coupling of the inhibitor with the substrate but not with the enzyme (6).This is not surprising in view of the strong binding of actinomycin by guanylic acid residues in DNA and of the high guanylic acid level in the sequence split by acid DNase (see below). A protein inhibitor has been extracted and partially purified from mouse liver by Lesca and Paoletti ( 4 5 ) .This protein inhibits acid DNases from different tissues and species but not pancreatic or E . coli DNases. Very interestingly, V vs. substrate concentration plots become sigmoid in the presence of the inhibitor provided that pH is lower than 5.6. The existence of a DNase-inhibitor complex is suggested by sucrose-gradient results. An unusual feature of the inhibitor is its ability to reactivate acid DNase preparations treated with 8 M urea. A weak competitive inhibitory effect of bases, nucleosides, and nucleoside mono- and polyphosphates has been reported ( 4 6 ) . 3’P-oligonucleotides are very weakly inhibitory. Inhibition by natural and biosynthetic polyribonucleotides of the type previously found for some bacterial DNases (47-50) has been demonstrated to occur in the case of spleen acid DNase (51, 5 2 ) . The inhibition is, as in the case of E . coli DNase ( 5 0 ) , of the competitive type. With the remarkable exceptions of poly A and poly C, which did not show any effect on the DNase activity, all synthetic polyribonucleotides tested, transfer RNA and ribosomal RNA exhibited an inhibitory activity. This was very weak with the single-stranded polymer poly U. The finding that polyribonucleotides having single-stranded structures, like poly U, have very weak inhibitory properties and that single-stranded DNA is a poorer substrate than double-stranded DNA underlines the weaker binding of the enzyme by single-stranded structures. It should be stressed that the competitive inhibition by polyribonucleotides is specific and does not simply represent the binding of a polyanion by a basic protein; in fact, some polyribonucleotides are ineffective as inhibitors, and an

45. P. Lesca and C. Paoletti, Proc. Nntl. Acad. Sci. 46. H. Slor, Ph.D. Thesis, Indiana University, 1966.

c’, S. 64,

913 (1969).

A. W. Bernheimer and N. K. Ruffier, J . E x p t l . M e d . 93, 399 (1951). A . W. Bernheimer, BJ 53, 53 (1953). L. Kozloff, Cold Spring Harbor S y m p . Quant. Bid. 18, 209 (1953). I. R. Lehman, G. G. Roussos, and E. A. Pratt, JBC 237, 819 (1962). G. Bernardi, B B R C 17, 573 (1964). A. Jacquemin-Sablon, J. Laval, J.-Y. Le Talaer, J.-B. Le Pecq. and C. Paoletti, C o m p t . R e n d . 259, 2551 (1964). 47. 48. 49. 50. 51. 52.

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excess of cytochrome c, a strongly basic protein, in the incubation mixture does not interfere with inhibition.

3. “Phosphodiesterase” Activity Acid DNase from hog spleen catalyzes the slow hydrolysis of p-nitrophenol from bis (p-nitrophenyl) phosphate and the p-nitrophenyl esters of deoxyribonucleoside 3‘-phosphates, but not from those of deoxyribonucleoside 5’-phosphates (9). Using bis (p-nitrophenyl) phosphate as the substrate, the pH optimum was found to be between 5.6 and 5.9. The activity in acetate is about twice as large as in succinate buffer. In acetate buffer, no significant changes occur upon addition of Mg2+ or EDTA. I n the 4-7 pH range, 0.01 A1 S O P and 0.01 M HPOi- g‘ive a very strong inhibition. Interestingly, plots of the initial velocity of hydrolysis of bis (p-nitrophenyl) phosphate vs. substrate concentration have an initial upward curvature, whereas they are hyperbolic when native D N A is used as the substrate. The conclusion of Bernardi and Griff6 (9) that the “phosphodiesterase” activity of acid DNase is an intrinsic property of the enzyme molecule has been recently challenged by Slor (46, 5 3 ) , Swensoii and Hodes ( 5 4 ) ,and Slor and Hodes ( 5 5 ) , who claimed to have obtained a separation of the two activities. I n fact, none of the reported results proves an actual separation of the two activities and constitutes an acceptable evidence against the two activities being carried by the same protein molecule. Some data suggest, however, that the “phosphodiesterase” activity may be inactivated prcferentially by some treatments. I n connection with the phosphodiesterase activity of acid DNase, see also Tables I and I1 in reference (56‘) and the related discussion (56‘a).

E. SPECIFICITY Determining the specificity of a DNase is a problem of great complexity since not only the enzyme itself must be extremely pure but also the other enzymes (exonucleases and phosphatases) used in chain length and terminal nucleotide determinations must be extremely pure ; in addition, extremely accurate anrtlytical methods are needed. I n retrospect, it appears that these requirements were only partially met in some 53. H. Slor, BBRC 38, 1084 (1970). 54. M. R. Swenson and M . E. Hodw. JBC 244, 1803 (1969). 55. H. Slor and M. E. Hodes, A B B 139, 172 (1970). 56. A . Bernardi and G. Bernurdi, this volume. 1’. 329. 56a. P. J. Sicsard. A . Obrenovitch, and G . Aubel-S;idron. FEBS Letters 12, 41 (1970) could not confirm the claim of Hodes el ctl. (54, 5 5 ) .

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investigations (5740) devoted to the specificity of acid DNase (61).

A series of investigations in some crucial areas such as the purification of ancillary enzymes (62-65), base analysis (66, 6 7 ) , and kinetics of the middle and terminal phases of acid DNase digestion (19, 65) were therefore made in the author's laboratory. As a result of this work, a new picture of the acid DNase specificity is emerging. The events characterizing the middle and terminal phase may be briefly described as follows. During the middle phase the composition of the 3'-P terminal and of the 5'-OH terminal and penultimate nucleotides of oligonucleotides obtained from calf thymus DNA by acid DNase digestion, in the incubation conditions of Fig. 1, is practically constant. Some results obtained in this phase are given in Table 111. Purine nucleotides form about 75% of the 3'terminals and of the 5'-OH penultimates with a predominance of G in

TABLE I11 SPECIFICITY OF SPLEEN ACID DNase ON CALFTHYMUS DNA 3'-Phosphate terminal nucleotide" A

G T C Purines Pyrimidines Average size (1

28 44 20 8 72 28 14-20

5'-OH 5'-OH terminal penultimate nucleotideb nucleotide* 22 34 13 31 56 44 9.6

52 23 16 9 75 25 9.6

From Soave et al. (19). From Ehrlich et al. (66).

57. S. Vanecko and M. Laskowski, JBC 236, 1135 (1961). 58. S. Vanecko and M. Laskowski, BBA 61, 547 (1962). 59. J. Doskocil and F. Sorm, Collection Czech. Chem. Commun. 26, 2739 (1961). 60. J. Doskocil and F. Sorm, Collection Czech. Chorn. Commun. 27, 1476 (1962). 61. M. Carrara and G. Bernardi, Biochemistry 7 , 1121 (1968). 62. A. Chersi, A. Bernardi, and G. Bernardi, BBA 129, 11 (1966). 63. A. Bernardi and G. Bernardi, BBA 155, 360 (1968). 64. A. Chersi, A. Bernardi, and G. Bernardi, BBA, submitted for publication. 65. S. Ehrlich, G. Torti, and G. Bernardi, Biochemistry (in press). 66. M. Carrara and G. Bernardi, BBA 155, 1 (1968). 67. G. Piperno and G. Bernardi, BBA (in press).

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the first case and of A in the second one. This indicates not only that enzyme splittings are very far from random, but also that the enzyme is able to recognize a sequence of a t least three nucleotides in DNA. It is possible, therefore, that acid DNase may be used to assess the relative concentrations of recognized sequences in different DNA's. During the terminal phase a drift in the composition of terminal nucleotides takes place, leading to a more random distribution of terminals.

IV. Distribution, lntracellular localization, and Biological Role

An acid DNase activity has been found in the cells of a number of animal tissues and species (see references 1-6 for reviews of the literature). An interesting problem is whether this activity is carried by similar protein molecules. An unequivocal answer could be obtained by comparing the chemical, physical, and enzymic properties of highly purified preparations obtained from different sources. This being a very long and difficult task, Cordonnier and Bernardi (35) compared the chromatographic and enzymic properties, the sedimentation coefficients, and the mechanism of action on native DNA exhibited by partially purified acid DNase preparations obtained from 15 different animal sources : (a) epithelial tissues-hog liver, hog kidney, and hog pancreas; (b) lymphatic tissues-hog spleen, calf spleen, and calf thymus; (c) tumor tissues-a fibroblastic sarcoma from AKR mice and a mammary epithelioma from C,H mice; (d) nonmultiplying cells-chicken erythrocytes, hog erythrocytes, and mackerel sperms ; (e) biological fluids-hog serum, bull seminal plasma, and human urine; and ( f ) an invertebrate, the clam Mercenuriu mercenariu. The results obtained strongly indicated that the enzymic activity is associated with protein molecules endowed with very similar properties. The enzyme levels in the different tissues examined by Cordonnier and Bernardi (5.5) were found to vary by as much as three orders of magnitude. The highest acid DNase levels were found in lymphatic and tumoral tissues; the lowest were found in cells (sperms and erythrocytes) that do not reproduce themselves anymore. This relationship between levels of acid DNase activity and capacity for proliferation or regeneration of a given tissue had already been observed by Allfrey and Mirsky (68). As far as the intracellular localization of acid DNase is concerned, 68. V. G. Allfrey and A. E. Mirsky, J. Gen. Physiol. 36, 227 (1952).

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such an activity was found to be associated with lysosomes by several authors (69). On the other hand, it has been recognized very early that acid DNase activity is latent and that tissue autolysis or acidic treatintent is necessary to release it. J. M. Van Dyck and G. Bernardi (unpublished experiments) found that the enzyme from rat liver tritosomes has the same sedimentation coefficient as spleen acid DNase. It may be interesting to recall that acid DNase has been the first lysosomal enzyme obtained as a homogeneous protein. Concerning the biological role of acid DNase, it has been suggested by several authors that this enzyme might be involved in some essential biological mechanism, like DNA replication, where it might play some accessory role, and DNA recombination. If, however, one considers that lysosomes appear to contain all enzymes needed to degrade nucleic acids to nucleosides (Table IV) , it seems more likely that, a t least under TABLE IV DEQRADATION OF NUCLEIC ACIDSBY LYSOSOMAL HYDROLASES DNA (Acid DNase)

RNA (Acid RNase)

oligonucleotides 3 ’ P

Mononucleotides 3 ’ P (Acid phosphatase)

Nucleosides ‘Hypothetical.

69. J. T. Dingle and H. B. Fell, eds., “Lysosome in Biology and Pathology.” North-Holland Publ., Amsterdam, 1969.

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normal conditions, the biological role of acid DNase is a degradative one. Its diplotomic mechanism of action might be an extremely effective way of degrading foreign DNA. From the information gathered so far on acid DNase and other lysosomal hydrolases, it appears that these enzymes share some properties, like the basic character and the absence of SH groups, of digestive enzymes which are excreted from pancreatic cells into the intestinal tract. In this connection, it may be pertinent to mention here that a set of hydrolases of nucleic acids, recalling those of lysosomes, are found in bacterial cells such as E . coli and that they are localized between cell wall and cell membrane (7'0).

70. C. Cordonnier and G. Bernardi, BBRC 20, 555 (1965).


Deoxyribonuclease I M. LASKOWSKI, SR. I. Introduction . 11. Chemical Nature 111. Active Center . IV. Inhibitor . . V.Ions . . . VI. Kinetics . . VII. Specificity . . VIII. Physiological Role

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297 299 302 303 308 310

I. Introduction

During the past decade a previously accepted notion that the deoxyribonucleic acid (DNA)-deoxyribonuclease (DNase I) reaction runs a uniform course with a uniform specificity began to be seriously doubted. It is now realized that striking differences exist between the early and terminal stages of the same reaction. The observed differences are not limited solely to the rate of the reaction but include variations in endoor exonucleolytic character, in the effect of the divalent cation, and, finally, in the specificity toward the bases adjacent to the bond that is cleaved. Several reviews and books devoted exclusively t o methodology for following the action of DNase I exist (1-5). For the purpose of this 1. “Methods in Enzymology,” Vol. 2, Sect. 2, 1955. 2. “Methods in Enzymology,” Vol. 6, Sects. 1 and 2, 1963. 3. “Methods in Enzymology,” Vol. 12, Parts A and B, 1967, 1968. 4. G. L. Cantoni and D. R. Davies, Procedures Nucleic Acid Res. Sect. A (1966). 5. N. Kurnick, Methods Biochem. Anal. 9, 1 (1962). 289

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review it suffices to say that it is possible by the use of the p H stat to measure the number of internucleotide bonds cleaved. This method is independent of the stage of the reaction or of the location of the bond within the molecule. Methods measuring changes in molecular weight reflect the number of double-strand scissions. Finally, with the use of two other enzymes, polynucleotide kinase and DNA ligase (see below), it is possible to evaluate the number of “nicks” inflicted on one of the strands without causing a scission. Methods based on spectrophotometry reflect the collapse of ordered structure. Finally, methods measuring the appearance of mononucleotides reflect exonucleolytic activity. Among endonucleases which hydrolyze DNA one seldom finds an enzyme that attacks double-stranded and single-stranded substrates with equal ease. If the enzyme shows preference for double-stranded substrates (as DNase I does) autoretardation is observed. This decrease in the reaction rate is caused by the gradual disappearance of the preferred, double-stranded substrate and an increase in the concentration of less susceptible, single-stranded substrate. Differences in rates between the early and terminal phases of the reaction of the order of 1OOOfold have been described ( 6 ) . The opposite case, autoacceleration, is seen with those enzymes that show preference for the single-stranded structure, e.g., micrococcal nuclease ( 7 ) . I n the original meaning (8,9)endonucleases and exonucleases were conceived as retaining their character throughout the whole course of the reaction. It is now established that at least some typical endonucleases acquire exonucleolytic character toward the end of the reaction (10). Proximity of the newly created monophosphoryl or hydroxyl group is responsible for this change. Many DNases are known to be activated by a divalent cation. However, only from the work of Bollum (11) did it become clear that the nature of the cation may qualitatively change the specificity of the enzyme toward adjacent bases. Quantitative changes in the requirements for the divalent cation (10) have been observed during different stages of the same reaction, e.g., micrococcal nuclease (7) where the increased Ca2+concentration causes a decrease in the average size of the terminal product. Finally, it was shown with a number of DNases that during the 6. S. Vanecko and M. Laskowski, Sr., JBC 236, 3312 (1961).

7. E.Sulkowski and M. Laskowski, Sr., JBC 243, 4917 (1968). 8. M.Privat de Garilhe and M. Laskowski, Sr., JBC 223, 661 (1956). 9. M.Laskowski, Sr., G. Hagerty, and U.-R. Laurila, Nature 180, 1181 (1957) 10. M. Laskowski, Sr., Advan. Enzymol. 29, 165 (1967). 11. F.J. Bollum, JBC 240, 2599 (1965).

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course of the reaction cleavages become less specific (10). By extrapolation, the first few cleavages must be very specific. The direct proof for this statement is delayed by technical difficulties in determining terminal nucleotides in fragments of 1000 or more monomers. I n spite of this, it seems safe to predict that in the near future the DNases previously considered to be nonspecific will be used to inflict a very limited number of very specific cleavages. Regulation of ionic medium, pH, temperature, and exposure time can be expected to significantly improve specificity in the early stages of the reaction. On the other hand, it becomes equally evident that in the terminal stages of the reaction, in addition to the base specificity the effect of the monophosphoryl group determines the end point of the reaction (1.2). The reasons for selecting pancreatic DNase I as one of the two representative of mammalian DNases are to a large extent historical. Deoxyribonuclease I was the first enzyme to be recognized as specific for DNA (13-16),the first DNase to produce 5’-monoesterified products (16, lY), the first DNase to be crystallized (It?), the first DNase to have a specific protein inhibitor (19-23),the first DNase shown to produce “nicks” on one strand in preference to scission of both strands ( 2 4 , 2 6 ) .A new first has been added recently (25a) ; DNase I was covalently coupled to porous glass, thus supplying an insoluble DNase. The articles on DNases in previous editions of “The Enzymes” (26,xT) discussed several of these issues in historical perspective. The historical discussion will not be repeated in the present edition except when new information requires an introduction. 12. A. J. Mikulski, E. Sulkowski, L. Stasiuk, and M. Laskowski, Sr., JBC 244, 6559 (1969). 13. J. P. Greenstein and W. V. Jenrette, J . Natl. Cancer Inst. 1, 845 (1941). 14. M. Laskowski, Sr. and M. K. Seidel, A B B 7, 465 (1945). 15. M. McCarty, J . Gen. Physiol. 29, 123 (1946). 16. J. L. Potter, K. D. Brown, and M. Laskowski, Sr., BBA 9, 150 (1952). 17. R. L. Sinsheimer and J. F. Koerner, JACS 74, 283 (1952). 18. M. Kunitz, J . Gen. Physiol. 33, 349 (1950). 19. W. Dabrowska, E. J. Cooper, and M. Laskowski, Sr., JBC 177, 991 (1945). 20. E. J. Cooper, M. L. Trautman, and M. Laskowski, Sr., Proc. SOC. Exptl. B i d . M e d . 73, 219 (1950). 21. L. Cunningham and M. Laskowski, Sr., BBA 11, 590 (1953). 22. U. Lindberg, Biochemistry 6, 323 (1967). 23. U. Lindberg, Biochemistry 6, 343 (1967). 24. S. Zamenhof, G. Griboff, and N. Marullo, BBA 13, 459 (1954). 25. E. T. Young, I1 and R. L. Sinsheimer, JBC 240, 1274 (1965). 25a. A. R. Neurath and H. H. Weetall, FEBS Letters 8, 253 (1970). 26. M. Laskowski, Sr., “The Enzymes,” 1st ed., Val. 1, p. 956, 1951. 27. M. Laskowski, Sr., “The Enzymes,” 2nd ed., Val. 5, p. 123, 1961.

M. LASKOWSKI, SR.

II. Chemical Nature

Almost as soon as bovine pancreatic crystalline DNase I was obtained, doubts concerning its homogeneity arose. Even before crystallization (18) it was shown (28,282sa) that DNase I cocrystallizes with chymotrypsinogen B. The chronologically first crystalline product contained about two-thirds chymotrypsinogen B and about one-third DNase I (28, 28a). It would, therefore, be expected that the reverse also occurs. I n fact, Potter (29) showed that a commercial sample of crystalline DNase can be separated into five protein-containing bands on cellulose acetate strips. One of these bands was identified as chymotrypsinogen B. With a sensitive method of detection (31) the presence of one part of RNase per 100,000 parts of DNase I was found in the crystals. This activity could be further reduced by continuous flow electrophoresis (S2), or more efficiently by chromatography on DEAE-cellulose (33). Lindberg (34) passed the solution of commercial crystalline DNase I through a column of Sephadex G-100, removed contaminants of smaller molecular weight, and obtained a preparation of high purity. A series of beautiful papers from the laboratory of Moore and Stein ( 3 5 3 9 a ) recently appeared, confirming that crystalline DNase I is contaminated with about one-third chymotrypsinogen B and chymotrypsin B. It was first established (35) that commercial DP grade ( 3 2 ) DNase I is composed of a t least two, enzymically active glycoproteins. Figure 1, reproduced from the work of Price et al. ( S 5 ) , shows the separation on SE-Sephadex into two equally active components B and A. A modified procedure (39) of chromatography on phosphocellulose (Fig. 2) led to three peaks, C, B, and A (in order of their appearance from the column). A further refinement of technique (39a) allowed visualization of a small additional 28. M. Laskowski, Sr., JBC 166, 555 (1946). 28a. M. Laskowski, Sr. and A. Kazenko, JBC 167, 617 (1947). 29. J. L. Potter, personal communication, quoted by Laskowski (SO). 30. M. Laskowski, Sr., Procedures Nucleic Acid Res. p. 83 (1966). 31. J. Polatnick and H. L. Bachrach, Anal. Biochem. 2, 161 (1961). 32. Worthington, catalog (1965). 33. S. B. Zimmerman and G. Sandeen, Anal. Bioehem. 14, 269 (1966). 34. U. Lindberg, Biochemistry 6, 335 (1967). 35. P. A. Price, T.-Y. Liu, W. H. Stein, and S. Moore, JBC 244, 917 (1969). 36. P. A. Price, S. Moore, and W. H. Stein, JBC 244, 924 (1969). 37. P. A. Price, W. H. Stein, and S. Moore, JBC 244, 929 (1969). 38. B. J. Catley, S. Moore, and W. H. Stein, JBC 244, 933 (1969). 39. J. Salnikow, W. H. Stein, and S. Moore, Federation Proc. 28, 344 (1969). 398. J. Salnikow, S. Moore, and W. H. Stein, JBC 245, 5685 (1970).

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peak D. Each component had comparable specific activity. Peaks A:B:C were present in a constant ratio of 4:l:l. The major peak A corresponded to peak A of the previous separation (35). Figure 2 shows a portion of the chromatographic pattern obtained on phosphocellulose. Peak A has been studied further. Table I, reproduced from the paper of Price et al. (35),compares amino acid content of peak A with that obtained by Lindberg (34) for the purified DNase I. The composition reported from the two laboratories for preparations obtained by two different methods is surprisingly similar. The linear sequence is being determined in the laboratory of Moore and Stein (39b). The possibility that the three fractions are artifacts caused by the exposure to a strong acid during the preparation procedure has been ruled out by the experiment with a freshly collected pancreatic juice. The juice was first chromatographed on DEAE-cellulose according to Keller et al. (40) and the fraction containing DNase was then rechromatographed on phosphocellulose (39) and gave three identical peaks as were seen with crystalline DNase. To elucidate the mode of attachment of the carbohydrate moiety to the protein of DNase I, Catley et al. (38)digested the peak A DNase with Pronase and subjected the digest to gel filtration on Sephadex G-25. All of the carbohydrate was recovered in a mixture of the dipeptide SerAsp and the tetrapeptide Ser-Asp-Ala-Thr. Removal of serine by an Edman degradation demonstrated that all of the carbohydrate was in association with aspartic acid. Analysis of the carbohydrate moiety demonstrated two residues of glucosamine, five residues of mannose, and one residue of ammonia, leading to the conclusion that the saccharide moiety is attached at a single position on the enzyme through an aspartamidohexose linkage. An analysis of the peak B DNase identified the same sugars in the same proportions except that sialic acid was present in fractional quantities. Since sialic acid in the intact peak B DNase analyzed for 0.2 residue and in a tetrapeptide fraction for 0.06 residue, it was considered to be part of an impurity not associated with the heptasaccharide moiety. If the presence of fractional quantities of sialic acid is accidental, the most probable reason for the separation of peaks C, B, and A appears to be the number of amides since the gross amino acid composition is not different. Some minor corrections of the values shown in Table I are required. The latest work (39a) shows that peaks A and B have identical amino acid composition. Peak A contains two residues of N-acetylglucosamine 39b. S. Moore and W. H. Stein, personal communication (1970). 40. P. J. Keller, E. Cohen, and H. Neurath, JBC 233, 344 (1958).

M. LASKOWSKI, SR.

oo

0

- 100 >

0

o o o

c

V

._ c

'"

-,o

N

+

3

0

0)

c

e

n P

a

10 0

10 0 Effluent(ml)

40

60

80

FIG.3. Sephadex G-100 chromatography on DNase I, inhibitor 11, and mixture containing the two proteins. (A) DNase I only, (B) inhibitor I1 only, (C) and (D) both components with different molar excess of inhibitor, (E) equimolar amounts of inhibitor and enzyme, (F) and ( G ) both components with a different excess of enzyme. Absorbance a t 215 nm (solid line) was measured after 20-fold dilution with water using a similarly diluted blank of the elution buffer (0.5 M potassium phosphate, p H 7.6). Each chromatogram was analyzed for DNase activity (01, inhibitor activity ( O ) , and for the presence of DNase-inhibitor complex, in this figure represented as DNase I activity which was measured on samples of the fractions after adjustment of the pH to 3.5 with HCl ( 0 ) .[From Lindberg (3 4 ). Copyright 1967 by the American Chemical Society. Reprinted by permission of the copyright owner.]

12. DEOXYRIBONUCLEASE I

30 1

(49,000 instead of 59,000 for spleen inhibitor 11) and in the maximum stability which is a t pH 6 for thymus and pH 7 for spleen. One of the most exciting aspects of the problem is the exact mechanism of the union of the two proteins. Striking progress in understanding of the mechanism of the union between trypsin and trypsin inhibitor has been made in the laboratory of Laskowski, Jr. (46, 4 7 ) . The trypsin inhibitor-trypsin complex is essentially a Michaelis-Menten complex. During the union one trypsin-sensitive bond in the inhibitor (reactive site) is cleaved without affecting the inhibitory power of the protein, thus creating “the modified form” of the inhibitor. If one makes an analogy to enzyme other than trypsin, it would be expected that the inhibitor for DNase I should be a specifically resistant DNA. However, the protein nature of both partners, DNase I and spleen inhibitor 11, is well established. Obviously, not all protein-protein interactions must be of the trypsin inhibitor-trypsin type, but it may be worthwhile to check the DNase inhibitor for possible proteolytic activity. One is tempted to speculate that in this case the inhibitor I1 may be a protease specific for a peptide linkage involving histidine. To prove it experimentally, it would be necessary to trap the “modified form of DNase I” before the bond is re-formed. Several years ago the question of whether DNase I is a strictly digestive enzyme was argued. On the basis of experiments in which DNase I type of activity was found in the minced tissue only after the previous exposure to an acid pH, known to destroy the inhibitor, it was concluded (21) that the DNase I type of enzyme is intracellular and ubiquitous. Recently, Lee and Zbarsky (48) used pigeon crop gland inhibitor to identily a DNase I type of activity in the intestional mucosa of the rat. The crop gland inhibitor and presumably the spleen inhibitor I1 react with DNase I of many species. It appears that no species specificity exists in the reaction of complex formation. The above considerations could bring us to the discussion of the occurrence of DNase I in different tissues and in different organelles of the same cell. A fairly extensive literature exists, particularly in reference to different pathological conditions. It will not be considered in this review. In many papers only two criteria are used to classify an enzyme as DNase I : (1) It requires Mg”, and (2) it has an optimum a t pH 7. 46. M. Laskowski, Jr., S y m p . , Structure-Pri?iction Relationship of Proteolytic En(P. Desnuelle, H. Neurath, and M. Ottessn, eds.) p. 89. Munksgaard, Copmliagen, and Academic Press, New York. 1970. 47. M. Laskowski, Jr. and R . W. Senlock, “The Enzymes,” 3rd ed., Vol. 111, p. 376, 1971. 18. C. Y. Lee and S. 11. Zbarsky, Can. J . Biochem. 45, 39 (1967). zymes, 1968

302

M. LASKOWSICI, SR.

Historically, these criteria are justified, but they are no longer sufficient. Additional information characterizing the enzyme as a 5'-monoester former would be desirable, but the ability to react with the specific DNase I inhibitor appears to be the most important criterion. Without having tested for it the enzyme should not be called DNase I.

V. Ions

Many years ago it was observed that the presence of either Mg2+or MnZ+increased the rate of hydrolysis of DNA by DNase I whereas high concentrations of NaCl decreased it [see review (10)1. The revolutionary finding comes from the work of Bollum (11) who showed that the nature of activating cation qualitatively affects specificity. Deoxyribonuclease I was presented with (dI),.(dC), as substrate. Only (dI), but no (dC), was hydrolyzed when 1 0 m M Mg2+ was the sole activating divalent cation. If, in addition to Mg2+,2 mM Ca2+was introduced, both strands were hydrolyzed. The same result (both strands digested) was obtained when 10 m M Mn2+ alone was present instead of the mixture of Mg2+and Ca2+.A recent paper (48a) describes the elegant separation and identification of di- and trinucleotides in the DNase I digest. The composition of digests obtained in the presence of Mn2+is different from that obtained in the presence of MgZ+. This result strongly supports Bollurn's (11) conclusion. This discovery of Bollum (11) makes obsolete a number of previous excellent studies including those on mutual interdependence between concentrations of divalent metal, monovalent metal, hydrogen ion, and substrate. Unless the bond affected by the metal in question is specified, an overall rate represents a number with little value. The problem is further complicated by the suspected (by analogy to other nucleases) quantitative changes in requirements for metal ions a t different stages of the reaction. So far no such data are available for DNase I. One is tempted to add, luckily, because in view of the uncertainty of qualitative effects such data would hardly be expected to have a long survival time. Melgar and Goldthwait (49, 50) used a method in which isotopically labeled DNA was incorporated into acrylamide gel. The suspension of the gel containing DNA was used as substrate. The average molecular 48a. E. Juchnowicz and J. H. Spencer, Biochemistry 9, 3640 (1970). 49. E. Melgar and D. A . Goldthwait, JBC 243, 4401 (1968). 50. E. Melgar and D. A. Goldthwait, JBC 243, 4409 (1968).

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weight of fragments released from the gel was approximately 400,000. In the presence of Mg2+ alone the rate of release of these fragments with DNase I showed a lag period. No such lag period was observed with either DNase I1 or E . coli endonuclease I, known to make doublestrand scissions. The lag was eliminated when Mn2+,Caz+, or Co2+,or Mg2+plus Ca*+was used. Sodium added to Mn2+,to Ca2+,or to Ca2+plus Mg2+reestablished the lag. The results are interpreted as indicating that only single-strand cleavages occur during the lag period, whereas double-strand scissions release the fragments from the gel. This interpretation was confirmed by viscometry and ultracentrifugation. Eichhorn et al. (51) concluded that Co*+is a better activator of DNase I than any of the previously used metals. As a criterion of activity the authors used the formation of acid-soluble products, corresponding to terminal stages of the reaction. I n neither of these papers (49, 51) was the attempt made to characterize the split bond.

VI. Kinetics

As mentioned in the Introduction, a characteristic aspect of the kinetics of DNase I acting on native DNA is autoretardation (10). Autoretardation is caused by the continuous formation of products which are poorer substrates than those from which they are derived. Three types of experiments were performed to prove this (6). Experiments of the first type were performed in a p H stat. The reaction was allowed to run until the originally fast rate reached a plateau. At this time a 100-fold excess of enzyme was added. The initial rate of the reaction was restored, then slowed down, and reached a second plateau. Further addition of a fourfold amount of enzyme (the total enzyme concentration was now 500-fold that of the original) resulted in a new burst of activity, but the rate was slower than the original. A second type of experiment was performed by isolating the reaction products a t different stages of the degradation and using them as substrate for a fresh sample of DNase I. The reaction again was followed in a p H stat. The rate was highest with native DNA. To obtain comparable rates the amount of enzyme needed was 15-fold with the “I-min digest,” 500-fold with the “10-min digest,” and 2500-fold with “oligonucleotides” obtained from the reaction in which the first plateau was reached. 51. G. L. Eichhorn, P. Clark, and E. Tarien, JBC 244, 937 (1969).

304

M. LASKOWSKI, SR.

The third type of experiment was performed to show that inhibition by the products is insufficient to account for the observed difference in rate. The substrate consisted of a 50:50 mixture (in terms of optical density) of native DNA and “oligonucleotides.” With this mixture the rate of hydrolysis decreased to about one-half of that for native DNA. This result agrees with the previously established (52) competitive-type inhibition by the products. It also shows that the inhibition by products accounts for changes in rate within one order of magnitude, whereas the decreasing affinity toward the newly formed substrates accounts for more than three orders of magnitude. With so pronounced a difference in susceptibility of substrate in the very early and in the very late phases of the reaction, the two phases must be considered independently. Kinetics of the early phases of the reaction have been studied by Dekker and Schachman (53),Schumaker et al. (54),Thomas ( 5 5 ) , and Young and Sinsheimer ( 2 5 ) . All the results demonstrate that DNase I hits a t random and that several hits on a single strand occur before one double-strand scission is detected. The estimates of numbers, however, vary; Thomas considered that an average of 200 single hits occur before the average molecular weight has decreased by a factor of 2. Young and Sinsheimer (25) estimated that an average of 4 hits occur before h-phage DNA is inactivated, possibly as a result of a doublestrand scission. The ability of DNase I to inflict a number of a single-strand breaks (or nicks) before producing a detectable decrease in molecular weight has been utilized by Richardson and his colleagues to create a detecting system in their studies of polynucleotide kinase and DNA ligase (56, 5 t h ) . The scheme of sequential reactions involved in nicking and labeling the nicked ends is reproduced in Fig. 4. Figure 5 shows the dependence between the number of single-strand breaks and concentration of DNase I. In view of the impact of this work on several lines of nucleic acid research some of the experimental details are given in the legends. Figure 6 shows the scheme of a procedure by which the labeled nicks are first closed with the aid of DNA ligase. Deoxyribonucleic acid is then degraded enzymically in such a manner that allows the identification of the nucleosides adjacent to the labeled internucleotide linkage. Table I11 shows the composition of nucleotides adjacent to the nick 52. L. F. Cavalieri and B. Hatch, JACS 75, 1110 (1953). 53. C. A. Dekker and H. K. Schachman, Proc. Natl. Acad. Sci. U . S. 40,894 (1954). 54. V. N. Schumaker, E. G. Richards, and H. K. Schachman, JACS 78,4230 (1956). 55. C. A. Thomas, Jr., JACS 78, 1861 (1956). 56. B. Weiss, T. R. Live, and C. C. Richardson, JBC 243, 4530 (1968). 56a. B. Weiss, A. JaqueminSablon, T. R. Live, G. C. Fareed, and C. C. Richardson, JBC 243, 4543 (1968).

32PJ-

L-

-P -"J

1

Polynucleotide Kinase APP+p3'

P32

.-I

PY

FIG.4. Scheme of the preparation containing =P-labeled phosphomonoesters a t single-strand breaks, The two strands of TT DNA duplex are schematically represented by two parallel lines and only the 5' termini are designated. After the introduction of single-strand breaks into DNA by incubation with pancreatic DNase, the phosphomonoesters formed are removed by phosphatase a t 65". The 5' termini are then labeled by incubation with polynucleotide kinase. From Weiss et al. (66).

DNase concentralion,units /ml

1 k . 5. Production of single-strand breaks by pancreatic DNase. T, DNA w m incubated with varying amounts of pancreatic DNase. After each reaction, the number of single-strand breaks (internal phosphomonoesters per 4.0 X 10' nucleotides) was measured by end group labeling (see Fig. 4 ) . Crystalline pancreatic DNase I (11 mg. 1 vial Worthington) was dissolved in a 1-ml solution containing 10 m M sodium acetate buffer, pH 5.5,5 mM MgCl,, 0.2M NaC1, and 0.5 mg/ml of bovine plasma albumin. The mixture was stored at 0" for up to 1 month during which time it gradually lost 10-25% of its activity. Immediately before use it was diluted with the same solution and assayed spectrophotometrically by the method of Kunitz. One unit of enzymic activity was defined as the amount of enzyme causing an increase in Az, of 0.001/min/ml of assay solution at 25". Deoxyribonucleic acid was incubated in 5 ml volume containing 1.3 mM DNA, 67 mM tris-HC1 buffer (pH 8.0), 5 m M MgCl,, and 0.5-5 units of DNase I at 20" for 30 min. To stop the reaction EDTA (0.5M,pH 7.5) was added to attain 16 mM concentration. The niixture was dialyzed for 8 hr at 4" against 20 n d 1 NaCl-10 mM tris-HC1 buffer (pH 8.0) and stored up to 6 months at 0".From Weisa et al. (66).

306

M. LASKOWSKI, SR. X

Y

Z

A B C

- -- p l p J p .bz i p l

p

1,-- -

I

Ligose system

X Y Z A B C

- - -pJ P

1 k 1 1,- - P

diesterase sple~/

PJ P

k g a s e

2 HOJps 3-Mononwleotides

32p

4

OH

5'-Mononmleotides

FIG.6. Scheme for nearest neighbor analysis of phosphodiesters formed in the ligase reaction. From Weiss et al. (66a).

TABLE I11 NEARESTNEIGHBORANALYSISOF NUCLEOTIDES JOINED IN LIGASE REACTION"^^ DNA (Fig. 6) prior to the action of ligase 3'-Nucleotides 5'-Nucleotides 5'-Nucleotides DNA (Fig. 6) after the action of ligase

Nucleotide

(%I

(%I

(%)

dAMP d T MP dGMP dCMP

31

22 59

22

44 11

14

8 11

57 9 12

From W e i s et al. (664. A T? DNA preparation (300 mpmoles) containing 90% (the other sePwas terminal) of its a*Pin internal phosphomonoesters was incubated in the standard ligase reaction mixture for 30 min with 0.01 unit of DNA ligase. An aliquot of the reaction mixture was subjected to the standard assay procedure; 90% of 3 2 P in the DNA became insusceptible to phosphatase. The incubation mixture was dialyzed, incubated with phosphatase for 30 min a t 65" to remove any remaining phosphomonoesters. The protein was extracted with phenol, and the DNA was dialyzed against four changes of 0.01 M tris-HC1 buffer (pH 7.6)-0.05 M NaC1. One aliquot of DNA was hydrolyzed completely to 5'-mononucleotides (see Fig. 6) by the consecutive action of pancreatic DNase I and venom o-exonuclease. Another aliquot was hydrolyzed to 3'-mononucleotides by the action of micrococcal nuclease and spleen a-exonuclease. The only labeled nucleotides were those adjacent to the original DNase I cleavage (Fig. 4).

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which was originally inflicted by DNase I (Figs. 4 and 6). The data of Table I11 show that all four bases are present, but A and T are significantly favored. Little can be said concerning the location of nicks relative to the center of the molecule. Present evidence suggests th a t central locations are strongly favored (49, 5 3 ) . This is supported by inability to detect any sign of exonucleolytic character in the early phases of the DNase I reaction (67).A surprising and as-yet unexplained result of the latter work was the finding that in comparison to other nucleases DNase I showed an exceptionally high hyperchromic shift per cleavage during the early phase. As the reaction progressed a plateau was reached, even though titrimetry indicated a continuation of the reaction. One is tempted to speculate that the early nicks occur in such regions where the unwinding of an extended portion of the helix is possible (e.g., A,Trich regions). On the other hand, the cessation of optical changes prior to titrimetric changes suggests that a considerable portion of the oligonucleotides remains double stranded. An elegant study of the kinetics has been performed on single-stranded biosynthetic polymers (68),prepared by the method of Bollum et al. (69). Three polymers were used : (1) d ( [ 3H]PA) (PA)lzl (2) d ( ['HI PA) (PA)llii ( [ 2-14C]PA) 4 ~ ;7 and (31 d ( [3HI PT) (PA) i ~ The . digestion was carried out in the presence of Mg2+ and allowed to proceed in a pH stat until 10% of internucleotide bonds were hydrolyzed. The reaction was stopped by heating on a steam bath. The size of the products in the digestion mixture was determined by means of chromatography on a column of Bio-Gel P-60 previously calibrated with oligomers. The overall conclusion (58) is that the rate of attack on diester bonds within 10 nucleotides of an end is much smaller than on bonds in the central region, if the substrate molecule is several multiples of 10 nucleotides or less in length, but that this discrimination disappears as the substrate length increases. In connection with autoretardation it is interesting to note that with a single-stranded homopolymer, there is a greater probability of producing fragments larger than 10 than of producing shorter fragments. Fragments of about 15 are attacked slower than the larger ones. All these findings confirm autoretardation and extend it into a phase of the reaction, in which little or no double-stranded substrate remains. The observation that short fragments (less than 10) 57. E. J. Williams, S.-C. Sung, and M. Laskowski, Sr., JRC 236, 1130 (1961). 58. D. E. Hoard and W. Goad, J M B 31, 595 (1968). 59. F. J. Bollum, E. Goreniger, and M. Yoneda, Proc. Natl. Acad. Sci. U.S. 51, 853 (1963).

308

M. LASKOWSKI, SR.

are formed faster in the earlier than in the latter stages of the reaction implies that DNase I either recognizes structure up to 100 A long (3 x 10 oligomers) or recognizes such aspects of the tertiary structure of homopolymers that exist in the long, but not in short, fragments. As in the experiment of Bollum (11), Hoard and Goad ( 5 8 ) , observed that the thymidylic acid fragment was deiraded faster than the corresponding fragment containing adenylic acid. This was in contrast to the report of Ralph et al. (60). Since Bollum and Hoard and Goad used Mg2+whereas Ralph et a2. used Mn2+as activating cation, it seems possible that the observed differences were caused by the nature of the metal (see Section V ) . Entirely different kinetics characterize the terminal phase of the reaction. At this phase, the remaining substrates are quite resistant. Presumably, the end point of the reaction is primarily affected by the concentration of the activation ion.

VII. Speciflcity

The previous characterization of specificity of DNase I (26, H )was arrived a t from the analysis of digestion products at the time of termination of the reaction. Three tacit assumptions were made, all of which are probably false: (1) the specificity does not change during the whole course of the reaction, (2) the nature of the activating ion does not influence specificity, and (3)the end point of the reaction is characteristic for the enzyme and is independent of the medium (ions, pH, type of substrate, etc.) Although the earlier data are valid for the conditions under which they were collected their generality needs to be reexamined. A long time ago, the work in the Sinsheimer’s laboratory (61-63)and in ours (64) established that among the products present a t the so-called termination of the reaction, dinucleotides of the type pPu-pPy were either rare or absent. This was interpreted as evidence that the Pu-pPy bond is the most susceptible to DNase I [for details, see reviews (10, 27, 30)1. This conclusion rests on the validity of the first assumption (see above). The recent work of Scheffler et al. (65) is the only work that supR. K. Ralph, R. A. Smith, and H. G. Khorana, Biochemistry 1, 131 (1962). R. L. Sinsheimer and J. F. Koerner, JACS 74, 283 (1952). R. L. Sinsheimer, JBC 208, 445 (1954). R. L. Sinsheimer, JBC 215, 579 (1955). M. Privat de Garilhe, L. Cunningham, U.-R. Laurila, and M. Laskowski, Sr., JBC 224, 751 (1957). 65. I. E. Scheffler, E. L. Elson, and R. L. Baldwin, J M B 36, 291 (1968). 60. 61. 62. 63. 64.

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ports this premise. The authors digested biosynthetic d (A-T) polymer with DNase I in the presence of Mg” and found only the products of the type d(pTpA),, where n was any integral number. No fragments starting with A, and no odd numbered fragments were detected. These findings show that only the cleavages between A and p?” were occurring throughout the whole course of the reaction, and no other cleavage. This agrees with the postulated specificity Pu-pPy. However, in many respects d (A-T) polymer behaves differently from a representative natural DNA with its four bases. The most obvious difference is its complementarity, which allows it to retain double-stranded conformation essentially throughout the reaction. To accomlish the double-strand scission the hydrolysis of an identical A-pT bond is required. The work on d(A-T), suggests that synthetic polymers may be most useful in elucidating some aspects of base specificity. The conclusion, however, that specificity remains unchanged throughout the whole course of the reaction is not directly transferable to DNA, where early cleavages appear to be considerably more specific than the later ones. I n Section VI the comparison of the rate of hydrolysis of (dA), and (dT), was discussed. The only way to account for the contradictory findings in different laboratories is to ascribe the differences to the activating ion used. The evidence exists that during the late phase of the reaction factors other than the Pu-pPy specificity determine the lability of the internucleotide bond. Thus, Potter et al. (42) easily digested d-ApApTp to d-ApA and d-pTp, whereas Khorana (43) observed that d-CpApT was resistant to DNase I. Both compounds had Pu-pPy sequence in the p - y positions, but differed with respect to 3’-terminal phosphate, which exerts a labilizing influence on the proximal internucleotide bond (see below). Doubts concerning the validity of the second and third premise stem from two sources: studies of the effects of metals on kinetics of DNase I, and from a comparison with other nucleases. Hurst and Becking (66-68) and Hacha and Fredericq (69) showed that whereas Mgz+ and Mn2+ both accelerated the action of DNase I, each led to a different mixture of products. Both groups suggested that DNase I may be a mixture of a DNase and an oligonucleotidase. I n view of Bollum’s findings (11) the probable explanation for these phenomena is that Mn2+ and Mg2+ affected susceptibility of different bonds in the substrate. The experiments in which a direct effect of concentration of a divalent 66. G. C. Becking and R. 0. Hurst, Can. J . Biochem. Physiol. 40, 166 (1962). 67. R. 0. Hurst and G. C. Becking, Can. J . Biochem. Physwl. 41, 469 (1963). 68. G. C. Becking and R. 0. Hurst, Can. J . Bwchem. P h y h l . 41, 1433 (1963). 69. R. Hacha and E. Fredericq, Bull. SOC. Chim. Belges 72, 580 (1963).

310

M. LASKOWSKI, SR.

cation on the end point of the reaction was demonstrated have been performed so far only with micrococcal nuclease (7). It seems likely, however, that with all DNases requiring a divalent cation, the requirement increases as the reaction proceeds. One issue was so far avoided, namely, the origin of mononucleotides. I n a digest stopped a t end point of the fast reaction the digestion mixture contains about 1% of mononucleotides. The maximum amount of mononucleotides ever observed after an exhaustive digestion was 5% ( 6 ) . All four mononucleotides are present but in different amounts. Ralph et al. (60) concluded that the smallest unit from which mononucleotides can be derived is a tetranucleotide, because in agreement with others (6) they found di- and trinucleotides resistant to large doses of DNase I . There is only indirect evidence suggesting that mononucleotides may originate from the o terminus (41). Summarizing the present status of our knowledge of the specificity of DNase I it is necessary to reemphasize the difference between the early and the terminal phase of the reaction. The only exception is biosynthetic d(A-T), (66). With DNA as substrate the early cleavages are directed toward the center of the molecule and are predominantly singlestrand nicks. By analogy to other nucleases one should expect that they are specific also with respect t o the adjacent bases. I n the latter part of the reaction the Pu-pPy bond is preferentially cleaved. The reaction can be carried to the stage when products are essentially a mixture of dinucleotides and trinucleotides. At this stage the term preferentially cleaved linkage is obviously nonapplicable.

VIII. Physiological Role

As yet no definite physiological role can be assigned to DNase I. The role of such an enzyme in digestion is testified by its presence in the pancreatic juice. However, the presence of DNase I activity in practically all other tissues casts doubt that the digestive function is its major role. The prevailing opinion assigns the major function of DNase I t o inflicting nicks during the early stages of hydrolytic attack on DNA. Thus participation in repair phenomena rather than complete digestion appears to be its major function. ACKNOWLEDOMENTS The experimental work referred to in this article and performed in our laboratory was generously supported by the American Cancer Society PRP-30 and E-157, by

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the U. S. Atomic Energy Commission AT(30-1)3630, and the National Science Foundation GB-6058. The author is indebted to Dn. Bollum, Moore, Richardson, Sinsheimer, and Weinfeld for the critical reading of the manuscript, and to Drs. Lindberg, Moore, and Richardson for permission to reproduce their data.


13 Venonz Exonuclease M.LASKOWSKI, SR.

.

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

. . . . .

I. Introduction . . . . . . 11. Chemical Nature of the Enzyme . , . 111. Structural Characteristics of Substrates Affecting Susceptibility A. Conformation . . . . B. The Nature of Sugar . . . . . . C. The Nature of Bases . . . . . . . D.Effect of Monophosphoryl Group . . . . IV. Venom Exonuclease as a Tool for Structural Determination . A. Sequence in Ribooligonucleotides . . . . B. Identification of a and o Terminals . . . . . V. Other Venom Enzymes That Hydrolyze Phosphate Esters . .

.

.

. . .

. . . . . . .

313 317 319 319 320 320 322 324 324 326 328

I. Introduction

Venom has long been known to be a good source of several enzymes that hydrolyze esters of phosphoric acid. I t is not possible to discuss venom exonuclease without mentioning other enzymes of this group. An effort will be made, however, to limit the discussion of other phosphatases to the bare essentials and key references. The surveys of different species of snake with respect to these enzymes are fairly numerous (1-9) and allow several conclusions to be drawn. 1. C . 4 . Chen and C.-C. Su, Yamaguchi Med. Coll. J . 8, 570 (1959). 2. C.-C. Yang, C. Iwanoga, and S. Kawashi, J . Formosan Med. Assoc. 57, 525 (1963). 3. S. K . Vasilenko, Biokhimiya 28, 602 (1963). 4. G.T.Babkina and S. K. Vasilenko, Biokhimiya 29, 268, (1964). 5. F.E. Russell, F. W. Buess, and M. V. Woo, Tozicon 1, 99 (1963). 6.B.D. McLennan and B. G. Lane, Federation Proc. 24, 602 (1965). 7. B. D. McLennan and B. G. Lane, Can. J . Biochem. 46, 81 (1968). 8. G.M. Richards, G. du Vair, and M. Laskowski, Sr., Biochemktry 4,501 (1965). 9. D.Mebs, Z. Physiol. Chem. 349, 1115 (1968). 313

314

M. LASKOWSKI, SR.

First, exonuclease (phosphodiesterase) , endonuclease (8, 10-12), 5’nucleotidase (13),and nonspecific phosphatase (13) are present in all venoms that have been analyzed. Therefore, nature did not provide these materials as sources of any one of these enzymes free from undesirable contaminants which may be represented by the others. Second, a new ribonuclease has been reported (3, 4, 6 , 7, 14) in three different species of snake which are widely separated geographically. I n addition, a monophosphatase that specifically or preferentially (7) attacks pNp is strongly suspected. Third, the quantitative differences in concentration of the four enzymes vary appreciably therefore making several venoms an undesirable source of exonuclease (8). All partially purified preparations of venom exonuclease exhibit adenosine triphosphate-pyrophosphatase activity (cleavage of the a-p linkage). Pfleiderer and Ortanderl (14a) studied this issue and showed that during purification the ratio of the two activities remains constant, concluding that both activities are intrinsic properties of the same enzyme. The terms venom exonuclease and venom phosphodiesterase are a t present used interchangeably to designate the same enzyme. The reviewer prefers the first, because he would like to see phosphodiesterase restored to it original meaning as the general name for all enzymes attacking diesterified phosphate. During the recent past, venom exonuclease has been reviewed several times (15-21). Three books (ZZ-24) devoted to nucleases discuss venom exonuclease. The distinction between exonucleases and endonuclease was first formulated in our laboratory (11, 25). The originally erroneous designation of the terminus has been corrected (26-29). 10. H. A. Haessler and L. Cunningham, Exptl. Cell Res. 13, 304 (1957). 11. M. Laskowski, Sr., G. Hagerty, and U.-R. Laurila, Nature 180, 1181 (1957). 12. J. G. Georgatsos and M. Laskowski, Sr., Biochemistry 1, 288 (1962). 13. E. Sulkowski, W. Bjork, and M. Laskowski, Sr., JBC 238, 2477 (1963). 14. L. Brisbois, N. Rabinovitch-Mahler, P. Delori, and L. Grillo, J . Chromatog. 37, 463 (1968). 14a. G. Pfleiderer and F. Ortanderl, Biochem. 2.337, 431 (1963). 15. G. C. Butler, “Methods in Enzymology,” Vol. 2, p. 561, 1965. 16. M. Laskowski, Sr., Ann. N . Y . Acad. Sci. 81, 776 (1959). 17. H. G. Khorana, “The Enzymes,” 2nd ed., Vol. 5, p. 79, 1961. 18. W. E. Razzell, “Methods in Enzymology,” Vol. 6, 236, 1963. 19. M. Laskowski, Sr., Procedures Nucleic Acid Res. p. 154 (1966). 20. M. Laskowski, Sr., Advan. Enzynol. 29, 165 (1967). 21. W. Bjork, Acta Univ. Upsaliensis, Abstr. Uppsala Dissertations Sci. 106 (1967). 22. M. Privat de Garilhe, “Les Nuclkases.” Hermann, Paris, 1964 (in French). 23. M. Privat de Garilhe, “Enzymes in Nucleic Acid Research.” Hermann, Paris, 1967 (in English). 24. V. S. Shapot, “Nukleazy.” Meditzina, Moskva, 1968 (in Russian).

13. VENOM EXONUCLEASE

315

It is now clear that venom exonuclease attacks all polynucleotide chains from the w terminus regardless of the location of the monophosphoryl group (see below). During the past few years, a t the suggestion of Dr. Waldo E. Cohn we have used a to designate the 5' terminus and w for the 3' terminus. The Greek letter has an advantage over the symbol 5', when the terminal nucleotide is to be isolated either as N u or pN'p (a-nucleoside or (~-3~~5'nucleosidediphosphate instead of the alternative description as 5'terminal nucleoside or $-terminal 3',5'-nucleosidediphosphate, which is more cumbersome). I n this connection the reviewer would like to suggest the name of o-exonucleases for the type of enzyme exemplified by venom exonuclease and a-exonucleases for the type of enzyme exemplified by spleen exonuclease, as substitutes for types I and I1 of Razzell (18). This nomenclature is particularly useful in case of nucleases with mixed endo-, exonucleolytic properties as well as mixed mono-, diesterase properties. For example, mung bean nuclease I which was purified as an endonuclease, and thus diesterase, is inseparable from the w-monophosphatase activity (5052); both activities are probably intrinsic properties of the same molecule. An enzyme with similar properties (53) has been previously observed among exonucleases of E . coli (see Chapter 10 by Lehman, this volume). Presumably many more enzymes of this type will be found. Among the presently known nucleases only a few exhibit the W-monophosphatase activity. Venom exonuclease which belongs to the group of W-exonucleases does not have this property (see Section IV,B) . It has been known for some time (16)that venom exonuclease produces only 5'-mononucleotides and that it attacks both deoxyribonucleic acid (DNA) and ribonucleic acid (RNA). Recently, however, it was shown that it also attacks derivatives of arabinose (30,34, 36) and

25. 26. 27. 28. 29.

M. Privat de Garilhe and M. Laskowski, Sr., JBC 223, 661 (1955). M. F. Singer, R. J. Hilmoe, and L. A. Heppel, Federation Proc. 17, 312 (1958). W. E. Razzell and H. G. Khorana, JACS 80, 17770 (1958). W. E. Razzell and H. G. Khorana, JBC 234, 2114 (1959). G. M. Richards and M. Laskowski, Sr., Biochemistry 8, 1786 (1969).

30. W. J. Wechter, A. J. Mikulski, and M. Laskowski, Sr., BBRC 30, 318 (1968). 31. P. H. Johnson and M. Laskowski, Sr., JBC 245, 891 (1970). 32. A. J. Mikulski, P. H. Johnson, and M. Laskowski, Sr., Federation Proc. 25, 906 (1970). 33. C. C. Richardson, I. R. Lehman, and A. Kornberg, JBC 239, 251 (1964). 34. W. J. Wechter, J . M e d . Chem. 10, 762 (1967). 35. G. M. Richards, D. J. Tutas, W. J. Wechter, and M. Laskowski, Sr., Biochemistry 6, 2908 (1967).

316

M. LASKOWSKI, SR.

thus appears to be “totally blind” to sugar. Historically, venom exonuclease was considered to have a strong preference for short monostranded oligonucleotides. I n spite of claims to the contrary (%), it is capable of attacking native DNA (11, 37-40). The most important and as yet unanswered question with regard to venom exonuclease is concerned with its endonucleolytic activity. Is this activity a result of contamination with the known venom endonuclease (8, 10-12) or contamination with an as yet unidentified endonuclease, or is it an intrinsic property of the enzyme molecule? Traces of contaminating venom endonuclease, which is a #-monoester former (12) can undoubtedly be identified in all presently available preparations. There are, however, some facts suggesting the presence of an endonucleolytic activity that is distinct from the known venom endonuclease. Tener et al. (41) and Razzell and Khorana (28) found that cyclo-pTpT, cyclo-pTpTpT, and cyclo-pTpTpTpT were all hydrolyzed at an identical, very slow rate, and the only product was pT. No linear oligonucleotide was detected as an intermediate. Should the known endonuclease as a contaminant have been responsible for the opening of the ring, a compound having a 3’-monophosphoryl group would have been detected. Bjork (42) subjected phage T2 DNA to the action of deoxyribonuclease (DNaseI) and obtained a mixture of fragments containing glucosylated hydroxymethyl cytosine. This mixture was then used as substrate for repeated and exhaustive digestion with venom exonuclease. Products containing oligonucleotides were analyzed for glucose to phosphate ratio. It was assumed that glucosylated hydroxymethyl cytosine completely blocks the exonucleolytic action of exonuclease. From the composition of the oligonucleotides obtained after repeated exhaustive digestion, it was concluded that the endonucleolytic action of venom exonuclease requires a sequence of a minimum of three glucose-free nucleotides interposed between the glucose-containing units. Unquestionably the activity observed by Khorana et al. (28, 41) and by Bjork (42) must be ascribed to an enzyme that is a 5’-monoester former. The question as to whether or not this activity is an intrinsic 36. I. I. Nikolskaya, N. M. Shalina, and T. I. Tikhonenko, BBA 91, 354 (1964). 37. H. G. Boman and U. Kaletta, Nature 178, 1394 (1956). 38. H. G. Boman and U. Kaletta, BBA 24, 619 (1957). 39. J. Adler, I. R. Lehman, M. D. Bessman, E. S. Sims, and A. Kornberg, Proc. Natl. Acad. Sci. U.S. 44, 641 (1958). 40. E. J. Williams, S.-C. Sung, and M. Laskowski, Sr., JBC 236, 1130 (1961). 41. G. M. Tener, H. G. Khorana, R. Markham, and E. H. Pol, JACS 80, 6223 (1958). 42. W. Bjork, Arkiv Kemi 27, 515 (1967).

13.

VENOM EXONUCLEASE

317

catalytic property of pure exonuclease cannot be answered a t present. An interesting substance poly (adenosine diphosphate) in which two 5‘monophosphoryl groups are linked through a pyrophosphate bond to form a chain of not more than 25 oligomers has been recently used as substrate. The enzyme was commercial venom exonuclease that was further purified by passage through a column of Dowex 50W-X8. Matsubara e t al. (@a) concluded that in this system venom exonuclease had endonucleolytic type of action.

II. Chemical Nature of the Enzyme

Very little is known about the chemical nature and physical properties of venom exonuclease; we simply do not have a preparation pure enough to warrant such studies. This regrettable state of affairs is caused in part by the high price of the starting material. No laboratory can afford a macroscale purification procedure. Therefore, the major aim up to the present time has been to obtain a preparation free from the contaminants that interfere with a specific use of the enzyme. This approach resulted in a number of preparations that varied not only in specific activity of exonuclease but also in the nature, and quantity of contaminants. The criteria used to describe these remaining contaminating activities vary. Such statements as “below the level of detection” are helpful only if the level is specified. Unfortunately, this is not always the case. Several methods for preparation of purified venom exonuclease have been described [see the reviews on methods (15,18, 19, 21) and books (22-24) 1. The major effort of purification was directed toward removing the contaminating monophosphatases. A successful and widely used step was introduced by Sinsheimer and Koerner ( 4 3 ) . At p H 4, monophosphatases are precipitated with a lower concentration of acetone; the remaining exonuclease is precipitated by a higher acetone concentration. Several modifications of this principle have been proposed (concentration of acetone, temperature during precipitation, etc.) , Commercially available preparations represent essentially this stage and contain per unit of 5’-nucleotidase, unit of nonspecific unit of exonuclease: phosphatase, and about the same amount of endonuclease. The last figure is only an approximation because of the difficulty of accurate determination. The more elaborate preparations have these contaminants 42a. H. Matsubara, S. Hasegawa, S. Fujimura, T. Shima, T. SugimurtL and M. Futai, JBC 245, 3606 (1970). 43. R. L. Sinsheimer and J. F. Koerner, JBC 198, 293 (1952).

318

M. LASKOWSKI, SR.

reduced by about two orders of magnitude. Recently significant progress was made (43a). Monophosphatases may be selectively inactivated by thermal denaturation below pH 4. Since the acetone precipitation step is not applicable to all venoms, the decision of which method to use depends on the availability of the starting venom: Crotalus adamanteus (35), Bothrops atrox ( 4 4 ), Hemachatus haemachates (45’), and Vipera lebetina ( 4 5 ) . The best that can be done a t present to characterize venom exonuclease is to state that it is a colorless protein, presumably having no prosthetic group. However, the absence of carbohydrate has not been definitely established. The enzyme is heat labile, and it is not significantly retarded on Sephadex G-100 (4.4), suggesting a comparatively large molecular weight. Either exonuclease itself (or a protein t o which it is strongly bound) behaves as a basic protein because it is readily absorbed on CM-cellulose and only slightly on DEAE-cellulose (4,46). Exonuclease activity peaked a t pH 8.7 when the whole venom of cobra (Naja naja atra) was subjected to electrofocusing ( 4 7 ) . Although virtually nothing is known concerning the detailed structure of the exonuclease molecule, speculations about its active center have been made (48, 49). Brown and Bowles (48) concluded that venom exonuclease requires the following for activity: intact tryptophan and tyrosine residues, SH groups, and S-S bridges. Wigler ( 4 9 ) ,on the basis of kinetic data, concluded that the active center contains two closely located acidic groups. The presence of acidic groups in the vicinity of the active center serves to explain the recent results of Richards and Laskowski (30, 5 0 ) , who found that a double negative charge on the 3’-monophosphoryl group confers an unusual resistance on this type of compound to exonuclease. Suggestions concerning the structural element to which the enzyme binds vary. Razzell and Khorana ( 5 1 ) , on the basis of minimal requirements for activity, considered the binding site identical with the hydrolytic site, which is limited to one doubly esterified 5’-nucleotide. Bjork (42) believed that besides a hydrolytic site additional binding sites exist. I n this connection it should be mentioned that whereas a #-mono43a. E. Sulkowski and M. Laskowski, Sr., Federation Pi-oc. (in press). 44. W.Bjork, JBC 238, 2487 (1963). 45. I. I. Nikolskaya, N. M. Shalina, and E. I. Budowski, BBA 64, 197 (1962). 46. F. Felix, J. L. Potter, and M. Laskowski, Sr., JBC 235, 1150 (1960). 47. J. Simon, L. Brisbois, and L. Grillo, J . Chromatog. 44, 209 (1969). 48. J. H.Brown and M. E. Bowles, U.S. A m g Med. Res. Lab., Fort Knox, Ky., R e p t . 627, 11 (1965); CA 15178b (1965). 49. P. W.Wigler, JBC 238, 1767 (1963). 50. G. M. Richards and M. Laskowski, Sr., Bbchemistm 8, 4858 (1969). 51. W. E.Razzell and H. G . Khorana, JBC 234, 2106 (1959).

13. VENOM

319

EXONUCLEASE

nucleotide is a minimal requirement for a good substrate, it is not an absolute requirement since di-p-dinitrophenyl phosphate is slowly hydrolyzed. Several years ago, Razzell and Khorana ( 5 1 ) ,using dinucleosidemonophosphates of deoxythymidylic acid, showed that the 5‘+5‘ linkage was hydrolyzed slightly faster than the normal 3 ’ 4 ’ linkage. With nucleosides heterogeneous with respect to sugar, both 2‘+5’ and 5‘+5‘ linkages were shown to be hydrolyzed slightly slower than the normal 3‘+5’ linkages (35). In Khorana’s and in our laboratories the differences were comparatively small. I n the case of the 5’+5’-linked nucleosides with different sugars, the cleavage occurred in such a manner that phosphate remained on a nucleoside containing sugar other than arabinose (34, 36) ’ 111. Structural Characteristics of Substrates Affecting Susceptibility

A. CONFORMATION In spite of some claims to the contrary, venom exonuclease is capable of attacking double-stranded high molecular DNA. I n fact, doublestranded DNA is a better substrate than denatured DNA. Bjork (@) studied the rates of degradation of native and heat-denatured DNA using a pH stat. Denatured DNA was degraded a t a steady rate, which was dependent on the ionic strength of the medium. An increase in NaCl concentration from 1 to 100 m M decreased the rate of hydrolysis by a factor of two. With native DNA a two-phase reaction was observed. The initial, very rapid, rate was independent of NaCl concentration. After about one-third of the linkages had been hydrolyzed, the rate slowed down to that of denatured DNA and became salt dependent. Similar biphasic kinetics was observed previously with DNA that was denatured by an exhaustive dialysis (40). With RNA as substrate the results are conflicting. Cousin (52) observed that poly I, poly U, and poly C were rapidly and completely hydrolyzed, whereas under identical conditions poly I poly C was hydrolyzed slowly and incompletely. Native rRNA and tRNA were resistant to 60 pg/ml of commercial exonuclease, but after heat denaturation rRNA was readily and tRNA partly digested (52). Nihei and Cantoni (53) digested tRNA to completion, as did Keller (64) who observed a two-phase reaction. In agreement with the last two authors (53, 54) but contrary to the results of Cousin ( 6 2 ) , Hadjiolov et al.

+

52. M. Cousin, Bull. SOC.Chim. Bwl. 45, 1363 (1963). 53. T. Nihei and G. L. Cantoni, JBC 238, 3991 (1963). 54. E. B. Keller, BBRC 17, 412 (1986).

320

M. LASKOWSKI, SR.

(65) found that poly U was hydrolyzed faster than poly A, whereas an equimolar mixture of poly A and poly U forming a double-stranded structure was hydrolyzed a t a rate that was halfway between the previous two. Heat denaturation did not affect the rate of hydrolysis of rRNA. It would appear, therefore, that with RNA as substrate, conformation exerts no effect on the rate of hydrolysis.

B. THENATUREOF SUGAR I t has been known for many years (15-24) that venom exonuclease is capable of hydrolyzing both DNA and RNA. Gray and Lane (56) showed that naturally occurring 2'-O-methyl-substituted ribose derivatives are also hydrolyzed, even though the rate of hydrolysis is slower than with the unsubstituted ribose. Interestingly, the 2'-O-methylated derivatives are totally resistant to 5'-nucleotidase (57, 5 8 ) . Wechter (34) and Richards et al. (35) showed that venom exonuclease is capable of hydrolyzing dinucleoside monophosphates with one or both nucleosides containing arabinose. Even though only four different sugars have been tested, it would appear that venom exonuclease is totally blind to sugar, a t least in the qualitative sense. Other enzymes capable of hydrolyzing DNA and RNA are incapable of hydrolyzing derivatives of arabinose, e.g., micrococcal nuclease (30). Quantitative differences between susceptibility of compounds containing different sugars exist. Richards et al. (35) made a systematic study of a number of dinucleosidemonophosphates (N"pN0) in which either the CY- or P-nucleoside contained arabinose, while the other nucleoside contained ribose or deoxyribose. All compounds were found susceptible to venom exonuclease. The surprising finding was that the extremes of differences in the values of V , and K,,, lay in a comparatively narrow range of 20-fold.

C. THENATUREOF BASES Venom exonuclease has no pronounced preference (or discrimination) toward any of the four common bases of DNA or RNA. The discrimina55. A. A. Hadjiolov, P. V. Venkov, L. B. Dolapchiev, and D. D. Genchev, BBA 142, 111 (1967). 56. M. W.Gray and B. G. Lane, BBA 134, 243 (1967). 57. M. Honjo, Y. Kanai, Y. Furukawa, Y. Mizuno, and Y. Sanno, BBA 87, 696 (1964). 58. B. G . Lane, Biochemistry 4, 212 (1965).

13.

VENOM EXONUCLEASE

321

tion against glucosylated hydroxymethyl cytosine has been suspected for a long time because these compounds were resistant to the combined action of DNase I and venom exonuclease (59, 60). Recent work of Nikolskaya et al. ( G I ) and Bjork (41) established that glucosylated bases blocked the action of venom exonuclease. A good source of uncommon bases is tRNA. It provides substrates for studying the effect of base on the rate of hydrolysis. Baev e t al. (82) showed that N2-dimethylguanylyl- (3'-5') -cytidine-3' phosphate (GZ"pCp) was hydrolyzed much slower than the usual GpCp. Venkstern (63) reported that Tpq was hydrolyzed very slowly. Naylor et al. (64) found that Cpq was hydrolyzed with half the rate of CpU. The same group of workers introduced (64, 6 5 ) a chemical block on uridine and pseudo-uridine residues by reacting RNA with 1-cyclohexyl-3- (2morpho-linyl- (4)-ethyl) -carbodiimide metho-p-toluene sulfonate. The modification of the uridine residues completely blocked the action of venom exonuclease and also blocked the action of pancreatic RNase. The action of venom exonuclease is also blocked by thymine dimers produced as a result of UV irradiation of DNA a t 280 nm. Setlow et al. (G6) subjected irradiated DNA to exhaustive digestion by venom exonuclease. They isolated and identified the products of the reaction, which were composed of large amounts of all four 5'-mononucleotides and of small amounts of trinucleotides of the type d-pNpTTT where N was any of four common nucleosides and TpT was the irradiation-induced dimer of thymidine. These trinucleotides were totally resistant to further digestion with venom exonuclease but became partially susceptible after UV irradiation at 240 nm, known to reverse dimerization. The authors picture the action of venom exonuclease as proceeding linearly from the 0 terminus until the block is encountered. At this point the enzyme (or an impurity) acts very slowly by splitting the internal bond one base beyond the dimer. From there on conventional hydrolysis is resumed until the next block. This experiment touches upon one of the most pressing problems connected with venom exonuclease. Is the endonucleolytic activity an intrinsic property of the enzyme? 59. R. L. Sinsheimer, Science 120, 551 (1954). 60. E. J. Vokin, JACS 76, 5892 (1954). 61. I. I. Nikolskaya, D. S. Kislina, N. M. Shalina, and T. I. Tikhonenko, Biokhimiya 30, 1236 (1965). 62. A. A. Baev, T. V. Venkstern, A. D. Mirzabekova, and R. I. Tatarskaya, Biokhimiya 28, 931 (1963). 63. T. V. Venkstern, Dokl. Akad. Nauk SSSR 170, 718 (1966). 64. R. Naylor, N. W. Y. Ho, and P. T. Gilham, JACS 87, 4209 (1965). 65. J . C. Lee, N. W. Y. Ho, and P. T. Gilham, BBA 95, 503 (1955). 66. R. B. Setlow, W. L. Carrier, and F. J. Bollum, B B A 91, 446 (1964).

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M. LASKOWSKI, SR.

D. EFFECT OF MONOPHOSPHORYL GROUP The effect of the proximity of the 5’-monophosphoryl group on susceptibility of a dinucleotide to venom exonuclease was recognized quite early (25, 2 8 ) . The removal of the 5’-monophosphoryl group decreased the rate of hydrolysis about 10-fold. The reported extreme values varied from 5- to 20-fold. The influence of the 5‘-monophosphoryl group (or of the lack of this group) is most pronounced with dinucleotides and is easily demonstrable with tri- and tetranucleotides. Intuitively one expects the effect to diminish with increasing chain length; indeed, with a homologous series of dephosphorylated compounds U (pU) Dolapchiev (67) showed that the longer the chain the more susceptible it becomes t o exonuclease. This finding indirectly supports Bj ork’s hypothesis regarding the existence of multiple sites of attachment of substrate to enzyme. The absence of the 5’-monophosphoryl group is felt by the enzyme even though the hydrolytic site is removed from the ry terminus by several monomers. Derivatives bearing a 3‘-monophosphoryl group were originally classified as totally resistant to venom exonuclease. As the quality of the enzyme preparation improved, these compounds were found susceptible but required 1000-fold more enzyme than was needed to hydrolyze 5’-monophosphate-bearing compounds. This unusual resistance led to another erroneous conclusion, that the polarity of exonuclease ehanges ( 2 Q ) .The basis for this belief were the experiments in which a mixture of tri-, tetra-, and pentanucleotides of the type d-NapN@pN7pNJpwere used as substrates. The early products were nucleosides and nucleotides, whereas 3’, 5’-mononucleoside diphosphates appeared considerably later. It is clear now that the mixture was contaminated with a small amount of dephosphorylated chains which were rapidly hydrolyzed to completion. The accompanying tabulation represents an approximation of several semiquantitative determination from different laboratories. These data %,

Compound

Relative amount of enzymes required for hydrolysis

d-pNupN@ d-N‘pN@ d-NapNap

1 10 1000 ~~

67. L. B. Dolapchiev, FEBS Letters 2, 185 (1969)

13.

323

VENOM EXONUCLEASE

were obtained a t pH 9, which is known to be an optimum for venom exonuclease attacking chains bearing 5'-monophosphate. Under the conditions in which the determinations were made these values are still essentially valid. However, it was recently found (29) that decreasing pH (and thus the negative charge on 3'-monophosphate) decreases the resistance of these compounds to venom exonuclease. Figure 1 shows the dependence of the rate of hydrolysis on pH. A change from pH 9 to 6 produces an increase in the rate of hydrolysis to 125% with ribonucleotides and a 500% increase with deoxyribonucleotides. This also means that the quantity of enzyme required to accomplish a complete hydrolysis of deoxyribodinucleotides is reduced by a factor of 5 (from lo00 to 200 relative units ; see the accompanying tabulation). In Fig. 2 the ratio of the rates of hydrolysis of phosphorylated to that of dephosphorylated dinucleotides is plotted vs. pH. The ordinate is on the logarithmic scale. The ratio changes from about 40 a t pH 5 to about 1700 at pH 9. [The previous estimate of the latter value (as 1000/10 = 100 in the tabulation) was an underestimate.] The lower curve of Fig. 2 compares the analogous ratio except that a ribodinucleotide terminated in 2',3'-cyclic phosphate serves as substrate. In this case the ratio decreases slightly when the pH is changed

0

E

z

25

-

20

-

t5-

.c 40 2

5 -

I

I

I

I

I

I

1

4

5

6

7

8

9

PH

FIG.1. Effect of pH on the hydrolysis of mixed-base 3'-phosphorylated ribodinucleotides ( 0 )and deoxyribodinucleotides (A). For ribonucleotides reaction mixtures (total volume 0.063 ml) were incubated for 1 hr a t 37" in sealed capillary tubes. Substrate Ant = 65, [enzyme] = 4.2 units/ml, [buffer] = 0.1 M (tris succinate), [MgCl,] = 1.0 mM. After electrophoresis on paper for 0.5 hr, 23 V/cm, pH 7.0, amounts of N were determined as percent of total material ( A n ] )recovered from the paper. With deoxyribodinucleotides [substrate] = 83 A,?,, [enzyme] = 5.1 units/ml; [buffer] and [MgCl,] as above. Reprinted from Richards and Laskowski (60). Copyright (1969) by the American Chemical Society. Reprinted by permission of the copyright owner.

324

M. LASHOWSKI, SR.

u)

2

e

.I -

4000300 -

>,

P R r

I00

-

30

-

v-

._ o c

B

I0 -

0

3 I '

4

I

I

1

I

L

5

9

7

0

9

PH

FIQ.2. Effect of removing negatively charged groups from the w end of a dinucleotide at several values of pH. The data were compiled from large number of experiments. ( 0 ) Calculated from the rate of appearance of nucleosides (d-Nu) ; ( A ) calculated from the rate of appearance of the &terminal unit, be it d-pNe (from d-NapN8) or d-pN@p (from d-NepNPp). The same system is used for A'pA8 and AapAR > p. Reprinted from Richards and Laskowski ($9).Copyright (1969) by the American Chemical Society. Reprinted by permission of the copyright owner.

from 6 to 9. It must be borne in mind that with both ApA > p and ApA the optimal rate of hydrolysis is a t pH 9. On the basis of these findings the experiments with deoxyribosetrinucleotides were performed (Fig. 3). The results clearly show that throughout the whole range of pH values an intermediate d-NapNP accumulated as d-pNyp was liberated. The experiment convincingly shows that no change in polarity in the action of venom exonuclease occurs with substrates bearing 3'-monophosphoryl group. The same W+(Y sequence of attacks occurs with all substrates regardless of termination.

IV. Venom Exonuclease as a Tool for Structural Determination

A.

SEQUENCE IN

RIBOOLIGONUCLEOTIDES

An elegant method of determination of sequence using monophosphatase, exonuclease, and alkaline hydrolysis has been proposed by

13.

325

VENOM EXONUCLEASE

d-NpN

5-

b a -

4-

-6

3 -

0 + +

r-

8 2 -

d-N

I X

4

5

6

7

8

9

PH

FIG.3. Effect of pH on the hydrolysis of purified mixed deoxyribotrinucleotides (dd-pN@,and ( 0 ) NnpN@pN'p)by venom exonuclease. ( A ) d-pNnp, ( X ) d-NapN8/2, (0) d-Na. Unhydrolyzed d-NapNBpNTp is not shown but is included in the total Am. Substrate = 83 A 27 1 , [enzyme] = 0.525 unit/ml for pH 4.0, 0.210 unit/ml for pH 5.0 and 9, and 0.105 unit/ml for pH 6-8. Electrophoresis for 2 hr, pH 6. Observed values for (percent of total A271) of the separated products were divided by 5.2 or 1 to compensate for the different amounts of enzyme used. Dashed line shows values for mononucleotides without such correction. Values of percent of total A m for d-NnpN@were divided by 2 to facilitate comparison with other products on a molar basis. Reprinted from Richards and Laskowski (29). Copyright (1969) by the American Chemical Society. Reprinted by permission of the copyright owner.

Holley et al. (68). A ribooligonucleotide is first dephosphorylated with rnonophosphatase. Its composition is established by a complete hydrolysis with KOH, which also defines the m-terminal nucleoside. The rest of the dcphosphorylated compound is subjected to a mild, incomplete hydrolysis with venom exonuclease. The mixture is then chromatographed on DEAE-cellulose according to the method of Tomlinson and Tener (69), giving a series of homologs each longer by one monomer than the preceding compound. The material of each peak is collected separately and hydrolyzed completely with KOH, giving a mixture of mononucleotides and one nucleoside representing the o terminus. The nucleoside in each peak is identified and this establishes the sequence. The method could be extended to deoxyribose derivatives by substi68. R. W. Holley, J. T. Madison, and A. Zamir, BBRC 17, 3846 (1964). 69. R. V. Tomlinson and G. M. Tener, Biochemktq/ 2, 697 (1963).

326

M. LASKOWSKI, SR.

tuting an enzyme for KOH to accomplish the complete degradation in order to obtain the W-deoxynucleoside. Until now this could have been done either with the aid of spleen a-exonuclease or Lactobacillus acidophilus a-exonuclease (70). Since the first enzyme is known to contain some monophosphatase, the risk of obtaining a false nucleoside existed, particularly with the longer oligonucleotides. The second enzyme also has limitation, it attacks only short oligonucleotides. Recently ( 7 I ) , it was found that micrococcal nuclease in extremely high concentration is capable of degrading oligomers to 3’-mononucleotides. Thus, an alternative enzymic degradation may be tried. I n the case of micrococcal nuclease the contamination with monophosphatase is insignificant.

B. IDENTIFICATION OF a A N D w TERMINALS Venom exonuclease has been widely used for the determination of the CY terminus in oligonucleotides. The normal procedure was to divide the dephosphorylated sample into two parts and digest one part with venom exonuclease. The terminus appeared as a nucleoside; the rest of the chain was degraded to 5’-mononucleotides. The other part of the dephosphorylated chain was degraded with spleen a-exonuclease. In this case the w terminus appeared as a nucleoside, the rest of the chain as 3’-mononucleotides. As the quality of venom exonuclease improved, more difficult tasks were tackled. The LY terminus was identified in tobacco mosaic virus (72). It required finding one nucleoside among 6400 nucleotides. Venom cxonuclease was also used for identification of both terminals in chains bearing 3’-monophosphate (IS,73). The CY terminus appears as a nucleoside, the w terminus as a nucleoside-3’,5’-diphosphate.The method did not gain a wide application because it required large amounts of highly purified enzyme. The recent finding that 3’-monophosphates are better substrates a t pH 6 than 9 (29) is likely to increase the use of this method. Roblin (74) used venom exonuclease to identify the a terminus in RNA isolated from bacteriophage R-17. The RNA was degraded with alkali and the mixture chromatographed according to Tomlinson and Tener ( 6 9 ) . The terminal appeared at the place in the pattern that corre(Y

70. W. Fiera and H. G . Khorana, JBC 238, 2789 (1963). 71. A. J. Mikulski, E. Sulkowski, L. Stasiuk, and M. Laskowski, Sr., JBC 244, 6559 (1969). 72. T. Sugiyama and H. Fraenkel-Conrat, Biochemistry 2, 332 (1963). 73. S. Venecko and M. Laskowski, Sr., BBA 61, 547 (1962). 74. R. Roblin, J M B 31, 51 (1968).

13. VENOM

327

EXONUCLEASE

sponded to tetranucleotides and had a spectrum of G. The amount of venom exonuclease required to produce a complete hydrolysis of G T P was established in a control experiment.

-

exonuclease

PPPG

faat

PG

+ PP

A fivefold amount of exonuclease was then used on the isolated derivative, and only three-fourths of it was hydrolyzed. The slow hydrolysis was expected because of the presence of 3‘-monophosphate. The product of this reaction, pGp, was identified by comparison with an authentic sample. Both reactions were run a t p H 9.0. Should a p H of 6 be used

-

exonucleaae

PPPGP

slow

PGP

+ PP

in the second reaction the hydrolysis would have been complete. In conjunction with other enzymes DNase I, phosphatase, polynucleotide kinase, DNA ligase, and another nuclease, venom exonuclease, have been used for identification of nicks inflicted by DNase I. Essentially the same method identifies the (Y terminus in a very long polynucleotide. The schematic representation of events preceding venom exonuclease is shown in Figs. 4 and 6 in Chapter 12, by Laskowski, Sr., this volume. From the schemes it is obvious that the simplified problem of determining only the terminus requires phosphatase and polynucleotide kinase, but no ligase. Upon a complete digestion to 5’-mononucleotides (venom exonuclease) the only radioactive nucleotide is the a! terminal. The contamination by venom endonuclease has no effect on identification. However, the contamination with 5’-nucleotidase leads to partial dephosphorylation and a false estimate of the chain length. Venom exonuclease was also used to determine the o terminus and its adjacent four nucleotides in tobacco mosaic virus ( 7 6 ) .The sequence was determined from the rate of appearance of monomers. In passing, it may be mentioned that o-exonuclease activity presumably simiIar to venom exonuclease has been studied histochemically in a variety of tissues. Several substrates have been developed specifically for this purpose (76-79). The newest addition to the family of substrates (Y

75. B. Singer and H. Fraenkel-Conrat, BBA 72, 534 (1963). 76. H. Sierakowska, H. Szemplinska, and D. Shugar, Acta Biochim. Polon. 10, 399 (1963).

77. H. Sierakowska and D. Shugar, BBRC 11, 70 (1963). 78. J. P. Horwitz, C. V. Easwaran, P. L. Wolf, and L. S. Kowalczyk, BBA 185, 143 (1969).

79. M. Erecinska, H. Sierakowska, and D. Shugar, European J . Biochem. 11, 465 (1969).

328

M. LASKOWSKI, SR.

useful for distinguishing a- from o-exonuclease are 2,Cdinitrophenyl esters of 3‘- and 5’-phosphates of thymidine (80). A comprehensive review (81) on histochemical aspects has been published.

V. Other Venom Enzymes That Hydrolyze Phosphate Esters

Two monophosphatases of venom are known (31). Both have pH optima of about 9 and thus represent dangerous contaminants when exonuclease is used to identify a nucleoside among a large number of mononucleotides. One of the monophosphatases, 5’-nucleotidase, is quite specific and attacks only 5’-rnononucleotides. This enzyme has been used to determine the amount of mononucleotides in a mixture of 5’-monophosphates of various chain length. It was also used to distinguish between 3‘- and 5’-mononucleotides in a mixture of both. Venom endonuclease is ’#-monoester former with a preference for the Gp-G bond and an optimal pH of 5 (1%’).It represents the most dangerous contaminant of exonuclease in experiments designed for the identification of terminals in long chains bearing 3‘-monophosphates because it leads to false terminals. More details concerning the properties of these contaminating enzymes may be found in reviews (17-21)and books (22-24) devoted to nucleases. ACHNOWLEDQMENTS The experimental work referred to in this article and performed in our laboratory was generously supported by the American Cancer Society, PRP-30 and E157; by the U. S. Atomic Energy Commission, AT(30-1)3630; and by the National Science Foundation, GB-6058. The author is indebted to Drs. Barnard, Heppel, Holley, Lehman, Tener, and Weinfeld for the critical reading of the manuscript.

80. R. G.von Tigerstrom and M. Smith, Bwchemtstry 8, 3067 (1969). 81. D.Shugar and H. Sierakowska, Prog. Nucleic Acid Res. 7, 369 (1967).

Spleen A cid Exonuclease ALBERT0 BERNARDI

GIORGIO BERNARDI

I. Introduction . . . . . . . . . . 11. Isolation, Purity, and Physical Properties . . . 111. Catalytic Properties . . . . . . . . A. Activity on Natural Substrates . . . . B. Activity on Artificial Substrates . . . . C. pH-Activity Curves-Activators and Inhibitors IV. Distribution and Intracellular Localization . . .

. . . .

.

.

.

.

.

. . .

. . . .

.

.

.

.

.

329 330 331 331 333 334 336

I. Introduction

Spleen acid exonuclease is an enzyme particularly useful in sequence studies of oligonucleotides, derived from both ribonucleic acid and deoxyribonucleic acid, since it splits off, in a sequential way, nucleoside3’-phosphates starting from the 5’OH end. The enzyme has also been called spleen phosphodiesterase (I, 2 ) and phosphodiesterase I1 ( 3 ) .We prefer to use the term phosphodiesteruse as a general name for the broad group of enzymes hydrolyzing phosphodiester bonds whether between nucleosides or not ( 4 ) . Table I gives a few examples of such enzymes. The term phosphodiesterase II (S), intended to mean an enzyme releasing nucleoside-3’-phosphates, seems to be an unhappy one, like that of deoxyribonuclease I1 (6) from 1. L. A. Heppel, R. Markham, and R. J. Hilmoe, Nature 171, 1152 (1953). 2. D. M. Brown, L. A. Heppel, and R. J. Hilmoe, J . Chem. SOC.p. 40 (1954). 3. W. E. Razzell, Experientia 23, 321 (1967). 4. G. Schmidt and M. Laskowski, “The Enzymes,” 2nd ed., Vol. 5, p. 1, 1961. 5. G. Bernardi, “The Enzymes,” 3rd ed., Vol. IV, p. 271, 1971.

329

330

A. BERNARDI AND G. BERNARDI

which i t is derived. I n fact, to mention just one of several criticisms which can be raised, roman numerals are more and more used to indicate new enzymes both of bacterial (6) and animal ( 7 ) origin and have little bearing on the position of phosphate in the products of digestion. An alternative, shorter, nomenclature for the enzyme is spleen exonuclease. After the initial and fundamental work of Heppel and Hilmoe, and Razzell and Khorana, already reviewed in the previous edition of “The Enzymes” (8) and in two other articles (3,9 ) , the major advances have been the preparation of spleen exonuclease in a very highly purified form (10, 11), and the recognition that the enzyme has no endonucleolytic activity and that it can attack oligonucleotides carrying a terminal phosphate in the 5’ position (12) ; this represents, however, a strong ratelimiting step.

It. Isolation, Purity, and Physical Properties

A method for the partial purification of spleen exonuclease was described by Heppel and Hilmoe in 1955 (IS) and by Hilmoe in 1960 (14) ; this was later improved by Razzell and Khorana (15) and Richardson and Kornberg (16). I n 1966, we described a novel purification procedure (10) leading to an enzyme preparation with a specific activity comparable to that of the best preparation of Razzell and Khorana (15). Enzyme yields were, however, low; the method was therefore modified and satisfactory results were obtained (11). The new method involves the preparation of a crude enzyme obtained essentially as in the case of acid deoxyribonuclease (6, 1 7 ) . The main differences are that acidification to pH 2.5 is avoided and (NH,),SO, fractionation is done between 35 and 60% saturation. The crude enzyme is then purified by chroma6. I. R. Lehman, Procedures Nucleic Acid Res. 2, 84 (1963). 7. T. Lindahl, J. A. Gally, and G. Edelman, Proc. Natl. Acad. X c i . U. S. 62, 597 (1969). 8. H. G. Khorana, ‘The Enzymes,” 2nd ed., Vol. 5, p. 79, 1961. 9. W.E. Razzell, “Methods in Enzymology,” Vol. 6, p. 230, 1963. 10. G. Bernardi and A. Bernardi, Procedures Nuclek Acid Res. p. 144 (1966). 11. A. Bernardi and G . Bernardi, BBA 155, 360 (1%). 12. A. Bernardi and G. L. Cantoni, JBC 244, 1468 (1969). 13. L. A. Heppel and R. J. Hilmoe, “Methods in Enzymology,” Vol. 2, p. 565, 1955. 14. R.J. Hilmoe, JBC 235, 2117 (1960). 15. W.E.Razzell and H. G . Khorana, JBC 236, 1144 (1961). 16. C. C. Richardson and A. Kornberg, JBC 239, 242 (1964). 17. G. Bernardi, A. Bernardi, and A. Chersi, BBA 120, 1 (1966).

14.

SPLEEN ACID EXONUCLEASE

331

tography on CM-Sephadex, hydroxyapatite, Sephadex G-75, and rechromatography on hydroxyapatite. The final product may be freezedried with only a small loss in activity. The enzyme is usually stored as a frozen solution or in 50% glycerol a t -15". The enzyme obtained by this purification procedure (11), when tested for contaminants under very stringent conditions, was found to be completely free from phosphatase, DNase, ribonuclease, and adenosine deaminase activities. The sedimentation coefficient of spleen exonuclease, measured by centrifugation in a sucrose density gradient, using cytochrome c as the reference protein, was 4.6 S (11). The enzyme is eluted from Sephadex G-100 between acid phosphomonoesterase (s = 5.6 S) and acid DNase (s = 3.4 S ) . The thermal inactivation curve of the enzyme in 0.15 M acetate buffer0.01 M ethylenediaminetetraacetate (EDTA) , pH 5.0, showed that a 50% inactivation was obtained by heating for 20 min a t 56" (11).


A. ACTIVITYON NATURAL SUBSTRATES 1. Mechanism of Action

Spleen exonuclease is active on the 5'-OH oligonucleotides of both the rib0 and the deoxyribo series. These are sequentially split from the 5'-OH end with formation of 3'-mononucleotides. It has been suggested that an enzyme-product intermediate may exist in the form of nucleoside-3'phosphoryl-enzyme complex (3)since transfer of nucleoside-3'-phosphate to available 5'-hydroxyl functions (or other alcoholic functions) occurs a t high substrate (or acceptor) concentrations (15,18). Bernardi and Cantoni (12) have investigated in detail the mechanism of action of spleen exonuclease on tRNA. They showed that a t pH 4.8 the enzyme is practically unable to digest phosphorylated tRNA, whereas it can digest dephosphorylated tRNA perfectly well. I n contrast, at p H 6.2 the enzyme attacks phosphorylated tRNA though a t a slower rate than dephosphorylated tRNA. Since no inorganic phosphate is liberated in the degradation of phosphorylated tRNA, it is likely that in this case the enzyme begins its attack by releasing the terminal nucleotide as a diphosphate. Following the release of labeled serine at the opposite end of 18. L. A. Heppel and P. R. Whitfield, BJ 80, 1, (19%).

332

A. BERNARDI AND G . BERNARDI

the initial enzyme attack permitted to see that exonuclease degrades tRNA molecules “jumping” from one substrate molecule to another; in fact practically no serine is liberated up to 50% digestion, indicating that the enzyme digestion is progressing a t the same rate in all molecules. I n the case of phosphorylated tRNA, clearly the splitting of the terminal nucleotide represents a strong rate-limiting step since, once this obstacle is overcome, the resulting dephosphorylated substrate molecules are rapidly digested to the end and therefore the liberation of labeled serine takes place early in the digestion course. The mechanism of action of spleen exonuclease is similar to that seen for venom exonuclease (19-21)but different from the processive type of attack exhibited by E . coli RNase 11, sheep kidney exonuclease, and polynucleotide phosphorylase (22,23), in which cases each polynucleotide molecule is completely degraded before the enzymes attack a new molecule. The results of Bernardi and Cantoni (12)contradict the previous beliefs that the enzyme has an intrinsic, though weak, endonucleolytic activity (8) and that a phosphate group in a terminal 5’ position makes a polynucleotide chain completely resistant to the enzyme (16, 94, 26). 2. Eflect of Secondary Structure

The enzyme is very sensitive to the secondary structure of the substrate. Native calf thymus DNA is quite resistant to enzymic attack by spleen exonuclease, being split a t less than 4% the rate a t which alkalidenatured DNA is split (11). Long deoxyribonucleotides (average chain length 20-50), which still have complementary double-stranded structure, are rather resistant to the enzyme (26). Some limited results obtained with synthetic polyribonucleotides (11) are rather puzzling since poly C was found to be completely resistant, whereas poly A, poly I, and poly U were degraded a t comparable rates. I n the solvent used (0.15 M acetate buffer401 M EDTA, p H 5.0), poly A and poly C are believed to have 19. J. Preiss, P. Berg, E. J. Ofengand, F. H. Bergmann, and M. Dieckmann, Proc. Natl. Acad. Sn‘. U.S. 45, 319, (1959). 20. W. E. Razzell and H. G. Khorane, JBC 234, 2114 (1959). 21. T. Nihei and G. L. Cantoni, JBC 238, 3991 (1963). 22. G. W. Nossal and M. F. Singer, JBC 243, 913 (1968). 23. C. B. Klee and M. F. Singer, JBC 243, 923 (1968). 24. D. R. Harkness and R. J. Hilmoe, BBRC 0, 293 (1967). 25. L. A. Heppel and J. C. Rabinowitz, Ann. Rev. Bwchem. 27, 613 (1968). 26. C. h a v e , J.-P. Thiery, S. D. Ehrlich, and G. Bernardi, Biochemistry (submitted for publication).

14.


333

a similar double-helical structure with co-parallel strands, whereas poly U is in a disordered configuration (27). 3. Effect of Glucosylation and Other Chemical Modifications

Glucosylated oligonucleotides obtained from T4 phage DNA by acid DNase digestion are resistant to spleen exonuclease (28). It has been reported that acetylation of the 2'-OH groups of tRNA completely inhibits the action of the enzyme, whereas venom exonuclease is not affected (29). The naturally occurring methylation of sugars and bases in tRNA does not seem to hinder the action of spleen exonuclease.

B. ACTIVITY ON ARTIFICIAL SUBSTRATE~ Other substrates for spleen exonuclease are the p-nitrophenyl esters of nucleoside-3'-phosphates and bis (p-nitrophenyl) phosphate, which is split only very slowly. These substrates are also split by enzymes having quite different natural substrates (Table I) (30-37). I n fact, not only phosphodiesterases, in a broad sense, such as acid DNase, micrococcal nuclease, spleen and venom exonucleases, and cyclic phosphodiesterase but also enzymes such as nucleoside phosphoacyl hydrolase and nucleoside polyphosphatase split these substrates. As pointed out by Spahr and Gesteland (367,this may be explained by the fact that these substrates are not true diesters but rather mixed phosphoanhydrides because of the acidic character of the phenolic OH. It is evident that the use of the synthetic substrates, advocated by Razzell (3) as specific substrates for exonucleases, may be very misleading. Table I1 shows the distinctive characters of three spleen enzymes active on bis (p-nitrophenyl) phosphate which are present in the crude extracts from which acid exonuclease is prepared. 27. A. M. Michelson, J. Massoulie, and W. Guschlbauer, Pro,. Nucleic Acid Res. 6, 84 (1967). 28. C. Soave and G. Bernardi, unpublished experiments (1968). 29. D. G. Knorre, N. M. Pustoshilova, and N. M. Teplova, Bwkhimiya 31, 666 (1986). 30. G. Bernardi and M. GriffB, Biochemistry 3, 1419 (1964). 31. P. Cuatrecasas, M. Wilcher, and C. B. Anfinsen, Bwchemwtry 8, 2277 (1969). 32. M. Laskowski, Procedures Nucleic Acid Res. p. 154 (1966). 33. Y. Anraku, Procedures Nucleic Acid Res. p. 130 (1966). 34. M. Laskowski and B. Filipowicz, Bull. SOC.Chim. Bwl. 4 4 1885 (1958). 35. A. Bernardi and G. Bernardi, BBA 155, 371 (1968). 36. P. F. Spahr and R. F. Gesteland, European J . Bwchem. 12, 270 (1970). 37. T. Term and T. Ukita, J . Biochem. (Tokyo) 58, 163 (1986).

334

A. BERNABDI AND G. BERNARD1

TABLE I SOME ENZYMES ACTIVE ON ~NITROPHENYL PHOSPHODIESTERS p-Ni trophenyl derivatives of Enzyme

Ref.

Natural substrate

Acid D N w Micrococcal nuclease Spleen exonuclease

(30) (31)

DNA DNA, RNA

Bis(p-nitrophenyl) 3'5'phosphate Nucleotides Nucleotides

+ + +

5'-OH oligonucleotides Snake venom (3s) 3'-OH oligoexonuclease nucleotides Cyclic (33) Nucleosidef phosphodiesterase 21, 3'-cycli c phosphates Nucleoside (34, 36) ATP, ADP polyphosphatase Nucleoside (36) Aminoacyl Competitive phosphoacyl adenylate inhibition hydrolase Pancreatic (37) ?b phosphodiesterase (16)

+

+ + +

+"

-

+

+

-

+

-

-

+ +

Split with liberation of p-nitrophenyl phosphate.

* Nucleic acids and oligonucleotides are resistant.

It should be pointed out that the successful purification of spleen exonuclease ( 1 1 ) was greatly helped by use of a DNA hydrolyzate produced by spleen acid DNase as the substrate, since the synthetic substrates are nonspecific, and RNA "core" (the water-undialyzable ribooligonucleotides obtained by exhaustive digestion of RNA with pancreatic RNase) is also hydrolyzed by both acid and basic spleen ribonucleases (38,39). Spleen exonuclease is unable t o hydrolyze cyclic phosphates (14).

C. pH-ACTIVITYCURVES-ACTIVATORS AND INHIBITORS Using an acid DNase hydrolyzate as the substrate and 0.1 M succinate and phosphate buffers as the solvents, the pH optimum was found to be close to 5.5; a higher value, between pH 6 and 7, was found in 0.1 M acetate. The addition of 0.02 M Mg2+did not affect very sensibly the pHactivity curves, although a shift to lower values could be detected. These 38. A. Bernardi and G. Bernardi, BBA 129, 23 (1966). 39. M. E. Maver and A. E. Greco, JBC 237, 736 (1962).

14.

336


TABLE I1 PROPERTIES OF SPLENIC ENZYMES ACTIVE ON BIS(~NITROPHENYL) PHOSPHATE’

Properties

1. Sedimentation coefficientb 2. pH optimum”

3. Substrates Bis(p-nitrophenyl) phosphate p-Nitrophenyl eaters of thymidine5’-phosphate thymidine3’-phosphate Native DNA ATP, ADP, etc. 3’-Phosphate oligonucleotides 4. Inhibitors’ HPOP

sod*-

Polyribonucleotides 5. Thermal inactivation (50%)

6. Chromatographic properties DEAE-Sephadex (pH 6.8) Hydroxyapatite (pH 6.8) CM-Sephadex 7. Kinetica (V vs. S)

Nucleoside polyphosphatase

Acid DNase

Exonuclease

3.4 5.6-5.9

5.8

3.2 6.8

+

+

+

4.6

-

+ +-

+ + + 60” 0.05 M KP.

0.3 M K P pH 6.8; 0.2 M K P Sigmoid

0.05-0.1M 0.12 M K P pH 6.3; 0 . 2 M KCl Hyperbolic

0.05MKP 0.12 M K P pH 5.7; 0.11 M K P Hyperbolic

From Bernardi and Bernardi (36). As determined by sedimentation in sucrose gradient, using cytochrome c as a reference protein; enzymic assays were done on both bis(p-nitrophenyl) phosphate and the natural substrates; the results were the same. c Using bis(p-nitrophenyl) phosphate in 0.25 M succinate buffer as the substrate. The enzyme degrades polyribonucleotides to 3‘P mononucleosides; these are inhibitory. Eluting molarity; K P is potassium phosphate buffer. 0

results (11) are in general agreement with those of Hilmoe (14), which were obtained on RNA core using lower salt concentrations. The pH-activity curve using bis (p-nitrophenyl) phosphate as a substrate showed an optimum a t pH 5.8, but considerable activity could be detected between pH 6 and 7 (11). Using RNA “core” as the substrate, EDTA and sulfhydryl reagents are activators; MgZ+,MnZ+,and, more effectively, CuZ+,Hg”, and Zn2+ are inhibitors; arsenite and fluoride are weak inhibitors (14 ) . Deoxyribonucleoside-3’-phosphates are competitive inhibitors of the activity on acid DNase digests.

336

A. BERNARD1 AND G. BERNARDI

IV. Distribution and lntracellular localization

It has been claimed that spleen acid exonuclease has its counterpart in other tissues (3, 40). It is very likely that this claim is correct, in spite of the fact that it was based on the wrong assumption that a hydrolytic activity a t pH 6.0 on p-nitrophenyl thymidine-3’-phosphate could be equated with acid exonuclease activity (see Tables I and 11).In fact, an acid exonuclease activity has been shown in fish muscle (41, 42) and, using as a substrate an acid DNase digest, in rat liver ( 4 3 ) . In this latter case, it was checked that the activity on p-nitrophenyl thymidine-3’phosphate was not inhibited by S04z- and that it was lower than on thymidylyl (3’ + 5’) thymidine ( 4 4 ) . As far as the intracellular localization of the acid exonuclease activity is concerned, Razzell (40), using the synthetic substrate, found it in both the mitochondrial-lysosomal fraction and in the supernatant. Van Dyck and Bernardi (46) found that an enzymic activity on acid DNase hydrolyzates could be extracted from rat liver lysosomes and that the sedimentation coefficient of this activity, as determined by sucrose density gradient centrifugation, was the same as for spleen acid exonuclease. Subsequent work (43, 44) confirmed the idea that acid exonuclease is a lysosomal enzyme like other acid hydrolases of nucleic acids [see Table IV and related discussion in Bernardi ( 6 )1. For a recent review on the problem of intracellular localization of exonucleases, the reader is referred to Shugar and Sierakowska (46‘).

40. W. E. Razzell, JBC 236, 3028 (1961). 41. N. Tomlinson, Can. J . Biochem. Physiol. 36, 633 (1958). 42. N. Tomlinson, Can. J . Biochem. Physiol. 47, 945 (1959). 43. J. M. Van Dyck and R. Wattiaux, European J . Biochem. 7 , 15 (1968). 44. M. Erecinska, H. Sierakowska, and D. Shugar, European J . Biochem. 11, 465 (1969). 45. J. M . Van Dyck and G. Bernardi, unpublished experiments (1968). 46, D. Shugar and H. Sierakowska, in “Progress in Nucleic Acid Research” (J. N. Davidson and W. E. Cohn, eds.), Vol. VII, p 369. Academic Press, New York, 1967.

Nucleotid e Phosphomonoesterases GEORGE I. DRUMMOND

MASANOBU YAMAMOTO

I . 5'-Nucleotidase . . . . . . . . . . . . . A . Bacterial 5'-Nucleotidase . . . . . . . . . B . Yeast iY-Nucleotidase . . . . . . . . . . C . Snake Venom 5'-Nucleotidase . . . . . . . . D. Bull Seminal Plasma 5'-Nucleotidase . . . . . . . . . . . . . . . E . Liver 5'-Nucleotidase F. Intestinal 5'-Nucleotidase . . . . . . . . . . . . . . . . G . 5'-Nucleotidase from Pituitary H . 5'-Nucleotidase from Nerve Tissue . . . . . . I . 5'-Nucleotidase from Cardiac Tissue . . . . . . J . 5'-Nucleotidase from Other Vertebrate Tissues . . . K . 5'-Nucleotidase from Ehrlich Ascites Tumor Cells . . . L. 5'-Nucleotidase from Potatoes . . . . . . . M . Comparison of the Enzymes . . . . . . . . 11. 3'-Nucleotida~e . . . . . . . . . . . . . . . . . . . . . A. Rye Grass 3'-Nucleotidase B. M u g Bean 3'-Nucleotidase . . . . . . . . C . 3'-Nucleotidase from Wheat Seedlings . . . . . . D . 3'-Nucleotidase from Microorganisms . . . . . .

337 338 341 342 342 343 345 346 346 347 348 348 349 349 352 353 353 353 354

.

I S'-Nucleotidase

5'-Nucleotidase (5'-ribonucleotide phosphohydrolase. EC 3.1.3.5) is widely distributed in nature and a voluminous literature has appeared in the past decade on the enzyme from vertebrate tissues. seminal fluid. snake venoms. yeasts. and bacteria . Studies regarding the discovery and early investigations of the enzyme have been reviewed by Heppel ( 1 ) and

.

1 . L . A . Heppel. "The Enzymes. " 2nd ed., Vol . 5. p 49. 1961.

337

338

G. I. DRUMMOND AND M. YAMAMOTO

more recently by Bodansky and Schwartz ( 2 ) . The enzymes from these various sources have some properties in common ; however, numerous differences exist with respect to substrates and physical and chemical properties. Because of the diversity of studies each enzyme will be reviewed according to its source; this will be followed by a synopsis of the main similarities and contrasting features. A. BACTERIAL 5'-NUCLEOTIDASE In 1963, Kohn and Reis (3) first drew attention to the fact that extracts from many species of bacteria-Proteus, Hemophilus, Staphylococcus, Escherichia, and Clostridium-were capable of hydrolyzing both ribonucleoside 3'- and 5'-monophosphates. From their studies on relative activities with the two substrates, pH optima, and the effects of metal ions, they concluded that bacterial 3'- and 5'-nucleotidases were distinct and separate enzymes. Since that time both activities have been examined closely. Neu and Heppel ( 4 ) found that the 5'-nucleotidase of E . wltwas released into solution when spheroplasts were prepared with ethylenediaminetetraacetate (EDTA)-1ysozyme (6). The enzyme is also released from E. coli cells by osmotic shock. In this procedure, cells, preferably in the exponential growth phase suspended in hypertonic sucrose, are centrifuged and rapidly dispersed in a medium of low ionic strength (6). A number of degradative enzymes including 5'-nucleotidase, alkaline phosphatase, and cyclic phosphate diesterase are released into solution (7, 8) (see also Chapter 16 by Drummond and Yamamoto, this volume). Using the osmotic shock technique, Neu (9) has achieved a 5000-fold purification of 5'-nucleotidase from E. coli. The enzyme was pure as judged by molecular sieve chromatography, gel electrophoresis, and ultracentrifugation. A molecular weight of 52,000 was determined. The purified preparation hydrolyzes all 5'-ribo- and 5'-deoxyribonucleotides with preference for 5'-AMP. It does not attack 2'-AMP, 3'-AMP, cyclic 2',3'-AMP, or inorganic pyrophosphate. It appears unusual in that in addition to 5'nucleotides, adenosine triphosphate (ATP) , uridine diphosphate glucose, and bis (p-nitrophenyl) phosphate are hydrolyzed. The ratio of activities 2. 3. 4. 5. 6. 7. 8. 9.

0. Bodansky and M. K. Schwartz, Advan. Clin. Chem. 11, 277 (1968). J. Kohn and J. L. Reis, J Bacterial. 86, 713 (1963). H. C. Neu and L. A. Heppel, BBRC 17, 215 (1964). R. Repaske, BBA 30, 225 (1958). L. A. Heppel, Science 156, 1451 (1967). H. C. Neu and L. A. Heppel, JBC 240, 3685 (1965). N. G. Nossal and L. A. Heppel, JBC 241, 3055 (1968). H. C. Neu, JBC 242, 3896 (1967).

15.

339

NUCLEOTIDE PHOSPHOMONOEGTERASES

for these unrelated substrates remained constant during purification to apparent homogeneity; heat inactivation curves with respect to each substrate were parallel. These facts suggested that the various hydrolytic activities were associated with a single protein (9). This would seem to be a rather unusual enzyme since the substrates are quite unrelated and hydrolysis of bis (p-nitrophenyl) phosphate is usually indicative of diesterase activity. The 5’-AMP hydrolytic activity is stimulated 100-fold by 0.5 mM Co2+;it is inhibited by Zn2+and chelating agents. For stimulation of 5’-AMP and ATP hydrolysis, MnZ+can replace Co2+but Coz+is not needed for UDPG hydrolysis. Glaser et al. ( 9 a ) have studied extensively the nucleoside diphosphosugar hydrolase activity of E. coli and have also concluded that this activity and 5’-nucleotidase are associated with the same protein. Thus, the ratio of 5’-nucleotidase to UDP-sugar hydrolase activity remained constant over a 1000-fold range of purification and both were inactivated equally by heat treatment. These investigators showed that 14C-uridine-labeled UDP-D-glucose was hydrolyzed to uridine and inorganic phosphate without extensive mixing with a pool of nonlabeled 5’-UMP. This suggested an enzyme-bound complex of 5‘UMP as intermediate. Based on these findings they concluded that UDPG is hydrolyzed by the following sequence: a-glucose-1-P

UDP-D-glucose

+E

UDP-D-glucose-E

+

E-UMP

-+

It E + 5’-UMP

E

+ uridine + Pi

The enzyme also cleaves UDP-D-galactose and UDP-N-acetyl-D-galactosamine; other nucleosidediphosphate sugars containing adenine, guanine, and cytosine as the base are hydrolyzed at less than 5% of the rate of uridine nucleotides. The authors suggested (9a) that the enzyme is likely concerned with intracellular (rather than extracellular) UDPG, acting to maintain suitable levels of nucleotides in the cell. With such a wide range of specificity, the name 5’-nucleotidase for this bacterial enzyme seems inappropriate. When E. coli were grown in the presence of low concentrations of EDTA, a striking reduction in the activity of 5’-nucleotidase (also alkaline phosphatase and cyclic phosphate diesterase) occurred (10) and the data suggest that EDTA acts by binding a trace metal essential for activity. To investigate this further, E . coli were grown in the presence of s5Zn2+and subjected to osmotic shock (11).5’-Nucleotidase was purified from the shock fluid to apparent homogeneity. The purification was ac9a. L. Glaser, A. Melo, and R. Paul, JBC 242, 1944 (1967). 10. H. F. Dvorak, JBC 243, 2640 (1968). 11. H. F. Dvorak and L. A. Heppel, JBC 243, 2647 (1968).

340


companied by an enrichment with respect to 65Zn which could not be removed by dialysis. This and other evidence suggest that 5’-nucleotidase (and nucleoside cyclic phosphate diesterase) are metalloproteins, possibly zinc metalloproteins. The fact that 5’-nucleotidase is released into solution during spheroplast formation and during osmotic shock suggests a surface localization of the enzyme. Other evidence as to surface localization has also been provided. Thus, when an E . coli mutant (U-7a) which lacks alkaline phosphatase was grown on 2’-AMP as a carbon and phosphate source a severe lag in growth occurred (12). I n the presence of 5’-AMP growth took place rapidly. The inference was that 5‘-nucleotidase, being a surface enzyme, was capable of cleaving 5’-AMP to provide a carbon and phosphorus source for the cell. The possible location of these enzymes between the cell wall and cell membrane in the “periplasmic” space has been considered by Heppel ( 6 ) and is also discussed in Chapter 16 by Drummond and Yamamoto, this volume. Osmotic shock has also been used to release the enzyme from various Enterobacteriaceae: Shigella sonnei, Salmonella heidelberg, and Proteus vulgaris (13, 14). The enzyme from all these organisms exhibit properties similar to the E . coli enzyme with regard to the p H optimum, ion stimulation, substrate specificity, and physical properties. Mauck and Glaser (15) have recently purified a periplasmic enzyme from Bacillus subtilis which catalyzes the hydrolysis of several nucleosidediphosphate sugars ( ADP-glucose, GDP-glucose, GDP-mannose, CDP-glucose, etc.) to the corresponding nucleoside, inorganic phosphate and sugar phosphate. The enzyme shows 5’-nucleotidase activity and both activities seem to be catalyzed by the same protein. Unlike the E . coli enzyme, no divalent cations are required and it does not hydrolyze ATP. A specific protein inhibitor for 5’-nucleotidase has been purified from E. coli cell cytoplasm (10, 16). It prevents the action of the enzyme on 5’-AMP, ATP, and UDPG. It also inhibits the hydrolysis of 5’-AMP by the 5‘-nucleotidases from A . aerogenes, S. sonnei, and S. typhimurium (10). Other Enterobacteriaceae also possess similar intracellular protein inhibitors (13) which inhibit all hydrolytic activities of the 5’-nucleotidase of these organisms. The relevance of this inhibitor protein to the action of the enzyme in vivo is not known. 12. H.C.Neu, JBC 242, 3905 (1967). 13. H.C.Neu, Biochemistry 7, 3766 (1968). 14. H.C.Neu, J . Bacterbl. 95, 1732 (1968). 15. J. Mauck and L. Glaser, BiochemGtry 9, 1140 (1970). 16. H.F.Dvorak, Y. Anraku, and L. A. Heppel, BBRC 24,628 (1966).

15.

NUCLWTIDE

PHOSPHOMONOESTERASES

341

B. YEAST 5’-NUCLEOTIDASE A 5’-nucleotidase from yeast (Saccharomyces oviformis) has been purified to electrophoretic homogeneity and studied kinetically in detail by Takei (17-21). The enzyme hydrolyzes all ribo- and deoxyribonucleoside 5’-phosphates. It does not hydrolyze nicotinamide mononucleotide, 3‘- or 2’-AMP, sugar phosphates, or P-glycerol phosphate. Like the bacterial enzyme, yeast 5’-nucleotidase is markedly activated by Co2+and Ni2+ (18) and inhibited by EDTA. It was found that purified preparations of the enzyme possessed nucleotide pyrophosphatase (EC 3.6.1.9) activity. Substrates for the nucleotide pyrophosphatase are NAD, NADH2, FAD, ATP, and to a lesser degree, NADP and inorganic pyrophosphate (21). This activity could not be eliminated during purification of the 5’-nucleotidase to electrophoretic homogeneity (19). The pH profile of both activities were the same (pH 6.3-6.5) ; both activities were equally labile on heating a t temperatures between 40” and 60” for 5 min; their stabilities to UV irradiation and urea treatments were identical ; and both were similarly inhibited by N-bromosuccinimide, iodine, Zn2+, Ag+, and Cu2+ ( 2 1 ) . These data indicate that both 5’-nucleotidase and nucleotide pyrophosphatase reside in the same enzyme protein in S . oviformis. The enzyme seems to catalyze the catabolism of NAD in this organism as follows:

+ + adenosine + Pi + NhlN

(1) NAD -+ 5’-AMP NMS (2) 5’-AMP -+ adenosine I’,

(3) NAD

-+

The K , values for 5’-nucleotides are in the range of 0.2 mM for purine ribonucleotides and higher (2 mM range) for pyrimidine ribonucleotides (20). The enzyme is inhibited competitively by purine and pyrimidine bases, nucleosides, 2’- and 3’-mononucleotides, and NMN; NAD and NADP display a mixed type of inhibition against 5’-AMP hydrolysis ( K i values of 1 and 7 mM, respectively). Takei has concluded that the active sites for the two activities although residing in the same protein are not identical. It seems possible that the enzyme may be composed of two protein subunits each with a separate activity and with active centers 17. 9. Takei, Agr. Bwl. Chem. ( T o k ~ o 29, ) 372 (1965). 18. S.Takei, Agr. Bwl. Chem. ( T o k y o ) 30, 1215 (1966). 19. S.Takei, Agr. Biol. Chem. ( T o k y o ) 31, 917 (1967). 20. S.Takei, Agr. Bwl. Chem. (Tokyo) 31, 1251 (1967). 21. S. Takei, J. Totsu, and K . Nakanishi, Agr. BWZ. Chem. (Tokyo) 33, 1251 (1969).

342


in part common to each other. In general, this enzyme seems remarkably similar to the bacterial 5’-nucleotidase.

c. SNAKE VENOM

5’-NUCLEOTIDASE

That various snake venoms contain potent 5’-nucleotidase activity has been known for over 30 years. Until fairly recently only relatively crude preparations have been available ( 2 2 ) .Sulkowski et al. have purified the enzyme 1000-fold from Bothrops atroz venom ( 2 3 ) .The preparation was free of alkaline phosphatase and phosphodiesterase activities which are rich in venom of several species (24). The enzyme hydrolyzes all riboand deoxyribo-5’-nucleotideswith greatest reactivity for 5’-AMP. It does not attack 3’-nucleotides, ATP, ribose-5-phosphate1 inorganic pyrophosphate, p-nitrophenyl phosphate, nor dinucleotides of the type d-pXpY or pXpYp ( 2 3 ) .Specificity studies indicate a requirement that C-1 of ribose must contain a nitrogenous base and that the hydroxyl group on C-3 must be free. The enzyme is strongly inhibited by EDTA (0.1-1 mM) and this inhibition is reversed by CW+, Mg2+,and Ni2+.A similar enzyme has been purified from venoms of Crotalus adamanteus, Hemachatus haemachates, and Vipera m s e l l i (26, 2 6 ) . The H . haemachates enzyme is activated by 2’- and 3‘-mononucleotides and by 0-amino acids (26) which seem to do so by increasing the dissociation of enzyme and product. This enzyme is activated by Co2+and Mg2+,and activation is increased in the presence of 3’-AMP (26). 5‘-Nucleotidase has been purified to electrophoretic homogeneity from cobra (Naja naja atra) venom ($7). Properties of this enzyme are again similar to the B . atroz venom enzyme with regard to substrate specificity, activation by Mg2+ and Mn2+, and inhibition by Zn2+and Ni2+.It differs in that the pH optimum is 6.5-7.0.

D. BULL SEMINALPLASMA B’-NUCLEOTIDASE Some of the properties of the enzyme from this source were reviewed by Heppel ( 1 ) . More recently, attention has been focused on certain kinetic properties of the enzyme, especially its double pH optimum. Using 22. 23. 24. 25. 26. 27.

W. Bjork, BBA 49, 195 (1961). E. Sulkowski, W. Bjork, and M. Laskowski, JBC 238,2477 (1903). G. M. Richards, G. du Vair, and M. Laskowski, Sr.,Biochemiatry 4, 501 (1965). W. Bjork, BBA 89, 483 (1964). W. Bjork, Arkiv Kemi 27, 555 (1967). Y. Chen and T. Lo, J . Chinese Chem. SOC.(Taiwan) 15,M (1988).

15.

NUCLEOTIDE PHOSPHOMONOEST~ASES

343

a partially purified enzyme, Bodansky and Schwartz (68)found that in the presence of MgZ+,L-histidine inhibited the enzyme below pH 7.5 but activated i t above this pH value, shifting the optimum from 7.5 to 9.3. In the absence of MgZ+,L-histidine produced inhibition below p H 9. The second pH optimum was independent of buffer, but was Mg2+ and temperature dependent (29). It was also dependent upon the nature of the substrate since the phenomena was exhibited only with 5’-AMP, 5’-GMP, and 5’-IMP. From these studies, Levin and Bodansky (29) have proposed a model to explain the role of Mg2+ in producing a second optimum a t pH 9. The model involves four binding sites: one for the C-2 hydroxyl of ribose, one for water, another for phosphate, and one for Mg2+.The model proposed is contingent upon the absence of isozymes, and none was detected by starch gel electrophoresis. Pilcher and Scott (SO), however, have resolved bull seminal plasma 5’-nucleotidase into three active components by electrophoresis on polyacrylamide gels. It is thus possible that the double pH optimum is a reflection of heterogeneity.

E. LIVER5’-NUCLEOTIDASE 5’-Nucleotidases have been studied in liver from various species and activity has been identified in lysosomes, cytoplasmic supernatants and plasma membrane preparations. Arsenis and Touster (31) have purified a 5’-nucleotidase from rat liver lysosomes to apparent homogeneity. The enzyme is unusual in that it hydrolyzes 2’-, 3’-, and 5’-mononucleotides equally well with preference for 5‘-dAMP. It also hydrolyzes FMN, p nitrophenyl phosphate, and ,&glycerol phosphate, but not inorganic pyrophosphate or bis (p-nitrophenyl) phosphate. Unlike the 5’-nucleotidases described thus far, divalent cations such as Co2+,Mn2+,and Mg2+have no activating effect, but EDTA is inhibitory. In spite of the broad substrate specificity kinetic experiments indicate that a single enzyme is involved. Because of its broad substrate specificity it has been suggested (31) that it may play a key role in lysosomal catabolism of nucleic acids. An apparently different 5’-nucleotidase has been partially purified (50-fold) from acetone powder preparations of chicken liver (32): 5’-IMP and 5’-GMP are hydrolyzed by this preparation more rapidly than other 5‘-nucleotides ; 5‘-AMP, 5’-UMP, and 5’-CMP are hydro28. 0. Bodansky and M. K. Schwartz, JBC 238, 3420 (1963). 29. S. J. Levin and 0. Bodansky, JBC 241, 51 (1966). 30. C. W. Pilcher and T. G. Scott, BJ 104, 41c (1967). 31. C. Arsenis and 0. Touster, JBC 243, 5702 (1968). 32. R. Itoh, A. Mitsui, and K. Tsushima, BBA 146, 151 (1967).

344


lyzed a t rates only 510% that of 5‘-IMP. The enzyme is inactive in the absence of divalent metal and is maximally active in the presence of 10 mM Mgz+or Co2+;MnZ+is less effective. It is competitively inhibited by nucleosides, inosine being the most potent. A similar enzyme preparation has been obtained from acetone powder preparations of rat, frog, and pig liver (33). Again 5’-IMP and 5’-GMP are the preferred substrates; the enzyme requires divalent cations and is inhibited by nucleosides. From a study of the dephosphorylation of pyrimidine nucleotides in the soluble fraction of rat liver, Fritzon (34, 35) has provided evidence that two 5’nucleotidases exist. One of the enzymes had a broader specificity for 5’-nucleotides than the other, which acted mainly on dTMP and dUMP. The former enzyme was partially purified from the 100,000 x g supernatant fraction of rat liver (36) essentially free of nonspecific phosphatases. This enzyme is activated by divalent metals, the pH optimum is 6.3 and 5’-IMP is the preferred substrate. The general properties indicate that it is identical to the one isolated from liver acetone powder preparations by Itoh et al. (33).It is entirely reasonable that the cytoplasmic enzyme alone would be extracted into solution from acetone powders. I n spite of the presence of lysosomal and cytoplasmic 5’-nucleotidases in liver, much evidence exists that most of the enzyme in this tissue is membrane bound. Using procedures for isolating subcellular structural components, Song and Bodansky have reported (37) that activity resides in membrane fragments that constitute a part of the microsomal membranes. The distribution of the enzyme between the subfractions of microsomal preparations subjected t o density gradient centrifugation suggested that most of the activity was in the heavier fraction, i.e., in those membranes with attached ribosomes (38) and therefore deriving from the endoplasmic reticulum rather than the plasma membrane. However, liver plasma membrane preparations isolated by sucrose density gradient procedures (39, .lo) show enrichment with respect to 5’-nucleotidase (41-43), and a recent modification employing CaC12 as a mem33. R. Itoh, A. Mitsui, m d K. Tsushima, J . Biochem. (Tokyo) 63, 165 (1968). 34. P. Fritzon, European J . Biochem. 1, 12 (1967). 35. P. Fritzon, BBA 151, 716 (1968). 36. P. Fritzon, BBA 178, 534 (1969). 37. C. S.Song and 0. Bodansky, JBC 242, 694 (1967). 38. C. S. Song and A. Kappas, Ann. N . Y. Acad. Sci. 166, 585 (1969). 39. P. Emmelot, C. J. Bos, E. L. Benedetti, and P. H. Rumke, BBA SO, 126 (1964). 40. D. M. Neville, Jr., BBA 154, 540 (1968). 41. R. Coleman and J. B. Finean, BBA 125, 197 (1966). 42. J. M. Graham, J. A. Higgins, and C. Green, BBA 150, 303 (1968). 43. P. Emmelot and C. J. Bos, BBA 120, 369 (1966).

15.

NUCLEOTIDE PHOSPHOMONOESTERASES

345

brane stabilizer gives especially high specific activities with respect to 5’-nucleotidase (44). The membrane-bound enzyme from both rat liver and human liver (45) has been solubilized using deoxycholate (46) and sonic oscillation ( 4 7 ) . On precipitation to remove deoxycholate, the enzyme reassociates with phospholipids and other membrane proteins to regenerate vesicular membranes ( 4 6 ) . Widnell and Unkeless (48) have obtained a highly purified 5‘-nucleotidase from rat liver microsomes and plasma membranes using classic fractionation procedures in the presence of detergent. The enzyme has been shown to be a lipoprotein containing only one phospholipid, sphingomyelin. The enzyme hydrolyzes 5’-AMP and 5’-UMP more rapidly than other 5’-nucleotides. The substrate specificity differs considerably from that of the enzyme isolated from acetone powder extracts (33) and is probably identical with the one identified in plasma membrane preparations by Song and Bodansky (37). Thus, there is likely as many as three enzymes with 5’-nucleotidase activity in liver, one lysosomal, one cytoplasmic, and one membrane bound. Their specificities and kinetic properties appear to be distinctly different. This would suggest specialized physiological functions not yet understood.

F.

INTESTINAL

5’-NUCLEOTIDASE

Center and Behal (49) have resolved 5’-nucleotidase from calf intestinal mucosa into three fractions using DEAE-cellulose chromatography. One of these was obtained free of nonspecific phosphatase. It had a pH optimum of 6-6.5, MnZ+,Mg2+,and Co’+ (1-10 mM) all enhanced activity and complete inactivation was produced with 1 mM EDTA. This enzyme hydrolyzes all 5’-ribonucleotides a t similar rates and hydrolyzes 5’deoxribonucleotides more slowly. These properties indicate that it is strikingly similar to the one obtained from acetone powder preparations of chicken and rat liver (32, 33) and from soluble supernatants of rat liver ( 3 6 ) .The other two activities (which were not fully characterized) (49) could possibly have originated from particulate material or membranes because the authors employed deoxycholate in the early phase of purification. T. K. Ray, BBA 196, 1 (1970). C. S.Song and 0. Bodansky, BJ 101, 5 C (1966). C. S. Song, B. Tandler, and 0. Bodansky, Biochem. Med. 1, 100 (1967). C. S. Song, J. 8. Nisselbaum, B. Tandler, and 0. Bodansky, BBA 150, 300 (1968). 48. C. C. Widnell and J. C. Unkeless, Proc. Natl. Acad. Sci. U.S. 61, 1050 (1968). 49. M. S.Center and F. J. Behal, ABB 114, 414 (1966). 44. 45. 46. 47.

346 G. 6’-NUCLEOTIDASE


FROM PITUITARY

Some of the kinetic properties of a partially purified (60-fold) 5’nucleotidase from bovine pituitary gland have been described (50-54). The specificity of this enzyme seems different from that of other tissue in that 5’-GMP and 5’-UMP are the preferred substrates (61, 5 4 ) . The enzyme is strongly inhibited by EDTA and this is reversed by Mgz+ but not MnZ+ ( 5 0 ) .It is inhibited by Zn2+and competitively inhibited by 2’- and 3’-mononucleotides and nucleosides, particularly adenosine ( 5 4 ) .The approximate molecular weight was determined to be 237,000. It cannot be determined for certain whether this is a cytoplasmic enzyme. Pituitary glands were homogenized in 0.1 M ammonium sulfate and centrifuged at low gravitational force so that membranous material could have been present in the early stages of purificatian.

H. 5’-NUCLMYTIDASE

FROM

NERVE TISSUE

The enzyme has been partially purified (70-fold) from 38,000 X g supernatant fluid from sheep brain homogenates by Ipata (55-58).The, enzyme (MW 140,000) is reported to be specific for 5’-AMP and 5’-IMP although the substrate specificity does not appear to have been examined closely. 2‘- and 3’-AMP are not hydrolyzed ( 5 6 ) . Unlike the enzyme from many sources the brain enzyme does not require divalent cations and indeed Co2+, which stimulates several other 5’-nucleotidases, was inhibitory a t 5 mM. The enzyme is strongly inhibited by very low concentrations of ATP, UTP, and CTP (50% inhibition by 0.3 & ATP) but not by GTP. 2’-AMP, 3’-AMP, and a variety of other nucleoside monophosphates, nucleosides, and sugar phosphates do not inhibit. A kinetic examination of ATP, UTP, and CTP inhibition (56-58) revealed that inhibition curves were sigmoidal, indicating cooperativity between inhibitor molecules and an allosteric type of interaction between inhibitor and protein. The metabolic significance of ATP inhibition is 50. J. Lisowski, Arch. Immunol. Therap. Exptl. 12, 542 (1964). 51. J. Lisowski, Arch. Immunol. Therap. Ezptl. 14, 195 (1966). 52. J. Lisowski, Arch. Immunol. Therap. Exptl. 14, 209 (1966). 53. J. Lisowski, Arch. Irnmunol. Therap. Exptl. 14, 217 (1966). 54. J. Lisowski, BBA 113, 321 (1966). 55. P. L.Ipata, Nature 214, 618 (1967). 56. P. L. Ipata, BBRC 27, 337 (1967). 57. P. L. Ipata, Biochemistry 7, 507 (1968). 58. P. L. Ipata, Frog. Brain Res. 29, 527 (1968).

15.


347

not established. Again it is not entirely clear whether this enzyme is cytoplasmic, being recovered from homogenates centrifuged a t 38,000 X g. 5’-Nucleotidase activity has been examined histochemically (6941) in brain tissue and such studies indicate that more than one 5‘-nucleotidase is present or that the enzyme exists as isozymes (62). It is certainly possible that multiple activities exist in a tissue such as brain with such a diversity of cell types. Activity is present in both white and gray matter (63, 64) and is greatest in cortical tissue, cerebellum, and spinal cord (66). 5‘-Nucleotidase activity has been reported to be greatly diminished in demyelinated areas of the brain in patients with multiple sclerosis (66).

I. 5’-NUCLEOTIDASE FROM CARDIAC TISSUE The enzyme in the myocardium has recently attracted attention because of the possibility that adenosine is a physiological regulator of coronary blood flow (67) (adenosine is a potent coronary dilator). Most of the 5’-nucleotidase activity in rat heart is membrane bound, and a partially purified preparation has been obtained by extracting acetone powder preparations with deoxycholate (68). All 5’-nucleotides are hydrolyzed. The enzyme is strongly inhibited competitively by ATP (Ki 1.8 &). Whether this provides a regulatory mechanism for adenosine formation in the heart is not known. Histochemical evidence has indicated that the enzyme resides in the sarcoplasmic reticulum and transverse tubular system of rat myocardium (69). However, other evidence indicates that enzymic activity is also localized within the walls of the coronary blood vessels (70-72). Recent histochemical studies (73) have shown that the enzyme resides not 59. T. G. Scott, J . Comp. Neurol. 122, 1 (1964). 60. T. G. Scott, J . Comp. Neuro2. 129, 97 (1967). 61. D. Naidoo, J . Histochem. Cytochem. 10, 421 (1962). 62. T. G. Scott, J . Histochem. Cytochem. 13,657 (1965). 63. H. B. Tewari and G. H. Bourne, J . Anat. 97, 65 (1963). 64. K. Nandy and G. H. Bourne, Arch. Neurol. 11, 547 (1964). 65. N. Robinson and B. M. Phillips, Clin. Chim. Acta 10, 414 (1964). 66. K. D. Barron and J. Bernsohn, Ann. ‘N. Y . Acad. Sci. 122, 369 (1965). 67. R. M. Berne, Am. J. Physwl. 204, 317 (1963). 68. H. P. Baer, G. I. Drummond, and L. Duncan, Mol. Pharmacol. 2, 67 (1966). 69. J. Rostgaard and 0. Behnke, J . Ultrastruc. Res. 12, 579 (1965). 70. J. R. Williamson and D. L. Dipietro, BJ 95, 226 (1965). 71. H. P. Baer and G. I. Drummond, Proc. Soc. Ezptl. Bwl. Med. 127, 33 (1968). 72. E. Bajusz and G. Jasmin, Acta Histochem. 18, 222 (1964). 73. K. Nakatsu, H. Clarke, and G. I. Drummond, Federation Proc. 29, 351 (1970).

348


within the cardiac cell but exists almost exclusively in the capillary endothelial cells and small blood vessels of the coronary vasculature. Location of the enzyme a t these sites could have considerable significance with regard to the role of adenosine as an autoregulator of blood flow to the heart. .J. 5’-NUCLEOTIDASE FROM OTHERVERTEBRATE TISSUES 5’-Nucleotidase present in 48,000 x g supernatant fractions of rat and guinea pig skeletal muscle extracts has been examined briefly (74).5‘UMP seems to be the preferred substrate. The enzyme from fish skeletal muscle has also been studied (75).This enzyme hydrolyzes all riboand deoxyribonucleoside 5’-phosphates (except dCMP and dTMP) with preference for 5’-IMP and 5’-UMP. The enzyme is strongly activated by Mn2+; Mg2+is a less powerful activator, and Zn2+ and EDTA are inhibitors. This enzyme thus appears similar to the soluble activity from mammalian liver (33,36). 5’-Nucleotidase in mammary gland hydrolyzes all 5’-ribonucleotides and shows a decrease from pregnancy to early lactation (76).Rats injected with glucagon show increased 5‘nucleotidase in pancreatic islet tissue (77).The enzyme in mouse kidney has been examined histochemically and electrophoretically and found to exist as isoaymes (78).Electrophoretic techniques have also provided evidence that the enzyme exists as isozymes in many other tissues of the mouse such as liver, spleen, intestine, testes, and heart (79).

K. 5‘-NUCLEOTIDASE FROM EHRLICH ASCITES TUMOR CELLS Murray and Friedrichs (80) have obtained a 5‘-nucleotidase from a particulate fraction of Ehrlich ascites tumor cells using deoxycholate. The relative rates of hydrolysis of 5’-UMP, 5’-AMP, 5’-CMP, 5’-GMP, and 5‘-IMP are 129, 100, 93, 83, and 79, respectively. Adenosine and thymidine triphosphate are competitive inhibitors of 5’-AMP hydrolysis 74. I. Cozzani, P. L. Ipata, and M.Ranieri, FEBS Letters 2, 189 (1969). 75. H. L. A. Tarr, L. J. Gardner, and P. Ingram, J. Food Sci. 34, 637 (1969). 76. D. Y. Wang, BJ 83, 633 (1962). 77. S. Johanason and I. B. Toljedal, Endocrinology 82, 173 (1968). 78. M. J. Hardonk and 6. Koudstaal. Hitochemie 15, 290 (1968). 79. M. J. Hardonk and H. G. A. de Boer, Histochemie 12,!29 (1968). 80. A. W. Murray and B. Friedrichs, BJ 111, 83 (1989).

15.

NUCLEOTIDE PHOSPHOMONOESTERASEE

349

a,

(Ki 0.4 and 4.8 respectively). The enzyme is strongly inhibited by Znz+.5’-Nucleotidase in these cells has also been examined by Paterson and Hori (81) who found the enzyme located primarily in nuclei. Nuclei prepared from a 6-mercaptopurine-resistant subline were markedly deficient in enzymic activity. L. 5’-NUCLEOTIDASE

FROM POTATOES

The presence of an enzyme with 5’-nucleotidase activity in extracts of potato were referred to by Heppel (1).The enzyme has been purified 200-fold by Klein (8B) and studied kinetically (83). All major 5’nucleotides are hydrolyzed a t similar rates. The preparation also hydrolyzed 3’-nucleotides a t a substantial rate (2040% that of 5’-AMP). However, kinetic data (83)suggested that the purified preparation was perhaps a mixture of specific 5’- and 3’-nucleotidases.

M. COMPARISON OF

THE

ENZYMES

It seems clear that the 5‘-nucleotidases are a somewhat heterogeneous group of enzymes with many differences and yet having certain properties in common. It must be emphasized that most of the enzyme preparations do not represent pure proteins, and there can be many variations in experimental procedures which might account for some of the differences. Table I presents a summary of some of the similarities and contrasting features of the enzyme from several sources. It can be seen that the relative rates of hydrolysis of the major 5’-nucleotides differ in a seemingly random manner. The E . coli and S. sonnei enzymes differ from those of other sources in that they possess nucleoside diphosphosugar hydrolase activity and also hydrolyze ATP. The B. subtilis enzyme is different from that of the two above organisms in that it does not hydrolyze ATP and converts UDPG to nucleoside and sugar phosphate (16) whereas the E . coli and S. sonnei enzymes degrade this compound to base, sugar, and inorganic phosphate (9,13). The yeast enzyme is unique in that it possesses nucleotide pyrophosphatase activity, converting NAD to NMN, adenosine and inorganic phosphate. The liver lysosomal enzyme appears to have yet a different substrate profile, hydrolyzing both nucleoside 2‘- and 3’-phosphates in addition to 5‘-phosphates. 81. A. R. P. Paterson and A. Hori, Can. J . Biochem. Physwl. 41, 1339 (1963). 82. W. Klein, 2. Physwl. Chem. 307, 247 (1957). 83. W. Klein, 2. Physwl. Chem. 307, 254 (1957).

W

cn

0

TABLE I SUMMARY OF PROPERTIES OF VARIOUS 5!-NUCLEOTIDASES

Apparent Enzyme murce

E. wli

Cellular localisstion

K, Other eubstrah

5'-AMP (mM)

PH optimum

Activators

Deoxyribo0.03(0.12 6.0for5'Cd+.Mn'+, nucleotidea for ATP) AMP; 6.8 Ca*+ ATP. UTP, for ATP GTP. UDPglucose E . sonnei Surface A > U > G > C > I Deoxyribo0.012 (0.13 5.8 for 5'Cd+. Mn*+ periphic nucleotidea for ATP) AMP: 7-8 for UDPG ATP, ADP. (0.11 for UDP-glucoee UDPG) 8 UDPG. CDPG, 0.0018 for B. uubtilis A >G >U >C ADPG UDPG 5.57 C d + , Ni'+ A>>G = I > U > C NAD. NADH, 0.2 Yesst FAD, ATP S.wiformis

Snake venom B. droz

Surface periplaamic

Relative rate of hydrolyaia of 5'ribonucleotideaa A > G > C > U > I

A > C > I > G > U

Deoxyribonucleotidea NMN

9

W+.Mg'+ and Ni'+ reveme EDTA inhibition

Inhibitor8

MW

EDTA citrate 52,000

D%grS?% of purity

Ref.

5000-fold ( 9 )

urea

EDTA citrate 44,000-

3000-fold

(13)

p

(16)

P

(17$1 )

1OOO-fold

(83)

53 ,000

137,000

Zn*+, Cu'+. EDTA nucleosidea. particularly adenoeine En'+, EDTA

Rat liver

Rat liver

Rat liver Calf intestinal mucw Bovine pituitary

Lyemornea

>U

=G

>I

I =G >U >A I00.OOO X u supernatant U = A > C > G > I " P b membrane" U >C =G >A

Membrane bound

F i h skeletal muscle Ascitea Nuclear tumor cells membrane a

A

6.5-7 7 . 8 and 9 . 2

C > A > I > G

Sheep brain

Rat heart

0.13

A > C = U > C

Snake venom N. naja olra BuU seminal P b

2'-, 3'-Mononucleotides. 5'-deo4nucleotidea CDeoxyribonucleotidea

3.7-5.5

Mn'+. Mg'+

NP+, Znz+

Cd+.Ni'+

Znt+. Cu'+ nucleonidea nucleotidea Zn*+. NP+

Cd+,Mn*+.

6.8

6.3

Nucle&dea

P P

Cd+. MIL¶+, ME¶+

EDTA nucleosidea

G > U = C > I >A

0.06

9

ME'+ r e v e m EDTA

EDTA Zn'+ nucleonidea

A = I

0.007

7.3

C > U = A > I >G

0.018

9

U > A > C > G > I

P

P

6.04.5

>C >G

P

ME'+

0.05

U =I

Hihly

and Niz+ do n d activate

0.012 5'-Deoryribcnuoleotiden

1O.OOO

8

5'-Deoxyrib* nucleotidea 0.067

inhibition None required

Mnt+, ME'+

237 ,OOO

ATP. UTP. Cot+. En:+ ATP (Ri 1.8 r M ) EDTA. ZnZ+ ATP, Zn'+

P

P

D P

120,OOO

Nuclec&de-5'-monophospha~are represented only by the baee letter. Degree of purity designated nu p representa partial purification.

P

352

Q. I. DRUMMOND AND M. YAMAMOTO

The most notable similarities relate to activation and inactivation by metal ions and other materials. I n most instances (Table I) each is activated by one or more of the cations Co2+,Mn2+,NiZ+,Mg2+,or Ca2+ of which the former three are usually most effective. Here again, however, there are differences. Thus, the B. atrox enzyme is not activated by ions, but they serve to reverse EDTA inhibition (23). The rat liver lysosomal enzyme is also not activated by divalent metals (31) ; the sheep brain enzyme does not seem to require divalent cation and in fact it is inhibited by Co2+ (67). In most cases EDTA has been reported to be an effective inhibitor; Znz+and Cu2+are frequently inhibitory. Whereas ATP is a substrate for the bacterial enzyme, it and other nucleosidetriphosphates are powerful inhibitors of the enzyme from several mammalian sources (a7, 67,68). The enzymes differ markedly in molecular weight, varying from 10,OOO for the snake venom (Naja atra) enzyme (27) to 237,000 for the enzyme from bovine pituitary (64). It appears certain that there is more than one 5‘-nucleotidase present in most mammalian tissues. This is best established for liver. In other cases it has not been possible to determine the exact intracellular origin because of the nonselective extraction procedures used. However, those enzymes isolated from acetone powder preparations of chicken liver and rat liver appear to have properties essentially identical to the enzyme present in 100,000 x g supernatant fraction of rat liver and therefore may be cytoplasmic in origin. This could also be the case for the intestinal mucosa enzyme. Very little can be said about the physiological function of the enzyme except that it is obviously involved in normal cellular catabolism of nucleosidemonophosphates. Its surface localization in microorganisms must have metabolic relevance; its presence in membrane structures in mammalian tissues also points to specialized functions. Perhaps, even the nucleoside product has physiological functions yet to be discovered.

II. 3’-Nucleotidase

In contrast to 5‘-nucleotidases, enzymes which hydrolyze nucleoside3‘-phosphates have attracted comparatively little attention. Enzymes possessing some specificity for 3’-nucleotides seem to occur predominantly in the plant kingdom and several have been only partially purified and characterized.

15.


353

A. RYE GRASS3’-NUCLEOTIDASE An enzyme purified from rye grass capable of specifically hydrolyzing 3‘-nucleotides has been available for some years (84). The enzyme has also been purified (50-fold) from germinating rye seedlings (85) and seems quite specific for 3’-nucleotides. 3’-Deoxymononucleotides are not attacked (86) nor is arabinonucleoside 3’, 5’-diphosphate (87) . Enzymic activity increases 10-fold during germination (85).

B. MUNGBEAN3’-NUCLEOTIDASE Walters and Loring (88) have purified a 3’-nucleotidase about 50-fold from mung bean sprouts (Phaseolus aureus Roxb.). The enzyme hydrolyzes 3’-AMP, 3’-GMP, 3’-CMP in decreasing order and also hydrolyzes the 3I-phosphate group of coenzyme A. (89), but it has no significant activity for 2’- or 5’-ribonucleotides. For 3’-GMP, 3’-AMP, 3’-UMP, and 3’-CMP, K , values are 0.67, 1.1, 7.7, and 15 mM, respectively. The enzyme preparation also contained acid stable ribonuclease activity (89). Both 3’-nucleotidase and acid ribonuclease were inactivated reversibly at pH 5.0 and by dialysis and this inactivation could be prevented by Zn2+.The two activities were similarly inactivated by heat at pH 5 and 7.5. Such data indicate that the two are metalloproteinsprobably zinc metalloproteins. These similarities and other kinetic data provide evidence that the 3’-nucleotidase and ribonuclease activities reside in the same protein.

c. 3’-NUCLEOTIDASE FROM

WHEAT

SEEDLENGS

An enzyme similar to the 3‘-nucleotidase of mung bean has been isolated from germinating wheat seedlings and purified 800-fold (90). The preparation possessed DNase, RNase, and 3’-nucleotidase activities. These three activities were similar in p H optima, requirements for Znz+ and sulfhydryl compounds, stability to storage, temperature inactivation L. Shuster and N. 0. Kaplan, “Methods in Enzymology,” Vol. 2, p. 551, 1955. L. Shuster, JBC 229, 289 (1967). L. Cunningham, JACS 80, 2546 (1958). G. R. Barker and G. Lund, BBA 55, 987 (1962). T. L. Walters and H. S. Loring, JBC 241, 2870 (1966). H. S. Loring, J. E. McLennan, and T. L. Walters, JBC 241, 2876 (1966). 90. D. M. Hanson and F. L. Fairley, JBC 244, 2440 (1969).

84. 85. 86. 87. 88. 89.

354


and reactivation and inhibition by metal ions and EDTA. The three activities also co-chromatographed on DEAE-cellulose and phosphocellulose and migrated identically on gel filtration. The three activities thus seem to reside in a single protein. The activity in wheat seedlings increases over 80-fold during germination (91). D. 3’-NUCLEOTIDASE

FROM

MICROORGANISMS

Numerous microorganisms possess a cyclic 2’, 3’-ribonucleoside phosphate diesterase which has 3‘-nucleotidase activity. This “double headed” enzyme has been vigorously studied and is described in the chapter on nucleoside cyclic phosphate diesterases (Chapter 16 by Drummond and Yamamoto, this volume). Suffice it to say that the ability of microorganisms producing this enzyme to catabolize 3’-nucleotides is well established. An enzyme from Bacillus subtilis, which hydrolyzes a variety of nucleoside 3’-phosphates, has been briefly described (92). The enzyme was found to be present in culture filtrates after removing the cells by centrifugation. Whether or not this enzyme is identical to the nucleoside cyclic phosphate diesterase in this organism is unclear. Because of its specificity for 3‘-nucleotides it has been proposed (93) as a specific method for preparing 2’-nucleotides. Becker and Hurwitz (94) have found that after infection of E . coli B with T-even bacteriophages a novel 3‘-deoxynucleotidase activity appears. They purified the enzyme 2000-fold. In addition to its attack on 3’-deoxymononucleotides, the enzyme selectively removes the 3’-phosphoryl groups from DNA. It does not attack 3‘-ribonucleotides, 3’phosphoryl groups of RNA, or 5’-phosphate esters. Like bacterial 5‘nucleotidases, this enzyme is markedly activated by Mg2+and Co2+and is inhibited by EDTA. The enzyme appears to be a phage-induced enzyme; the activity rises early after injection with T-even phages and formation of the enzyme is blocked with chloramphenicol.

91. L. Shuster and R. H. Gifford, Arch. Biochem. Biophys. 96, 534 (1962). 92. S. Igarashi and A. Kakinuma, Agr. Biol. Chem. ( T o k y o ) 26, 218 (1962). 93. A. Kakinuma and S. Igarashi, Agr. Biol. Chem. ( T o k y o ) 28, 131 (1964). 94. A. Becker and J. Hurwitz, JBC 242, 936 (1967).

Nucleoside Cyclic Phosphate Diesterases GEORGE I . DRUMMOND

MASANOBU YAMAMOTO

I . Introduction . . . . . . . . . . . . . I1. Ribonucleoside Y.3'-Cyclic Phosphate Diesterase with 3'-Nucleotidase Activity from Microorganisms . . . . . A . Properties of the Enzyme . . . . . . . . B . Cellular Localization . . . . . . . . . . C . Cyclic Phosphate Diesterase-A Metalloenzyme . . . D . Physiological Function . . . . . . . . . I11. Ribonucleoside Y.3 '-Cyclic Phosphate Diesterase from Vertebrate Nerve . . . . . . . . . . . . A . Properties and Substrate Specificity . . . . . . B . Intracellular Localization in Myelin . . . . . . C . Physiological Role of the Y.3 '-Cyclic Phosphate Diesterase IV . Nucleoside 3'.5 '-Cyclic Phosphate Diesterase . . . . . . A. Distribution of the Enzyme in Nature . . . . . B . Substrate Specificity . . . . . . . . . . C . Intracellular Localization . . . . . . . . . D . Metal Ion Requirement. pH Optimum and Substrate Affinity . . . . . . . . . . . . . E . Inhibitors and Activators . . . . . . . . . F. Possibility of Other Nucleoside Cyclic 3'.5 '-Phosphate Diesterases . . . . . . . . . . . . G . Physiological Function of Nucleoside 3'.5 '-Cyclic Phosphate Diesterase . . . . . . . . .

355 356 357 361 362 363 363 364 364 365 365 366 366 367 368 368

370 370

.

I Introduction

Enzymes are available from a variety of sources which split the phosphodiester bond of nucleoside 2'.3 '. and 3'.5 '.cyclic phosphates . The ability of the ribonucleases to hydrolyze ribonucleoside 2'.3 '.cyclic phosphates to the corresponding 3'-phosphates is well known . During 355

356


the past decade a number of enzymes have been discovered which appear to be specific for nucleoside cyclic phosphates and which do not possess nuclease action, i.e., they do not split internucleotide bonds. Early studies in this area were reviewed in the second edition of “The Enzymes” by Khorana ( 1 ) . Since that time three enzymes in particular have been reasonably well characterized, the cyclic phosphate diesterase from bacteria which possesses 3‘-nucleotidase activity, the nucleoside 2‘,3’cyclic phosphate diesterase from nerve tissue, and a nucleoside 3’, 5’-cyclic phosphate diesterase which is widely distributed in nature.

II. Ribonucleoride 2’,3’-Cyclic Phosphate Diesterare with 3’-Nucleotidase Activity from Microorganisms

In 1964, Anraku (2, 3) reported the isolation of an enzyme from Escherichia coli B which hydrolyzed ribonucleoside 2’,3’-cyclic phosphates. Enzyme fractions representing a 900-fold purification also possessed 3’-nucleotidase activity. Similar activities have subsequently been purified from Proteus mirabilis (4, 6),halophilic Vibrio alginolyticus (6, 7) , Bacillus subtilis (8),and various Enterobacteriaceae, specifically, Shigella sonnei, Salmonella heidelberg, Serratia marcescens, Proteus vulguris ( 9 ), and others (10).The enzyme from each organism is strikingly similar, but some differences are apparent. Although in no case has the enzyme been purified to homogeneity, much evidence exists that the ribonucleoside 2’,3’-cyclic phosphate diesterase activity and the 3’-nucleotidase activity reside in the same protein. Thus, in all cases the ratio of the two activities remained constant throughout purification which has varied from 130-fold for the P . mirabilis enzyme ( 4 ) to 2000-fold for the enzyme from V . alginolyticus ( 6 ) .Anraku (3) found that both activities from E . coli B had the same optimal pH, both showed the same behavior to activators such as Co2+, and to inhibitors [Zn2+, Cuz+, ethylenediaminetetraacetate (EDTA) 3 , both were activated simultaneously by heating at 55” for 5 min and 1. H. G. Khorana, “The Enzymes,” 2nd ed., Vol. 6, p. 79, 1961. 2. Y . Anraku, JBC 239, 3412 (1984). 3. Y . Anraku, JBC 239, 3420 (1964). 4. M. S. Center and F . J. Behal, JBC 243, 138 (1968). 5. M. S. Center and F. J. Behal, ABB 127, 391 (1988). 6. T. Unemoto and M. Hayashi, BBA 171, 89 (1969). 7. T. Unemoto, F. Takahashi, and M. Hayashi, BBA 185, 134 (1969). 8. K. Shimada and Y. Sugino, BBA 185, 367 (1969). 9. H . C. Neu, Biochemistry 7, 3774 (1968). 10. H. C. Neu and J . Chou, J . Bacteriol. 94, 1934 (1967).

16.

357

NUCLEOSIDE CYCLIC PHOSPHATE DIESTERASES

were lost at temperatures above 70”. Center and Behal ( 4 ) were unable to separate the two activities from P . mirabilis by starch gel electrophoresis or by density gradient centrifugation. Like the E . coli enzyme both activities in P . mirabilis were similarly affected by metal ions, both possessed identical heat stabilities, and treatment with trypsin and chymotrypsin inactivated each to the same extent. Similar studies with the enzyme from B . subtilis (8) and the Enterobacteriaceae (9) support the conclusion that the diesterase and 3’-nucleotidase represent a single protein.

A. PROPERTIES OF THE ENZYME 1 . Substrate Specificity

The activity of the enzyme from the various organisms for a variety of phosphate esters is given in Table I. The enzyme is highly specific TABLE I RELATIVE RATESOF HYDROLYSIS OF VARIOUSPHOSPHATE ESTERSBY NUCLEOSIDE DIE ST CYCLIC PHOSPHATE DIESTERASE

Compound 3’-UMP 3’-AMP 3’-CMP 3’-GMP Cyclic 2‘,3’-UMP Cyclic 2’,3’-AMP Cyclic 2‘,3’-CMP Cyclic 2’,3’-GMP 5’-UMP 5’-AMP 5’-CMP 5’-GMP bis(p-Nitrophenyl) phosphate p-Ni tropheny1 phosphate 2’-AMP Cyclic 3’,5’-AMP 4

E. coli B (Refs. 8,3 ) O 100 90 78 95 90 87 75 70 0.9 3.3 4.6 0.9 33

P. mirabilis (Ref. 4 P

6, 7)

100 100 88 138 65

100 100 106 153 350

51

117

71 0.005 0 0.005 0 151

0.01 0

V. alginolyticus (Refs.

0.0002 0

168

B. S. sublilzs sonnei (Ref. 8)b (Ref. 9)

P. vulgaris (Ref. 9)

100 100 86 100

100 111 98 105 76 55 58

100 100 90 103 65 55 60

0.01 0.01

0

0

0.86

36

48

0.0001

31

22

0.01

Mixed 3’(2’) isomers were actually used as substrates.

* The B. sublilis enzyme also hydrolyzes 3’-deoxynucleotides.

0.001 0

0.001 0

358

G . I. DRUMMOND AND M . YAMAMOTO

for ribonucleoside 3’-phosphates and ribonucleoside 2’,3’-cyclic phosphates ; bis (p-nitrophenyl) phosphate is also hydrolyzed. p-Nitrophenyl phosphate is hydrolyzed, but a t a rate much slower than 3’-AMP except with the enzyme from Enterobacteriaceae. The B. subtilis enzyme is unique in that 3’-deoxyribonucleotides are attacked. Ribonucleoside 2‘and 5‘-phosphates, adenosine 3’,5’-cyclic phosphate, ribonucleoside diand triphosphates, hexosephosphates and pentose phosphates, and pglycerol phosphate are not attacked. I n addition most preparations are free of DNase and RNase activities, and the enzyme clearly does not split internucleotide bonds. Its ability to split the cyclic diester group of cyclic-ended oligonucleotides does not seem to have been clearly delineated. It can open the cyclic phosphodiester linkage of short cyclicended oligonucleotides such as ApA-cyclic-p and ApApA-cyclic-p but a t rates much slower than for cyclic 2’,3’-AMP (11). It has also been reported capable of opening the cyclic phosphate ends in TMV-RNA but is unable to release the resulting monoesterified phosphate ( l a ) . 2. Physical and Chemical Properties Several characteristic properties of the enzyme from the various microorganisms are given in Table 11. In general, there are similarities with regard to pH optima, heat sensitivity, activation and inhibition by metal ions, and molecular weight. Differences, however, are apparent, namely, the lower pH optimum of the enzyme from E. coli B and B. subtilis, the marked activation of the E. coli enzyme a t 55”, and the marked activation of the enzyme from this organism and other Enterobacteriaceae by Co2+.I n addition the enzyme from slightly halophilic V . alginolyticus is unique in that the 3’-nucleotidase is activated by C1- while the cyclic diesterase activity is inhibited by this anion. Some of these differences of ,course may be experimental rather than real, resulting perhaps from slight variations in assay procedures and from the presence of nonenzyme impurities in the different preparations. 3. Kinetics and Mechanism of Action

Some kinetic constants for several substrates are given in Table 111; K , values for each substrate are roughly comparable for the enzyme from each organism, and V,,, values indicate the degree of reactivity for each substrate. Several kinetic studies have provided evidence that 11. Y. Anraku and D. Mieuno, J . Biochem. ( T o k y o ) 61, 81 (1967). 12. N. Pfrogner, A. Bradley, and H. Frankel-Conrat, BBA 142, 105 (1967).

SOME Organism

E. coli B P . mirabilis

PROPERTIES OF THE

Ref.

PH Optimum

(4 3)

6.4

(4)

7.5-8

(6,7)

7.68.3

B. subtilis

(8)

6.5

Enterobacteriaceae

(3)

7.2-8.0

V . alginolyticus

TABLE I1 2',3'-cYCLIC PHOSPHATE DIESTERASE FROM &IICROORG.4NISMS Heat sensitivity

Effect of metal ions

&fold activation a t 55"; inactivated a t 70" Partial inactivation at 55" and 65"

Activated 3- to 5-fold by Co2+(0.1 to 1 d). Inhibited by Zn2+and Cuz+ (1 mM) and by EDTA (0.1 mM) No activation by Co*+and slight inhibition a t 1 mM. Inhibited by Zn2+(0.1 and 1 mM) and by EDTA (1 mM) No activation by Cot+. Inhibited by Zn2+, Hg2+, and Cuz+ (0.2 mM) and by EDTA (0.2 mM). 3'-Nucleotidase markedly activated by C1(200 mM) but cyclic diesterase inhibited by ClNot activated by Co2+ and inhibited above 1 mM. Inhibited by Zn2+ (1 mM) Stimulated 2- to 3-fold by Coz+and Mn2+(10 mM). Inhibited by Zn2+ (10 mM)

Partial inact,ivation a t 55"

Stable a t 70" for 5 min Partial inactivation at 55"

Molecular weight 68,000

65 t o 0 0

50,000

57,400

360


TABLE I11 KINETICCONSTANTS FOR

THE

CYCLIC

3‘-AMPa

3’-UMPo

Organism

K m

Vmax

Km

E. wli P . mirabilis v.algionolyticvsb B. 8ubtdh S . aonnei P . VUlgaTis

0.50 0.22 0.80 0.40 0.45 0.50

900

0.53 1610

660

0.50

a

0.22 0.70

PHOSPHATE DIESTERASE

Cyclic 2‘,3’UMPo Km

Vmax

Vmsx

bi5 @Nitrophw-1) phosphate Km

Vmax

2 . 8 1000 5000 0.94 0.75 360 3 . 6 9 . 3 X 106 4 . 5 1020 7.0 77 C 0.2 6.7

K, values are those given X lo-‘ M, and Vmaxvalues are pmolelmg protein/hr.

* Values listed are for those assays without C1-. Value for cyclic 2’,3’-AMP is 2.7 X 106.

cyclic phosphate diesterase and 3’-nucleotidase activities represent different active sites on the enzyme protein. To examine this Anraku (3) measured the velocity constants (k, and k,) of both reactions involving hydrolysis of cyclic 2’,3’-UMP. ki

kt

cyclic 2’,3‘-UMP -+ 3’-UMP -+ U

+ Pi

These were differently affected by different procedures. For example, when the enzyme was activated at 55”, the increment in k , was slight, but k, increased 3.5-fold. Similarly, in the presence of EDTA, k , and k , values decreased independently, suggesting that the sites for both activities were different. Center and Behal ( 6 ) found that with the P . mirabilis enzyme, cyclic 2’,3’-UMP competitively inhibited the hydrolysis of bis(p-nitrophenyl) phosphate. The K i was 40 pl! very close to the K , for the cyclic nucleotide (Kml75 pl!) which indicated that the two compounds could serve as alternate substrates being hydrolyzed at the same active site. In contrast, 3‘-AMP was a mixed inhibitor of cyclic 2’,3’-UMP and bis (p-nitrophenyl) phosphate hydrolysis. Adenosine was a mixed inhibitor of bis (p-nitrophenyl) phosphate hydrolysis but a competitive inhibitor of 3’-AMP hydrolysis. From such kinetic studies Center and Behal ( 6 ) suggested that two separate and adjacent sites A and B are involved in the hydrolysis of the diester and phosphomonoester substrates. Site A serves as a binding site for hydrolysis of ribonucleoside 2’,3’-cyclic phosphates and together with site B catalyzes the hydrolysis of the diester bond. During this reaction 3‘-

16.


361

UMP becomes bound to site B which catalyzes the hydrolysis of the phosphomonoester bond. Adenosine and 3’-AMP by binding at site B could interfere with the breakdown of cyclic 2’,3’-UMP. Similarly, binding of bis (p-nitrophenyl) phosphate at site A could interfere with the breakdown of 3’-AMP. Cyclic 2’,3’-UMP and bis (p-nitrophenyl) phosphate compete for site A while adenosine competes with 3’-AMP for site B. Unemoto et al. ( 7 ) have examined the mutual inhibition of substrates and substrate analogs for the enzyme from halophilic V . alginolyticus. They also concluded that 3‘-ribonucleotides and ribonucleoside 2’,3’-cyclic phosphates are hydrolyzed a t difierent sites. However, because of the nature of the mutual inhibition between 3’-AMP and bis(p-nitrophenyl) phosphate, they suggested that part of the site for the latter substrate overlaps with the 3’-nucleotidase site. At this time the precise mechanism of action of the enzyme is not settled, but clearly there are two active sites, one a 3’-nucleotidase site and a cyclic phosphate diesterase site. Anraku (13) has described this protein as a “double-headed” enzyme.

B . CELLULAR LOCALIZATION The cyclic phosphate diesterase is found in the cell sap upon fractionation of bacterial extracts, and when such extracts are centrifuged a t 100,OOO x g i t is present in the supernatant fluid. Neu and Heppel (14) found the enzyme was released into solution when EDTA-lysozyme spheroplasts (16) were prepared from E . coli. This pointed to a surface localization of the enzyme as had been suggested for alkaline phosphatase, another enzyme known to be released during spheroplast formation ( 1 6 ) . Neu and Heppel (14) have also shown that the diesterase is released from E. coli cells by a process of osmotic shock. I n this procedure (17) cells preferably in the exponential growth phase are suspended in hypertonic sucrose, sedimented, and rapidly subjected to dispersal in solution of low ionic strength. About 4% of the cellular protein is released and most of the cells remain viable ( 1 7 ) . Cyclic phosphate diesterase is one of a family of degradative enzymes (17-19) including alkaline phosphatase, ribonuclease I, 5’-nucleotidase (see Chap13. 14. 15. 16. 17. 18. 19.

Y. Anraku, Procedures Nucleic Acid Res. p. 130 (1966). H. C. Neu and L. A. Heppel, BBRC 17, 215 (1964). R. Repaske, BBA 30, 225 (1958). M. H. Malamy and B. L. Horecker, Biochemistry 3, 1889 (1964). L. A. Heppel, Science 156, 1451 (1967). H . C. Neu and L. A. Heppel, JBC 240, 3685 (1965). N. G. Noasal and L. A. Heppel, JBC 241, 3055 (1966).

362

Q.

I. DRUMMOND AND M. YAMAMOTO

ter 15 by Drummond and Yamamoto, this volume) and others that are released into solution. The diesterase is also released during osmotic shock from other members of the Enterobacteriaceae such as Shigella, Enterobacter, Citrobacter, Serratia, and Salmonella (9, 10) ; this technique has provided an effective alternate procedure for purification (9, IS).The nature of this process suggests that these enzymes exist external to the cytoplasmia membrane and within the cell wall. Brockman and Heppel (20) have provided further evidence that the enzyme is located near the surface of E . coli external to the protoplasmic membrane. Thus, uridine 2’,3’-cyclic phosphate was hydrolyzed by intact cells suggesting the enzyme lay outside the permeability barrier for phosphate esters. Enzymic activity with intact cells, however, was less than that observed with equivalent amounts of cell extract. This suggested a partial barrier to penetration; the enzyme may be located between the cell wall and the cytoplasmic membrane in the “periplasmic space” (17’). Histochemical evidence (21) supports such a localization. The suggestion has also been made (10)that these enzymes are probably loosely bound to cytoplasmic membrane through the mediation of divalent cations. Whatever the precise location it seems probable that such degradQtive enzymes are confined in a compartment separate from those regions of the cell where synthetic reactions are taking place. Such compartmentation may be analogous to the localization of certain degradative enzymes in mammalian lysosomes. C. CYCLICPHOSPHATE DIESTERASE-AMETALLOENZYME When E . coli are grown in the presence of low concentrations of EDTA, a selective depression of cyclic phosphate diesterase (as well as 5’-nucleotidase and alkaline phosphatase) occurs. Dvorak (28) suggested that EDTA may act by binding a trace metal ion essential for the activity of the enzymes. To investigate this, E . coli cells were grown in the presence of radioactive s5Zn and cyclic phosphate diesterase and 5‘-nucleotidase were purified from shock fluid (23).The purification was accompanied by enrichment of 66Zn which could not be removed by dialysis. On chromatography and disc gel electrophoresis a superimposable peak of en2yrni.c activity and 66Zn were observed for both enzymes. Both enzymes were inactivated by prolonged exposure to EDTA and partial reactivation occurred upon the addition of Zn2+. The pos-

R.W. Brockman and L. A. Heppel, Biochemistry 7, 2554 (1968). 21. S. 5.Spicer, B. K. Wetzel, and L. A. Heppel, Federation Proc. 25, 539 (1966). 22. H. F. Dvorak, JBC 243, 2640 (1968). 23. H. F. Dvorak and L. A. Heppel, JBC 243, 2647 (1968). 20.

16. NUCLWIDE

363

CYCLIC PHOSPHATE] DIE ST ERAS^

sibility is thus considered (93)that the diesterase and 5’-nucleotidase of E . coli are zinc metalloenzymes.

D. PHYSIOLOGICAL FUNCTION Since ribonucleoside 2‘,3’-cyclic phosphates are formed from degradation of RNA by ribonuclease, Anraku (3) has suggested that the cyclic phosphate diesterase acting together with ribonuclease may be involved in the reutilization of nucleotides. Some evidence for this exists. Purified cyclic diesterase hydrolyzes ribonucleoside 2’,3’-cyclic phosphates about lo00 times more rapidly than purified rRNase (11). Furthermore, the ratio of their activities in vivo is about 600:l indicating that cyclic phosphate diesterase is the major enzyme concerned with the digestion of nucleoside cyclic phosphates. I n support of this, it has been shown that the diesterase is capable of opening the cyclic diester linkage of short cyclic-ended oligonucleotides such as ApA-cyclic-p and ApApAcyclic-p (11). Based on these facts Anraku and Mizuno (11) have proposed an RNase-cyclic phosphate diesterase system in which the diesterase acts on oligonucleotides with 2’,3’-cyclic phosphate end groups and catalyzes the rapid reutilization of cyclic phosphates by converting them to the corresponding nucleosides as shown in the scheme: RNA

___t

Xpyp

f

RNase

RNA

x p y p ----------- z p

/*

1

..--------. Z-cyc1ic-p _ _ _ t

- -

cyclic phosphate diesterase

X-cyclic-p

3’-nucleotidase

t/

I(

3’-XMP

nucleosides

+ Pi

A suggestion that such a system functions in the breakdown of ribosomal RNA in vivo has been provided by Maruyama and Mizuno (93a).They showed that the degradation of ribosomal RNA induced by phosphorus starvation was catalyzed by the primary attack of RNase on ribosomes followed by the accumulation of oligonucleotides with 2‘,3‘-cyclic phosphate end groups and these were subsequently converted to nucleosides and inorganic phosphate presumably by cyclic phosphate diesterase.

111. Ribonucleoside 2’,3‘-Cyclic from Vertebrate Nerve

Phosphate Diestemre

Prior to 1961, ribonucleoside 2’,3’-cyclic phosphate diesterase activity had been reported from two vertebrate tissues, calf spleen (94)and beef 23a. H. Maruyama and D. Mizuno, BBA 108, 593 (1965). 24. P. R. Whitfield, L. A. Heppel, and R. Markham, BJ 80, 16 (1956).

364

Q. I. DRUMMOND AND M. YAMAMOTO

pancreas (26) [see Khorana ( I ) ] . More recently, Drummond et al. (26) have described an extremely active diesterase from nerve tissue. A. PROPERTIES AND SUBSTRATE SPECIFICITY This enzyme converts ribonucleoside 2',3'-cyclic phosphates exclusively to the corresponding 2'-phosphates. The enzyme has greater specificity for 2',3'-cyclic phosphates bearing purine bases than those with pyrimidine bases. Activities in crude extracts of brain may be as high as 5-10 pmoles/min/mg of protein at 30". The K,,, for cyclic 2',3'-AMP is 1.9 d. The enzyme does not hydrolyze simple nucleoside phosphate esters and does not split internucleotide bonds of dinucleotides, oligonucleotides, or RNA. It does not hydrolyze ribonucleoside 3',5'-cyclic phosphates. The enzyme opens the cyclic diester linkage of 2',3'-cyclicended dinucleotides such as GpC-cyclic-p and ApC-cyclic-p with formation of the corresponding 2'-ended dinucleotide (26). The 2',3'-cyclic diester bond of longer cyclic-ended oligonucleotides, for example, Ap (Ap),A-cyclic-p, is also cleaved without rupture of internucleotide bonds. The enzyme possesses a pH optimum between p H 6 and 7; it is strongly inhibited by Zn2+and Hg2+and activated slightly by EDTA and citrate. Tissue from the central nervous system is the richest source of the enzyme; it is also present in peripheral nerve. White matter contains much more activity than gray matter. Relatively insignificant activity resides in tissues other than nerve.

B. INTRACELLULAR LOCALIZATION IN MYELIN The enzyme was originally found to be membrane bound and resisted solubiliaation and purification (26).Lundblad and Moore (27') , however, have reported solubilizing it using dilute (5 mM) sodium borate buffer at pH 9 after 16 hr a t 37". Studies on regional and subcellular distribution using density gradient techniques have revealed that the 2',3'-cyclic phosphate diesterase concentrates in those fractions containing myelin (28, 2 9 ) , and the conclusion has been reached that the enzyme is localized in the myelin sheath or intimately associated structures. Kurihara and 25. F. F. Davis and F. W. Allen, BBA 21, 14 (1956). 26. G. I. Drummond, N. T. Iyer, and J. Keith, JBC 237, 3535 (1962). 27. R.L. Lundblad and S. Moore, Brain Res. 12, 227 (1969). 28. T. Kurihara and Y . Tsukada, J . Neurochem. 14, 1167 (1967). 29. R. W. Olafson, G. I. Drummond, and J. F. Lee, Can. J . Biochem. 47, 961 (1969).

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Tsukada (30) have examined developmental changes of the enzyme in chick brain and spinal cord. Enzymic activity appears a t about the eighteenth day of incubation and increases rapidly until 3 days after hatching in the brain and between 18 and 21 days of incubation in the spinal cord. These are precisely the periods of active myelination in the brain and spinal cord of the chick, respectively. Similarly, brain tissue of the newborn rat is devoid of cyclic phosphate diesterase activity; it appears at about 8 days after birth and increases dramatically between the tenth and thirty-fifth day of life (29). This coincides precisely with the development of myelin in this species. The diesterase is essentially absent in the brain of the “jimpy” mouse (SI),a lethal mutant devoid of myelin in the central nervous system. It is also absent from the spinal cord of this mutant. The enzyme is about 50% deficient in brain tissue of the “quaking” mouse (29), a mutant with partial deficiency of myelin. There is no activity in nerve fibers and ganglia from a variety of invertebrates such as squid, octopus, crab, shrimp, and starfish. Nerve tissue in these organisms is nonmyelinated. All these observations point to an intimate association of the enzyme with myelin in vivo.

C. PHYSIOLOGICAL ROLEOF

THE

CYCLIC PHOSPHATE DIESTERASE

Nothing is known of the physiological function of this enzyme. No biological function is known for ribonucleoside 2’,3’-cyclic phosphates or cyclic-ended oligonucleotides, and if they occur in nerve tissue they are surely below the sensitivity of the usual methods of detection. Moreover, myelin is a tissue which has been considered essentially devoid of metabolic activity. The presence of such a highly active enzyme raises interesting possibilities regarding a role of nucleoside 2’,3’-cyclic phosphates in nerve function. The possibility should even be considered that the physiological substrate is something other than a nucleoside cyclic phosphate. The association of the enzyme with myelin is of interest with regard to its possible involvement in demyelinating diseases.

IV. Nucleoride 3’,5’-Cyclic Phosphate Diesterase

Following the discovery of adenosine 3’,5’-cyclic phosphate, an activity present in various animal tissues capable of destroying the compound 30. T. Kurihara and Y. Tsukada, J . Neurochena. 15, 827 (1968). 31. T. Kurihara, J. L. Nussbaum, and P. Mandel, Brain Re8. 13, 401 (1969).

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Q. I. DRUMMOND AND M. YAMAMOW

was reported by Sutherland and Rall in 1958 (38).The activity was shown to be a Mg2+-dependent diesterase which catalyzed the hydrolysis of cyclic 3’,5’-AMP at the 3’ position forming 5‘-AMP. Since that time knowledge of the physiological role played by cyclic 3’,5’-AMP as a mediator of hormone action and modulator of enzymic activity has expanded rapidly [for reviews, see Sutherland et al.’ ( 3 6 )]. The 3’,5’cyclic phosphate diesterase has attracted considerable attention because of its important role (along with adenyl cyclase) in regulating intracellular levels of cyclic 3’,5’-AMP. A. DISTBIBUTION OF

THE

ENZYME EN NATUBE

Early studies revealed that the 3’,5’-cyclic phosphate diesterase is present in all mammalian tissues (39, 33, 3 6 ) , being most active in cerebral cortex (36, 37). It has also been identified in extracts of liver fluke (FascioZa hepatica), the common earthworm ( L u m b r h terresth) , and fly larvae (36) ; and it has been studied in marine organisms (38), the cellular slime mold Dictyostelium discoideum (39, 40), and in E. coli (41). The enzyme has been partially purified from beef heart (369,dog heart, (42) and bovine brain (37,@) . No highly purified preparations have yet been obtained and most studies have been performed with relatively crude preparations.

B. SUBSTRATE SPECIFICITY The enzyme from brain hydrolyzes cyclic 3’,5‘-monophosphates bearing purine bases more readily than those bearing pyrimidines (37,&). 32. E. W. Sutherland and T. W. Rall, JBC 232, 1077 (1958). 33. E. W. Sutherland and T. W. Rall, Phamnacol. Rev. 12, 265 (1960). 34. E. W. Sutherland, G. A. Robison, and R. W. Butcher, Circulation 37, 279 (1968).

35. G. A. Robison, R. W. Butcher, and E. W. Sutherland, Ann. Rev. Biochm. 37, 149 (1908).

36. R. W. Butcher and E. W. Sutherland, JBC 237, 1244 (1962). 37. G. I. Drummond and S. Perrot-Yee, JBC 236, 1128 (1961). 38. M. Yamamoto and K. L. Massey, Comp. Biochem. PhvWZ. 30, 941 (1969). 39. Y. Y. Chang, Science 161, 57 (1968). 40. J. T. Bonner, D. S. Barkley, E. M. Hall, T. M. Konijn, T. W. Mason, G. O’Keefe, 111, and P. B. Wolfe, DeveZop. BWZ. 20, 72 (1969). 41. H. Bra& and F. Chytil, F o l k Mkrobiol. (Prague) 11, 43 (1965). 42. K. G. Nair, Biochemistry 5, 150 (1960). 43. W. Y. Cheung, BBA 101, 303 (1969). 44. G. I. Drummond, M. W. Gllgan, E. J. Reiner, and M. Smith, JACS 86, 1026 (1964).

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Cyclic 3‘,5‘-CMP is not attacked. Deoxyribonucleoside 3’,5’-cyclic phosphates are hydrolyzed a t rates slightly less than the corresponding ribonucleoside derivatives ( 4 4 ) . Similar specificity is shown by the dog heart (42) and rat liver enzyme (45).Tubercidin 3’,5’-cyclic phosphate (tubercidin is 7-deaza adenosine) is hydrolyzed about three times more rapidly than cyclic 3’,5’-AMP by the rabbit brain enzyme (46). I n all cases that have been examined, the product of hydrolysis is exclusively the 5’-phosphate (39, 36, 37, 4.4). These limited studies indicate a rather low degree of substrate specificity with regard to the base moiety. There seems to be a much more rigid requirement for an intact phosphate and sugar moiety. Thus, substitution of a butyryl group on C-2 of ribose greatly reduces reactivity (47) and N8,2’-O-dibutyryl cyclic 3‘,5’-AMP is completely resistant to hydrolysis (4.6, 46). Moreover, adenosine 3’,5’cyclic phosphorothioate and a 3’-methylene cyclic phosphonate analog (the 5’-cyclic ester of 9- [3’-deoxy-3’-dihydroxyphosphinylmethyl-~-~ribofuranosyl] -adenine) are neither substrates nor effective inhibitors (46). I n addition, the 5’-methylene cyclic phosphonate compound and adenine xylofuranosyl 3’,5’-cyclic phosphate are extremely poor substrates. Precise understanding of the mechanism of action of the enzyme must await the availability of highly purified preparations and systematic kinetic studies,

C. INTRACELLULAR LOCALIZATION Some investigators have found the enzyme primarily in the 100,000 x g supernatant fluid of extracts of rabbit brain (37), dog heart (&), rat liver (46), rat brain, and beef pineal gland (48). I n contrast, other

investigators have reported that the enzyme from beef heart (36)and rat brain (49, 50) is mostly particulate being present in all the primary fractions nuclear, mitochondrial, microsomal, and 100,000 x g supernatant. The reasons for these discrepancies are not entirely clear but most likely result from differences in fractionation technique. Evidence from density gradient procedures coupled with electron microscopic examination indicate that the enzyme is preferentially located a t nerve 45. L. A. Menahan, K. D. Hepp, and 0. Wieland, European J . Bwchem. 8, 435 (1969). 46. G. I. Drummond and C. A. Powell, Mol. Pharmacol. 6, 24 (1970). 47. Th. Posternak, E. W. Sutherland, and W. F. Henion, BBA 65, 558 (1962). 48, B. Weiss and E. Costa, Biochem. Pharmacol. 17, 2107 (1968). 49. E. De Robertis, G . R. Arnaiz, M. Alberici, R. W. Butcher, and E. W. Sutherland, JBC 242, 3487 (1967). 50. W. Y. Cheung and L. Salganicoff, Nature 214, 90 (1967).

368


endings in brain tissue (38, 49, 60).The enzyme has also been demonstrated in various tissues by histochemical techniques (61,66).

D. METALIONREQUIREMENT, pH OPTIMUM AND SUBSTRATE AFFINITY The diesterase from most tissues has a pH optimum in the neutral or slightly alkaline range (36, 37, 42, 46,5 3 ) , and requires a divalent metal for optimal activity. Usually, Mg2+and Mn2+are equieffective; Zn2+is a powerful inhibitor (3'7, 38, 5 3 ) . Several K,,, values of the diesterase for cyclic "$'-AMP have been reported: beef heart 0.1 mM (36))dog heart 0.49 mM (&), rat liver 0.62 mM (&), rat brain 0.1-0.3 mM ( 5 0 ) ,E . coli 0.77 mM (41), and for the slime mold 2 mM (39). Only rough approximations of the molecular weight are available; these are 300,000for the slime mold (39) and 200,000for the rat liver enzyme (45).

E. INHIBITOM AND ACTIVATORS A wide variety of compounds inhibit the diesterase in vitro; a summary is presented in Table IV. Inhibition by the methyl xanthines has been most studied (36, 42, 53, 54) and is of pharmacological interest because it seems to account for the effects of these agents on intracellular levels of cyclic 3',5'-AMP in a variety of tissues, especially in the presence of hormones that stimulate adenyl cyclase. The slime mold enzyme is not inhibited by caffeine a t 10 mM (39) and the E . coli enzyme is not inhibited either ( 4 1 ) . Cheung (53, 55) has found that the brain enzyme is inhibited by both ATP and inorganic pyrophosphate ; and since these compounds are substrate and product of adenyl cyclase, respectively, he has suggested the inhibition may represent a physiological mechanism for controlling the enzyme in vivo. A variety of phenothiazine tranquilizers such as chlorpromazine are more potent inhibitors of the rabbit brain enzyme than the methyl xanthines (54). Whether this has any meaning in terms of the behavioral effects of these agents is of course not known. Interest is increasing, however, in the possibility that cyclic 3',5'51. T. R. Shanta, W. D. Woods, M. B. Waitsman, and G. H. Bourne, Hbtochemie 7, 177 (1966). 52. B. McL. Breckenridge and R. E. Johnston, J . Hbtochem. Cytochem. 17, 505 (19e9). 53. W. Y. Cheung, Biochemistry 6, 1079 (1987). 54. F. Honda and H. Imamura, BBA 161, 287 (1968). 55. W.Y . Cheung, BBRC 23, 214 (1966).

16.

369


TABLE IV INHIBITORS OF NUCLEOSIDE CYCLIC PHOSPHATE DIESTERASE Ki or inhibitory conc.

Enzyme source

Type of inhibition

Theophylline

Beef heart

Competitive

Caffeine T heophylline Caffeine ATP, CTP, UTP, ITP inorganic pyrophosphate Puromycin

Dog heart Rabbit brain Rat brain Rat brain

Noncompetitive 0.5 Competitive 0.11 Competitive 0.3 Mixed 50% inhibition at 3 mM Competitive 0.8

Compound ~

(d) Ref.

~~~

3,3’,5’-Triiodo-~thyronine Chlorpromaaine and other phenothiazines Ethacrynic acid Chlorthalidone Acetazoleamide Diazoxide (3-methyl-7chloro-1,2,4-benaathiadiazine-1,l-dioxide)

Rat diaphragm Rat adipose tissue and beef heart Rabbit brain Rat kidney Rat kidney Rat kidney Beef heart

Competitive

0.1

0.4

Significant at 0.05 Noncompetitive 0.4 Noncompetitive 2.5 Noncompetitive 6.0 Noncompetitive 50y0 with 0.45

AMP is related to certain mental diseases and may be involved in the action of tranquilizers and antidepressant drugs (60). Whether the ability of diuretic agents such as ethacrynic acid and chlorthalidone to inhibit the enzyme in kidney (58) is related to their diuretic action is also not known. It has been suggested that inhibition of diesterase by diazoxide (59) may explain the hyperglycemic activity of this agent. Several materials are known to activate the enzyme. Imidazole produces strong activation of the enzyme from mammalian tissues (36, 38,4%’) but not from E . coli (41). It has been reported (61) that insulin activates the beef heart enzyme in vitro, but it is not known if this has relevance M. M. Appleman and R. G. Kemp, BBRC 24, 564 (1966). 57. L. R. Mandel and F. A. Kuehl, Jr., BBRC 28, 13 (1967). 58. G. Senft, K. Munske, G. Schulta, and M. Hoffman, Arch. Exptl. Pathol. 56.

Pharmakol. 259, 344 (1968). 59. G. Schultz, G. Senft, W. Losert, and R.Sett, Arch. Ezptl. Pathol. Phannakol. 253, 372 (1966). 60. Y. H. Abdulla and K. Hamadah, Lancet I , 378 (1970). 61. G. Senft, G. Schultz, K. Munske. and M. Hoffman, DkbetologM 4, 330 (1968).

370


to the effects of this hormone on cyclic 3’,5’-AMP levels in vivo. Most of these studies have shed little light on the kinetic properties and mechanism of action of the enzyme. Instead they have been motivated largely by efforts to describe the effects of various drugs and hormones on physiological processes. I n addition to the proposed regulatory role of ATP and pyrophosphate, some possibility exists that 3’,5’-cyclic phosphate diesterase is under physiological control. Such ideas arose through observations of Cheung (43, 62) that the partially purified enzyme from beef brain was markedly activated by snake venom. The stimulatory factor was labile a t extreme pH; it was not dialyzable and appeared to be a protein. A similar activating factor is also present in brain tissue (63) and is removed during purification of the diesterase. It seems t o interact stoichiometrically with the enzyme. The activator is destroyed by trypsin and is not proteolytic itself. The precise role of this protein in regulating the phosphodiesterase in vivo is not yet established, however.

F. POSSIBILITY OF OTHER NUCLEOSIDECYCLIC 3’,5’-PHOSPHATE DIESTERASES It must be emphasized that much of the information on the enzyme to date comes from partially purified preparations. Even though it seems uniquely specific for nucleoside 3’,5’-cyclic phosphates, the possibility cannot be excluded that in some tissues more than one diesterase may be present. I n fact, Hardman and Sutherland (64) have shown that a second 3’,5’-cyclic phosphate diesterase exists in heart muscle which is primarily specific for uridine 3’,5’-cyclic phosphate. This enzyme was found to be more sensitive to inhibition by theophylline and t o activation by imidazole. Further studies are needed to clarify its precise status; other such enzymes may be discovered in the future. G. PHYSIOLOGICAL FUNCTION OF NUCLEOSIDE CYCLIC PHOSPHATE DIESTERASE The physiological significance of the diesterase cannot be minimized. Cyclic 3‘,5’-AMP has many actions and its concentration in the cell is dramatically affected by various agents, particularly neurohormones of 62. W. Y.Cheung, BBRC ZQ, 478 (1967). 63. W. Y.Cheung, BBRC 38, 633 (1970). 64. J. G. Hardman and E. W. Sutherland, JBC 240,

PC3704 (1986).

16.


371

the sympathetic nervous system and other hormones that mediate important biological processes (33-35). It is paramount that cells have a mechanism for terminating the effects of this regulator in order to maintain homeostasis. The diesterase clearly performs this function. It is reasonable to assume that some hormones and even drugs may alter intracellular levels of cyclic 3’,5’-AMP by affecting its catabolism, that is, by an action on nucleoside 3’,5’-cyclic phosphate diesterase.


17 E . coli Alkaline Phosphatase TED W . REID

IRWIN B . WILSON

I. Introduction . . . . A . Historical Background

B . Distribution . . . C . Function . . . I1. Molecular Properties . . A . Purification . . . B . Composition . . . C . Subunits . . . D. Isozymes . . . E . Physical Properties . F. Crystal Structure . G . Chemical Modification 111. Catalytic Properties . . A . Specificity . . . B . Competitive Inhibitors C. The Phosphoryl Enzyme D . The Role of Zinc . E . Number of Active Sites F. Transphosphorylation G. Kinetic Studies . .

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373 374 376 376 377 377 378 380 384 387 389 389 392 392 394 396 401 404 406 409

.

1 Introduction

A . HISTORICAL BACKGROUND Alkaline phosphatase from E . coli is an enzyme of the 1960’s. Although one brief reference to a phosphatase from E . coli having an alkaline p H maximum was reported in 1933 (1). it was not until the discovery by

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1 A . Boivin and L Mesrobeanu. Compt . Rend . Soc Biol 112. 611 (1933)

373

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T. W. REID AND I. B. WILSON

Horiuchi et al. ( d ) , and Torriani (3) that orthophosphate repressed the formation of a nonspecific phosphomonoesterase in E . coli that research on this enzyme began. This work (2, 3) showed a maximum rate of synthesis of the enzyme occurred only when the phosphate concentration became low enough to limit cell growth. With sufficient phosphate, the amount of active enzyme is negligible. Under conditions of limiting phosphate, alkaline phosphatase accounts for about 6% of the total protein synthesized by the cell ( 4 ) .

B. DISTRIBUTION Although the enzyme sediments with intact cells, alkaline phosphatase appears in the supernate when broken cells are centrifuged. Malamy and Horecker ( 5 ) discovered that alkaline phosphatase is quantitatively released from the cell when E . coli are converted to spheroplasts by lysozyme and ethylenediaminetetraacetic acid (EDTA) in a sucrose medium. This evidence, supported by the observation that substrates such as glucose 6-phosphate are rapidly hydrolyzed by intact cells with release of most of the phosphate into the medium, led Malamy and Horecker (6) to suggest that alkaline phosphatase is localized in the periplasmic space, a region described by Mitchell (7) as lying between the protoplasmic membrane and the wall layer, and that it is not in association with the wall (8). Later, it was found that alkaline phosphatase was also released by osmotic shock: E . coli were exposed to 0.5 M sucrose containing dilute tris-HC1 buffer and EDTA, and then the centrifuged cells were rapidly dispersed in the ‘‘shock medium” of cold water or cold 5 X lo-’ M MgCL Although the cells were 80% viable with the latter case, almost all of the enzyme was released (9, 10). Other evidence indicates that the only important structural effect of EDTA is to increase the permeability of the cell wall (11, 1 2 ) . Escherichia coli grow normally in the 2. T. Horiuchi, S. Horiuchi, and D. Mizuno, Nature 183, 1529 (1959). 3. A. Torriani, BBA 38, 460 (1960). 4. A. Garen and C . Levinthal, BBA 38, 460 (1960). 5. M. Malamy and B. L. Horecker, BBRC 5, 104 (1961). 6. M. Malamy and B. L. Horecker, Biochemistry 3, 1889 (1964). 7. P. Mitchell, in “Biological Structure and Function” (T.W . Goodwin and 0. Lindberg, eds.), Vol. 2. Academic Press, New York, 1961. 8. W. Weidel and J. Primosigh, J. Gen. Microbial. 18, 513 (1958). 9. H. C. Neu and L. A. Heppel, JBC 240, 3685 (1965). 10. N. G. Nossal and L. A. Heppel, JBC 241, 3055 (1966). 11. L. Leive, Proc. nNatl. Acad. Sci. U.S. 53, 745 (1965). 12. L. Leive, BBRC 18, 13 (1965).

17. E. COli

ALKALINE PHOSPHATASE

375

presence of EDTA but with a selective depression in the levels of the enzymes alkaline phosphatase, cyclic phosphodiesterase, and 5-nucleotidase (IS), which are thought to be in the periplasmic space. Localization of the enzyme in the periplasmic space is also consistent with the selective release of alkaline phosphatase during growth of an E . coli mutant which is osmotically sensitive because of a defective cell wall (14) and with the fact that phosphate esters which do not penetrate the protoplasmic membrane can be hydrolyzed by intact cells (15).In these latter measurements the activities found with intact cells as compared with equivalent cell extracts varied over wide limits depending upon the substrate and its concentration. This difference was assumed to result from a difference in the ease of penetration of the wall barrier by different phosphate esters. Histochemical studies also suggest a localization between membrane and cell wall (16-19). Most of these studies (16, 18, 19) showed a localization of the enzyme in small regions of periplasmic space. Schlesinger e t al. (20) concluded, on the basis of in vitro rates of dimerization, that the dimerization of enzyme subunits in vivo would not be as rapid as observed unless the subunits were compartmentalized in the cell. The in vitro rate of dimerization seemed to be based upon reoxidation and dimerization of reduced monomers and showed a maximum a t 65 pg/ml with respect to protein concentration. The in vivo process may be rather different, however, and later studies by Schlesinger and Barrett (21) with unreduced monomers would seem to change this conclusion because their rates did not have a maximum with respect to protein concentration. By forming spheroplasts from normal cells, Torriani (22) showed that pools of monomer but no alkaline phosphatase (active dimers) exist in the endoplasm and concluded that dimerization occurred outside the endoplasm. Fifteen percent of the enzyme exists as monomers associated with particles in the endoplasm that are larger than ribosomes. 13. H. Dvorak, JBC 243, 2640 (1968). 14. G. Mangiarotti, D. Apirion, and D. Schlessinger, Science 153, 892 (1966). 15. R. W. Brockman and L. A. Heppel, Biochemistry 7, 2554 (1968). 16. J. Done, C. Shorey, J. Loke, and J. Pollak, BJ 96, 27c (1965). 17. S. S. Spicer, B. K. Wetzel, and L. A. Heppel, Federation Proc. 25, 539 (1966). 18. V. Kushnarev and T. Smirnova, Can. J . Microbiol. 12,605 (1966). 19. V. Kushnarev, T. Smirnova, and A. Bykov, Dokl. Akad. Nauk SSSR 175, 718 (1967). 20. M. J. Schlesinger, A. Torriani, and C. Levinthal, Cold Spring Harbor Sump. Quant. Biol. 28, 539 (1963). 21. M . J. Schlesinger and K. Barrett, JBC 240, 4284 (1965). 22. A. Torriani, J . Bacterial. 96, 1200 (1968).

376

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W.

REID AND I. B. WILSON

Schlesinger (23) has shown that under conditions permitting protein synthesis spheroplasts are able to produce the subunits that appeared in the medium, but they are not able to dimerize. It appears then that the monomers are transported out of the endoplasm before being dimerized (24, 2 5 ) . However, the movement of the monomers from the endoplasm to the periplasmic space followed by their dimerization still requires explanation. One possibility is that the concentration of monomers in the endoplasm is too low to produce dimers and that the monomers are concentrated in the periplasmic space by active transport. Evidence against active transport was obtained in episomal transfer of the structural gene from E . coli to S. typhimurium ( 2 6 ) .Even though S. typhimurium does not synthesize alkaline phosphatase, the enzyme was produced by the heterogenote and appeared in the periplasmic space. Schlesinger and Olsen (26) argued that it is unlikely that S. typhimurium would have a transport system for alkaline phosphatase monomers because it does not normally make the enzyme. Another possibility is that the plasma membrane is permeable to monomers and not to dimers and that the periplasmic space contains some factor that accelerates the formation of dimers. Although Zn(I1) comes immediately to mind, it cannot be this accelerating factor because Harris and Coleman (27) have obtained inactive dimers by osmotic shock of E. coli that were grown in the absence of Zn(I1). These dimers become active immediately upon the addition of Zn(I1). Evidently the process of in vivo dimerization and localization of the enzyme remains unexplained.

C. FUNCTION Although it is widely found in bacteria, the physiological function of alkaline phosphatase is still unknown. The enzyme is nonspecific (4, 28) , and this would be desirable if its role were to supply phosphate from phosphate esters under conditions of phosphate deprivation. Although the enzyme is repressed by orthophosphate in many strains of E . coli, it is constitutive in most other bacteria (29), thus phosphate deprivation 23. 24. 25. 26. 27. 28. 29.

M. J. Schlesinger, J. Bacterial. 96, 727 (1968). M. J. Schlesinger, JBC 242, 1604 (1967). S. Schlesinger and M. J. Schlesinger, JBC 242, 3369 (1967). M. J. Schlesinger and R. Olsen, J. Bacteriol. 96, 1601 (1968). M. Harris and J. Coleman, JBC 243, 5063 (1968). L. A. Heppel, D. Harknew, and R. Hilmoe, JBC 237,841 (1962). M. Kuo and H. Blumenthal, Nature 190,29 (1961).

17. E . coli

377


does not seem to be an important factor. Even though its role could be the nonspecific hydrolysis of phosphate esters, it is nonetheless reasonable to entertain the possibility of other functions such as phosphate transport. Since the enzyme readily binds phosphate, both covalently and noncovalently, especially in acid p H (30,31), it could in principle transport and concentrate phosphate from a more acidic medium to the interior of the cell under conditions of low phosphate.

II. Molecular Properties

A. PURIFICATION The methods (and procedures) for growing E . coli and purifying alkaline phosphatase have been extensively reviewed by Torriani (32, 33) (see also Table I ) . Early methods of purifying alkaline phosphatase from E. coli involved heat shock (4), disruption of cells with a French press (4,34, 35), and TABLE I SUMMARY OF PURIFICATION OF E . coli ALKALINE PHOSPHATASE Yield Fraction 1. Extraction (a) Sonic disruption (b) Spheroplast formation (c) Osmotic shock 2. Purification (a) Heat treatment (optional) (b) (NH&SO4 concentration (optional) (c) Batchwise DEAE-cellulose (optional) (d) DEAE-cellulose chromatography (e) (NH4)2SO4 precipitation and crystallization

(%)

Specific activity“

100 90 70-100

3-6 16 18-30

100 95 95 60 100

40-46 40-46 40-46 48-50 5ob

Micromoles p-nitrophenyl phosphate/min/mg protein at 25” in 1.0 M tris (pH 8.0). Value obtained by Malamy and Horecker (39)for a crystalline preparation containing a mixture of three isozomes, adjusted to 25”. b

30. D. Levine, T. W. Reid, and I. B. Wilson, BiochemLtry 8, 2374 (1969). 31. T. W. Reid, M. Pavlic, D. Sullivan, and I. B. Wilson, Biochemistry 8, 3184 (1969). 32. A. Torriani, “Methods in Enzymology,” Vol. 12, Part B, p. 212, 1968. 33. A. Torriani, in “Procedures in Nucleic Acid Research,” p. 224, 1966. 34. A. Garen and H. Echols, J . Bacterial. 83, 297 (1962). 35. J. Schwartr and F. Lipmann, Proc. Natl. Acad. Sci. U. S. 47, 1996 (1961).

378


the preparation of acetone powders (35, 36). Purification of the enzyme was greatly facilitated by the discovery of Malamy and Horecker ( 5 ) that the enzyme is released from E . coli when spheroplasts are formed by treatment with lysozyme and EDTA in 20% sucrose (3'7). Most of the endocellular proteins (6, 38, 39) are retained by the spheroplast. It was later shown by Neu and Heppel (9,4) that alkaline phosphatase can also be released by treating the cells with EDTA in 20% sucrose followed by osmotic shock in cold water. This method has the advantage that the cells retain enzymes such as RNase which are difficult to eliminate during purification. The enzyme can be further purified by various techniques including heating a t 80" for 15 min which denatures and precipitates many proteins but not alkaline phosphatase (4, 32, 33), concentration with (NH4)&304 (15), or batchwise addition to DEAEcellulose followed by elution with 0.1 M NaCl (41). The enzyme is then chromatographed on DEAE-cellulose using a sodium chloride gradient (4, 15, 27, 38, 39, 41-44). It has been shown recently that gradient elution can separate the isozymes of alkaline phosphatase (41, 43, 44) and is also valuable in the purification of the apoeneyme (27'). I t has recently been reported (45) that when E. coli cells were suspended in 0.05 N HCl subunits of the enzyme were quantitatively released into the medium. Subsequent reassociation and reactivation of these subunits provided an initial cell free extract that contained alkaline phosphatase which was 30% pure. The enzyme was first crystallized by Malamy and Horecker (39) using ammonium sulfate.

B. COMPOSITION 1. AnaZysis

The amino acid composition of alkaline phosphatase of E. coli is given in Table I1 (41, 43, 46, 47'). The values from Lazdunski and Lazdunski 36. D. J. Plocke, C. Levinthal, and B. L. Vallee, Biochemistry 1, 373 (1962). 37. R. Repaske, BBA 30, 225 (1958). 38. C. Levinthal, E. Signer, and K. Fetherolf, Proc. Natl. Acad. Sci. U. S. 48, 1230 (1962). 39. M. H. Malamy and B. Horecker, Biochemistry 3, 1893 (1964). 40. H. Neu and L. Heppel, BBRC 17, 215 (1964). 41. R. Simpson, B. Vallee, and G. Tait, Biochemistry 7 , 4336 (1968). 42. J. Derieux, D. Leblanc, and K. Han, Ann. Znst. Pasteur Lille 17, 65 (1966). 43. C. Lazdunski and M. Lazdunski, BBA 147, 280 (1967). 44. M. J. Schlesinger and L. Anderson, Ann. N . Y . Acad. Sci. 151, 159 (1968). 45. M. J. Schlesinger and R. Olsen, Anal. Biochem. 36, 86 (1970). 46. F. Rothman and R. Byrne, JMB 6, 330 (1963). 47. J. Reynolds and M. J. Schlesinger, Biochemistry 7 , 2080 (1968).

17. E .

379

C O l i ALKALINE PHOSPHATASE

TABLE I1 AMINOACIDANALYSIS Ref. (41)"

Ref.

Ref.

Ref.

Amino acid

(4W

(4W

(4V

Lysine Histidine Arginine Aspartic Glutamic Proline Glycine Alanine Valine Methionine Isoleucine Leucine Tyrosine Phenylalanine Serine Threonine Cysteine Tryptophan

50.3 16.4 23.1 92.5 88.4 39.4 86.0 124.4 43.0 14.4 27.7 75.4 20.3 16.0 76.0 42.0 8.0 7.9

48 19 25 91 85 40 86 120 44 13 31 72 20 16 42 71 8d 9

51 18 25 90 85 41 86 119 41 13 30 72 20 16 43 70 8d D

50 17 23 84 79 36 77 110 43 12 26 66 19 15 40 70 7 7

Ref. (47) ~

53 16 20 (174)

20

Residues per 89,000 g of protein. Residues per 86,000 g of protein. Residues per 80,000 g of protein. This value was taken from Schlesinger and Barrett ( b l ) ,who showed that there were 4 cystine and no free sulfhydryl. 4

a

( 4 S ) , represent the analysis of two different isozymes. The values of Reynolds and Schlesinger (47) are from acid-base and spectrophotometric titrations. It is also stated that there is no free N-terminal amino acid (47). The only unusual feature of the analysis seems to be the high proline content. Also, the values for serine and threonine reported by either Simpson e t al. (41) or others (43, &) are probably reversed. The differences between columns two and three are probably too small to indicate a different composition for the two isozymes. Although values for the Zn(I1) content of alkaline phosphatase generally range from 2 to 4 [Applebury and Coleman (48) have found values as high as 7, and the tetramer is believed to contain 16 ( 4 9 ) ] , there is general agreement that only two Zn(I1) are necessary for activity of the enzyme (27, 36, 60-54). 48. M. L. Applebury and J. E. Coleman, JBC 244, 308 (1969). 49. J. A. Reynolds and M. J. Schlesinger, Biochemistry 8, 4278 (1969). 50. D. Plocke and B. Vallee, Biochemistry 1, 1039 (1962).

380

T.

W.


Simpson et al. (41) also found, in addition to zinc, approximately 2 Mg, 1 Fe, and trace amounts of other metals, per mole of purified enzyme. 2. Sequence Work

The amino acid sequence around the serine that is phosphorylated in the presence of inorganic phosphate a t low p H can be seen in Table I11 (55-57). The sequence of Schwartz et al. (66) accounted for 56% of the peptides that contained 32Pi (20% or more of the peptides were excluded as extreme fractions when the peaks were pooled). The sequence, as far as it is known, is the same for alkaline phosphatase from a mammalian source ( 5 8 ) . It is interesting to note, as pointed out by Boyer and others (59-6,+),that many hydrolytic enzymes with a serine residue a t their active site have the same general sequence, i.e., Asp (Glu) Ser Ala (Gly) .

- -

C. SUBUNITS Alkaline phosphatase can be reversibly denatured by thiol reduction in the presence of urea (38),a treatment which dissociates the dimer. Proteins purified from alkaline phosphatase-negative mutants that are antigenically related to alkaline phosphatase are readily and reversibly dissociated by acid ( 6 5 ) . Normal alkaline phosphatase is more stable; but a t a lower pH, less than 3.0, it too forms monomers with release of zinc ions. However, chelating agents that remove zinc do not cause 51. R. T. Simpson and B. Vallee, Biochemistry 7,4343 (1968). 52. C. Lazdunski, C. Petitclerc, and M. Lazdunski, European J . Biochem. 8, 510 (1969). 53. J. A. Reynolds and M . Schlesinger, Biochemistry 8, 588 (1969). 54. M. Applebury, B. Johnson, and J. E. Coleman, JBC 245, 4968 (1970). 55. J. Schwartz, A. Crestfield, and F. Lipmann, Proc. Natl. Acad. Sci. U. S. 49, 722 (1963). 56. C. Milstein, BBA 67, 171 (1963). 57. N. Zwaig and C. Milstein, BJ 92, 421 (1964). 58. L. Engstrom, BBA 92, 79 (1964). 59. P. D. Boyer, Ann. Rev. Biochem. 29, 15 (1960). 60. B. S. Hartley, Brookhaven Symp. Biol. 15, 85 (1962). 61. G. H. Dixon, D. L. Kauffman, and H. N. Neurath, JBC 233, 1373 (1958). 62. H. S. Jansz, D. Brons, and M. Warringer, BBA 34, 573 (1959). 63. H. S. Jansz, C. Posthumus, and J. Cohen, BBA 33, 396 (1959). 64. F. Sanger and D. C. Shaw, Nature 187, 872 (1960). 65. M. L. Schlesinger and C. Levinthal, J M B 7, 1 (1963).

TABLE I11 AMINO

Ref. (65) (57) (58) (60) (61) (66) (63) (64) Q

ACID SEQUENCE"

8

Enzyme

Sequence

Alkaline phosphatase (E. coli) Alkaline phosphatase (E. coli) Alkaline phosphahse calf intestine Chymotrypsin Trypsin Butyrylcholinesterase Liver aliesterase Subtilisin

Thr-Gly-Lys-Pro-AspT yr-Val-Thr- AspSerP- Ala-Ma-Ser- A h (Thr or Ser) AspSerP-Ala AspSerP-Ala

Italics indicate serine a t the active site.

Ser-Gly-ValSer-Ser-Cys-MetcGly-Asp-Ser-Gly-Gly-Pro-~u AspNHrSer-Cys-Glu-Gly-Gly-Asp-Ser-Gly-Pro-Val-Asp Phe-Gly-Gl y-Glu-Ser- Ala-Gly Gly-Glu-Ser- Ma-Gly-Gl y Thr-Ser-MetrAla

$

M

382


the dimer to dissociate. Also, although the enzyme is metastable with respect to monomers a t pH 4.4, there is no exchange of 65Zn with the zinc bound to the enzyme. Other methods of forming monomers are treatment with 6 M guanidine-HC1 and heating a t 95" for 15 min (21). The properties (66) of the subunits are as follows: (1) They can be frozen and thawed several times a t pH 2.0 but cannot be stored for long periods a t -20". (2) They are stable a t 4" and p H 2.0 for several days. (3) They are unstable a t concentrations less than 10 pg/ml a t room temperature ; however, BSA will stabilize dilute solutions. (4) I n contrast to the dimer, the subunit is denatured by periodate, Pauly reagent, and ionic detergents, and is readily digested by proteolytic enzymes. (5) Subunits do not form a precipitate with antiphosphatase antibody ; however, there appear to be some antigenic determinates common to both subunits and active enzyme since subunits interfere with the precipitation of alkaline phosphatase. The alkaline phosphatase-antibody complex has 70% of the original enzymic activity as a suspension in solution. Therefore, the antibody does not bind to the active site of alkaline phosphatase, but it can still differentiate between monomers and dimers. Zinc increases the rate of dimerization (21, 65, 66) ; however, Applebury and Coleman (48) showed that zinc is not necessary for dimerization to occur. Starting at a high p H and slowly lowering the pH they found that all of the zinc is lost by the time the pH reaches 4.0,yet the molecule though inactive is still dimeric. However, upon increasing the p H of a solution of monomers, the dimer reforms by p H 5.0, yet the zinc does not bind completely until pH 6.0. Also, the optical rotatory dispersion (ORD) spectrum is the same for dimer a t p H 8.0 and monomer a t pH 4.0,but a spectral change occurs for the monomer a t pH 2.0 (48, 67). Applebury and Coleman (48) reasoned that there must be a kinetic barrier which prevents any rapid equilibration of the dimer monomer system a t intermediate pH values and that the same barrier exists in the hysteresis loop involved in the titration of carboxyl groups on the enzyme (21, 67). Rothman and Byrne (46) have used tryptic digestion to determine whether the subunits of alkaline phosphatase are identical. Since trypsin specifically cleaves a t lysyl and arginyl residues, there will be as many peptides formed as there are arginine and lysine residues if the mono66. M. J. Schlesinger, JBC 240, 4293 (1965). 67. J. A. Reynolds and M. J. Schlesinger, Biochemistry 6, 3552 (1967).

17.

E.

383


mers are completely different. If the monomers are identical, only half as many peptides will be formed. Alkaline phosphatase was labeled with 14C-lysine in one experiment and l'c-arginine in another. The number of tryptic peptides was approximately one-half the number of arginine and lysine residues (Table IV). Since their preparation of alkaline phosphatase was composed of isozymes, the authors thought that the low yield (weak) peptides were a consequence of isozyme modifications. These results suggested that the monomers are identical. Earlier data on phenylalanine-containing tryptic peptides are also consistent with identical subunits (68). Further evidence was obtained in the following way. If the subunits are nonidentical, there would be two types of tryptic peptides: type 1 which is present in only one subunit, and type 2 which is present in both subunits. A mutation producing a change in a type 1 peptide would shift the corresponding spot on the map, i.e., an old spot would disappear and a new spot would appear, while a mutation producing a change in a type 2 peptide would leave the original spot at +'z intensity and produce a new spot. Rothman and Byrne found that two peptides which were present in the largest amount, and therefore type 2 peptides, were completely missing in the double mutant C,F,,. Since the normal intensity of the spots was sufficiently strong that half the normal amount of the peptides would still be clearly visible, they concluded that the two subunits are determined by the same genetic locus (or set of loci). The conclusion drawn was that the monomers are identical. Recently, Reynolds and Schlesinger (49) reported that between p H TABLE I V NUMBEROF TRYPTIC PEPTIDES Observed Type

Strong

Lysine containing Arginine containing Ninhydrin positive Histidine containing Tyrosine containing Tryptophan containing

24 9 31

Weak 7 3 9

Total

Calca

31 12 40 7 8 3

50 23 73b-74 17 19 7

a The calculated value is from the amino acid composition and a molecular weight of 8.0 x 104. 73 if the C-terminal amino acid is lysine or arginine.

68. M. Richmond,

BJ 85, 9p (1962).

384


7.0 and 8.0, and at a Zn(I1) concentration greater than M , alkaline phosphatase forms an active tetramer. The tetramer binds 16 zinc ions and many phosphate ions. Figure 1 depicts the molecular states of alkaline phosphatase under various conditions.

D. ISOZYMES Purified preparations of alkaline phosphatase from E. coli, judged homogeneous when examined in the analytical ultracentrifuge, contain several isozymes, because several bands which contain enzymic activity are obtained in starch-gel and disc-gel electrophoresis. Although most workers find three brands (38,39, 41, 43, 69),four (44) and five (70) equally spaced bands have been found. Single gene mutations apparently affect each isozyme, for the mobility of all the bands are affected without altering the spacing (44, 67, 70). Levinthal et aZ. (70) therefore concluded that all of the isozymes were coded for by the same gene. ACTIVE

9.6s

THIOL

ACTIVE

INACTIVE

pH 2 7.0 CHELATE

pH4.0

2.0s

3.8s

FIQ.1. Molecular states of alkaline phosphatase under various conditions. 69. E. Signer, A. Torriani, and C. Levinthal, Cold Spring Harbor Symp. Quant. Biol. 26, 31 (1961). 70. C . Levinthal, A. Garen, and F. Rothman, Proc. 6th Intern. Cong. Bwchem., 1961 Vol. 1, p. 196. Pergamon Press, Oxford, 1963.

17. E. coli


385

The isozyme mobilities (44) did not change when the enzyme was ( 1 ) reversibly dissociated by low pH, 6 M guanidine hydrochloride or 6 M guanidine hydrochloride and 0.5 M mercaptoethanol; (2) reversibly inactivated with EDTA; (3) heated a t 90" for 15 min, (4) incubated at pH 12 for 60 min; and (5) incubated with metals, inorganic phosphate,

or various substrates. However, it appears that the type of isozymes that are formed depend upon the growth medium because changes in the growth medium result in different separations between bands ( 4 4 , 7 1 ) . Episomal transfer of deoxyribonucleic acid (DNA) from E. coli to S. marcescens was shown to produce E. coli type of alkaline phosphatase, S. marcescens type of alkaline phosphatase, and some new alkaline phosphatases (20, 69, 72, 7 3 ) . Further studies (38)in vivo and in vitro showed that the new enzymes produced by the cross between E. coli and S. marcescens were a dimer containing one monomer of the E. coli type and one monomer of the S . marcescens type. By crossing E. coli with an S. marcescens mutant that produced very little alkaline phosphatase, it was shown that the alkaline phosphatase isoeymes produced gave the same tryptic peptide map as the E . coli wild-type enzyme with the exception of two new peptides. Other work showed that active hybrids (38, 74) were obtained in vitro by mixing the monomers of two pseudo-revertants of E. coli that produce active enzymes which are electrophoretically different from the wild-type enzyme. Also, complimentation studies with E. coli mutants showed in vivo hybrid enzyme formation ( 7 6 ) . The fact that it is possible to form active hybrids using monomers from many different sources indicates that the isozymes from the different sources possess similar secondary structure despite their compositional and immunological differences. Lazdunski and Lazdunski (4) , separated three isoeymes on DEAEcellulose. They found that pure samples of either isozyme I (the isozyme with the least negative charge a t p H 7.0 is referred to as isozyme I) or isozyme 111,after dissociation and reassociation, gave only the original single band on disc-gel electrophoresis. Pure isozyme 11,after dissociation and reassociation, gave three bands, corresponding to isoeymes I, 11, and 111. It was later shown ( 4 4 ) , that when monomers of isoeymes I and I11 are mixed before reassociation, three bands are obtained. This 71. E. Signer, Ph.D. Thesis, M.I.T., Cambridge, Massachusetts, 1961. 72. C. Levinthal, E. Signer, and A. Torriani, Bull. N . Y . Acad. Med. [21 38, 365 (1962). 73. M. J. Schlesinger and C. Levinthal, JMB 7 , 1 (1963). 74. D. P. Fan, M. J. Schlesinger, A. Torriani, K. Barrett, and C. Levinthal, JMB 15, 32 (19s6). 75. A. Garen and S. Garen, JMB 7 , 13 (1963).

386


shows that isozyme I1 is composed of one monomer of isozyme I and one monomer of isozyme 111. Schlesinger and Anderson (44) separated the isozymes on starch-gel and DEAE-cellulose a t various times after 14C-amino acids were added to a culture of E . coli synthesizing alkaline phosphatase. They found that initially most of the counts appeared in isozyme I, but later they appeared mainly in isozymes I1 and 111. In another experiment where only enough radioactive amino acid was added to allow for 5 min of synthesis of radioactive protein, they found that the label is initially found in isozymes I and I1 and later in isozymes I1 and 111.These experiments establish that the monomers of isozyme I are precursors for isozymes I1 and 111. In another set of experiments, Schlesinger and Anderson (44) showed that the isozymes are formed in vivo by alteration of the dimer. Using an E . coli mutant that makes an altered subunit, which will only dimerize (in vitro or in v i m ) in the presence of phosphate or Einc, they found that the monomers produced by the cell growing in the absence of phosphate and zinc produced only one electrophoretic form when the monomer was converted to the dimer in vitro. However, if the medium is made 2 mM in phosphate and 10 y-M in zinc, in the exponential phase of growth, three isozymes are formed. Additional support for the conclusion that isozymes are made by alteration of the dimer comes from the fact that independent of when 14C-labeled amino acids are added to the growing culture, label appears first in isozyme I. It thus appears that a mechanism is available in the periplasmic space for the conversion of isozyme I to the other isozymes. Tryptic peptide fingerprint studies (44) of the pure isozymes I and I11 showed a ninhydrin-positive peptide in isozyme I that is missing in isozyme I11 and two weakly ninhydrin-positive spots present in isozyme I11 but not in isozyme I. The two peptides sequences present only in isozyme I11 are more negatively charged than the one present in isozyme I. Since there are 36 tryptic peptides, it appears that the structural differences between the isozymes are not extensive. Other studies on the pure isozymes (43, 4.4) showed that K,,, and V,,, for p-nitrophenyl phosphate and Ki for inorganic phosphate are the same for the different isozymes. The isozymes also have the same ORD spectrum and pH vs. pKt profile. The isozymes showed only slight differences in their rates of dissociation and reassociation. It has been suggested (4.4, 71) that the differences in the isoayme composition involve carbohydrate residues. Salanito (76) found that 78. J. Salanito, Ph.D. Thesis, University of Indiana, Bloomington, Indiana, 1968.

17. E .

387

CO2i ALKALINE PHOSPHATASE

R

co

I SOZY M E

ISOZYME

I

II

CELL MEMBRANE

1 ISOZYME

X

= CARBOHYORATE (7) n.? FIG.2. Possible scheme for isozyme formation.

ISOZYME

m

reaction of the isozymes with periodate leads to the formation of one species. Figure 2 represents a possible scheme for isozyme formation,

E. PHYSICAL PROPERTIES As seen in Table V, the molecular weight values that have been obtained for alkaline phosphatase vary over a wide range ( 4 , 39, 41, 47, 48, 65, 7 7 ) . A more complete table of values, determined by the equi-

TABLE V PHYSICAL PROPERTIES OF ALKALINE PHOSPHATASE Ref. Sedimentation coefficient Partial specific volume (calc) Intrinsic viscosity Electrophoretic mobility Isoelectric point Isoionic point Frictional ratio MWw MWZ MWzit Absorbance a t 278 nm for 1 mg/ml

s : ~ , ~= 6 . 0 a t

pH 8 . 0 0.73 ml/g 3 . 4 cma/g 3.3 X cm*/V sec at pH 7 . 6 pH 4 . 5 pH 6 . 3 1.05 (67-98) X 108 at pH 8.0 (88-99) X l@a t pH 8 . 0 (86-110) X loa a t pH 8 . 0 0.72

(48)

77) (63) (4)

(41

(4) (47)

(4) (48) (48) (48) (391 411

77. A. Ullman, M . Goldberg, D. Perrin, and J. Monod, Biochemistq 7, 261 (1968).

388


librium centrifugation method of Yphantis (78), was given by Applebury and Coleman (48) for various conditions of pH and ionic strength. They concluded that the obvious heterogeneity of the alkaline phosphatase system could be explained by a system which is either nonideal or paucidisperse or both. Possibly the formation of tetramers could be involved.

6.0-

(A 1

4.0 o

Schlieren Scanner

2.0 I

P

0 X

IZ FIQ.3. Sedimentation coefficient and molecular weight as functions of pH. (A) Sedimentation coefficients as a function of pH. (0) si,, values were determined for samples adjusted from neutral pH to each pH value. The concentrations were 7.2 mg/ml, except at pH 5 and pH 4, where they were 4.0 mg/ml in 0.01 M tris-0.01 M sodium acetate. ( 0 )s1",,, values were determined for samples adjusted from neutral pH to the given pH. Concentrations were 0.62 mg/ml in 0.1 M NaCI-0.01 M tris-O.01 M sodium acetate. s ~ ~ , values , were determined for samples prepared a t pH 2, then dialyzed a t the appropriate pH. Concentrations Determinawere 0.62 mg/ml in 0.1 M NaCl-O.01 M tris-O.O1 M sodium acetate. (0) tions with Schlieren optics; all other determinations were made with the use of ultraviolet optics with the photo&ectric scanner. (B) Weight average molecular weight as a function of pH. ( 0 )M, values were determined for samples adjusted from neutral pH to each pH indicated, by dialysis, 0.62 mg/ml, in 0.1 M NaCl-0.01 hf tris-O.O1 M sodium acetate. (0) values were determined for samples prepared at pH 2, then dialyzed at the appropriate pH, 0.62 mg/ml in 0.1 M NaCl-O.01 M tris-O.O1 M sodium acetate.

(m)

a,

78. D. A. Yphantis, Biochemistry 3, 297 (1964).

17. E .


389

Reynolds and Schlesinger (63,67) have determined values of the intrinsic viscosity at different temperatures and pH. These data show that the enzyme is compact down to a pH of 4.0 but appears to be a random coil at pH 2.0 and also in 6 M guanidine hydrochloride. This is in agreement with results obtained from circular dichroism and optical rotatory dispersion studies (48, 63). Several workers (4, 21, 27, 39, 41, 46, 48, 53, 7 7 ) have determined values for the sedimentation coefficient. The pH dependence can be seen in Fig. 3 (4.8). In electron micrographs, alkaline phosphatase appears as a compact sphere with a diameter of about 60 A (4, 6 6 ) . This is in agreement with the value of 57 A calculated for a spherical unhydrated protein based on the frictional ratio and a molecular weight of 80,000 ( 4 ) . Electron micrographs of the acid-prepared monomer show no discernible structure (66).

F. CRYST~LL STRUCTURE Recently, Applebury et al. (64) have obtained large single crystals of E . coli alkaline phosphatase. Initial X-ray studies by Hanson et al. [see ( 5 4 ) ] show the crystals to be of the space group P3121, each unit cell containing three dimers. The unit cell dimensions are a = 70.5 A, b = 70.5 A, c = 155.6 A, j3 = 120°, which is consistent with the globular form predicted by the hydrodynamic frictional ratio, The threefold screw axis is along the c axis; the twofold axes of rotation relate pairs of identical monomers. Assuming that the twofold rotational axis between monomer units of the protein observed in the crystal is a twofold axis relating the two subunits of a functional dimer, unique single sites per dimer would be present only along the twofold axis (54).

G. CHEMICAL MODIFICATION 1. Modification during Protein Synthesis

a. Phenylalanine Replacement. With a phenylalanine auxotroph of E . coli, p-fluoro phenylalanine, m-fluoro phenylalanine, P-2-thienylalanine, and P-3-thienylalanine were substituted for phenylalanine in alkaline phosphatase (79, 80). The various alkaline phosphatases obtained using the different precursors showed the same specific activity to p 79. R. L. Munier, Compt. Rend. 250, 3524 (1960). 80. R. L. Munier and G . Sarrozin, Compt. Rend. 254,2853 (1962).

390

T.

W.


nitrophenyl phosphate. Later studies with wild-type E . coli (68, 81-83), showed that p-fluorophenylalanine could compete with phenylalanine. The resulting alkaline phosphatase contained 56% p-fluorophenylalanine randomly distributed in place of phenylalanine. The altered enzyme had the same sedimentation coefficient, starch-gel electrophoretic mobility, and specific activity as the normal enzyme. b. Leucine Replacement. Adding norleucine in place of leucine to the medium in which E . coli is grown results in the synthesis of an altered alkaline phosphatase which has a low specific activity but reacts normally with anti-alkaline phosphatase serum (84).

c. Tryptophan Replacement. Using an E. coli tryptophan auxotroph is was found that 5-fluorotryptophan (85),6-fluorotryptophan (85), azatryptophan (86), and tryptazan (86) substituted for tryptophan in the medium had little effect on the structure and function of the alkaline phosphatase that was produced. d. Histidine Replacement. Substitution for histidine by triazolealanine (87) or 2-methyl histidine (88)in the medium of an E . coli histidine auxotroph resulted in the formation of subunits that were incapable of dimerization. The monomers were detected by antisubunit antibody. e. Arginine Replacement. The substitution of canavanine in place of arginine, for the growth of an arginine auxotroph of E. coli, resulted in the formation of subunits incapable of dimerization and active dimers (89).The dimers showed a lesser stability and a lower specific activity than the normal enzyme. It was thought that the active dimers might have resulted from the presence of arginine in the protein (in one or more regions of the subunit necessary for dimerization) derived from arginine pools that were not removed before canavanine was added.

f. Methionine and Cystine Replacement. Under conditions of sulfur deprivation (0.06 mM sulfate), addition of 3 mM NazSeOs to the growth medium of E . coli results in the formation of alkaline phosphatase with a specific activity that is 30% that of the normal enzyme. The selenium 81. M. Richmond, BJ 84, llOp (1962). 82. M.Richmond, BJ 85, 9p (1962). 83. M.Richmond, J M B 6, 284 (1963). 84. S. Neale and H. Tristram, BBRC 11, 346 (1963). 85. R.L. Munier and A. Drappier, Compt. Rend. D265, 1429 (1967). 86. S. Schlesinger, JBC 243, 3877 (1968). 87. S.Schlesinger and M. Schlesinger, JBC 242, 3369 (1967). 88. S. Schlesinger and M. Schlesinger, JBC 244, 3803 (1969). 89. J. Attias, M.Schlesinger, and S. Schlesinger, JBC 244, 3810 (1969).

17. E .


39 1

incorporation was S-lOPr, of the number of methionine and cystine residues (90). 2. Modification after Protein Synthesis Photooxidation of alkaline phosphatase in the presence of methylene blue and Rose Bengal causes loss of activity for both native and apoenzyme. I n the case of the native enzyme, zinc protects 2 to 3 of the 16 histidine residues. The rate of oxidation of tryptophan is not affected by zinc, and there was no loss of tyrosine. Also, photooxidation of the apoenzyme diminishes zinc binding. It would appear that histidine residues play a role in binding the two zinc ions necessary for enzymic activity (91). Treatment of the enzyme with N-bromosuccinimide oxidized 2 of the 8 tryptophan residues and 8 of the 20 tyrosine residues, but none of the histidine residues. This treatment causes the phosphotransferase activity with tris as an acceptor to double and the hydrolase activity to increase slightly. I n the case of cobalt alkaline phosphatase, the above treatment caused a threefold increase in hydrolase activity and the generation of an even greater phosphotransferase activity (91). Reaction of a 2000-fold molar excess of 5-diazonium-1H-tetrazole (DHT) with alkaline phosphatase, reduces hydrolase activity to 36% and phosphotransferase activity to 29% of the control. Since no color was produced by the reaction it was assumed that the reaction occurred with lysine residues. When the concentration of DHT is a 20,000-fold molar excess, color develops and the hydrolase activity falls to 11% while the phosphotransferase activity falls to 47%. I n this later DHT reaction, the spectral maxima produced are characteristic of azotyrosine and azohistidine (91). Acetic anhydride causes the same decrease in activity as DHT by acetylation of €-amino groups of lysine; no acetylation of tyrosine occurred (91). No loss of activity is detected upon incubation of alkaline phosphatase with 0.02 M iodoacetamide or iodoacetate a t 25" for 2-12 hr over the pH range of 4-7.5 (92). Recently, it was reported (93) that alkaline phosphatase can be covalently coupled to porous glass with a silane coupling agent. The 90. 91. 92. 93.

G. Ahluwalia and H. Williams, ABB 117, 192 (1966). G. Tait and B. Valee, Proc. Natl. Acad. Sci. U.S. 56, 1247 (1966). S. Plotch and A. Lukton, BBA 99, 181 (1965). H. H. Weetall, Nature 223, 959 (1969).

392


enzyme appears to behave in a normal manner when assayed with p-nitrophenyl phosphate a t various pH.


A. SPECIFICITY The equilibrium catalyzed by alkaline phosphatase is classically of the following general type: 0

II

R4-P-OH

I

0

+ H20

ROH

+ H O - PI14 H I

OH

OH

Since the equilibrium lies well to the right it is customary to say that alkaline phosphatase hydrolyzes phosphate esters, but some related compounds are also hydrolyzed (Table VI) (3, 4, 28, SO, 94-100). The enzyme also catalyzes transphosphorylation reactions in which a different alcohol substitutes for H,O as a phosphate acceptor. Compounds that are hydrolyzed have the general structure, 0

II

X-P-OH

I

OH

where X can be RO-, HO-,RS-, HS-, 0

I1

RO-P-0-

I

OH

and F-, but P-N and P-C bonds are not cleaved. The enzyme has no diesterase activity. A distinctive feature of the alkaline phosphatase-catalyzed hydrolysis is that the relative rates of hydrolysis of the many different phosphate and S-phosphorothioate esters are nearly the same (it is difficult t o get precise rate values because of product inhibition by phosphate, the K , 94. C. Lazdunski and M. Lazdunski, European J . Bwchem. 7,294 (1969). 95. S. Horiuchi, Japan. J . M e d . Sci. B i d . 12, 429 (1959). 96. M. Gottesman, R. Simpson, and B. Vallee, Biochemistry 8, 3776 (1969). 97. D. Harkness and R. Hilmoe, BBRC 9, 393 (1962). 98. D. Trentham and H. Gutfreund, BJ 106, 455 (1966). 99. H. N. Fernley and P. G. Walker, Nature 212, 1435 (1966). 100. H. Neumann, L. Boross, and E.Katchalski, JBC 242, 3142 (1967).

17. E .

393

CO2i ALKALINE PHOSPHATASE

TABLE V I SUBSTRATES OF ALKALINEPHOSPHATASE Compound

Relative rate

Ref.

Compound

Relative rate

Ref.

5'-AMP

Polymetaphosphate

0.9

(28)

3'-AMP

Ribose 5-phosphate

0.7

(988)

2'-AMP

8-Glycerol phosphate

0.9-1.0

(3,4, 28, 9.6, 96)

ATP dATP

Ethanolamine phosphate Glucose 1-phosphate

0.6-0.9

(S,4, 28,

dAMP dGTP dGMP

Glucose 6-phosphate Histidinol phosphate p-Nitrophenyl phosphate

0.9 0.8-0.9 1.0

94) (3, 28) (4, 28) (3, 4, 28, 94-97)

2'- and 3'GMP 5'-GMP

Riboflavin 5'-phosphate

0.7

(4)

o-Carboxyphenyl phosphate a-Naphthyl phosphate 8-Naphthyl phosphate

0.25

(94)

1.5 1.0

(94) (30)

2, 4-Dinitrophenyl phosphate Fructose lI6-diphosphate Phosphoenol pyruvate 4-methyl urnbellif eryl phosphate N-Acetylcysteamine Sphosphate S-(Carboxymethyl) phosphorothioate S-[2-(Methoxy carbonyl) ethyl] phosphorothioate Cysteamine S-phosphate

1.0

(96, 98)

0.9-1.0

(4, 96)

dCMP 2'- and 3'CMP 5'-CMP

0.8-1.2

(4, 96)

dCTP UDP 5'-UMP

1.05 1.o 0.8-1.3

($8)

0.9 1.1

(96) (96)

(28)

(4, 28,

2'- and 3'-

1.0

96) (96)

UMP dTTP

1.0

(288)

5'-TMP

0.9

(96)

5'-IMP APAP POlY c POlY 1 PPi PPPi

0.7

(98)

0.6 0.42 1.0

(99)

0.7

(100)

0.7

(100)

0.7

(100)

0.73

(100)

(3)

for many substrates is higher than the Ki for phosphate). The only exceptions are o-carboxyphenyl phosphate and phosphoenol pyruvate. Bamann and Schwarze (101) have shown that mammalian alkaline phosphatase, which is also nonspecific, preferentially catalyzes the hy101. E. Bamann and P. Sohwarze, 2.Physiol. Chem. 349, 192 (1968).

394

T. W. REID AND

I.

B. WILSON

drolysis of the L ( + ) isomer of phosphomandelic acid. One can speculate that there is a positively charged group (perhaps zinc ion) in the active site of alkaline phosphatase, which normally interacts with the phosphate group of substrates but which interacts instead with the negatively charged carboxyl group on compounds such as L(-) -phosphomandelic acid, o-carboxyphenyl phosphate, and phosphoenol pyruvate to produce a less reactive enzymesubstrate complex. Phosphoserine is hydrolyzed (102) at the normal rate by alkaline phosphatase; however, the carboxyl group in this compound is more distant from the phosphate group. Other compounds, not shown in Table VI, which are hydrolyzed by alkaline phosphatase are: T P N (28), poly A (28), phosphocellulose (28), pyrophosphoserine (102, 10S), phosphoserine (102-104), pyridoxine phosphate (104), pyridoxal phosphate ( l o $ ) , phosphothreonine (104), and phosphocholine (104). These compounds are all hydrolyzed a t approximately the same rate.

B. COMPETITIVE INHIBITORS There are few potent competitive inhibitors for alkaline phosphatase (Table VII) (4, 21, 94, 96, 100, 105-109). Phosphate, thiophosphate, and arsenate have low values for Ki. These substances are actually substrates and can form a covalent intermediate. However, this is probably not the reason why they are potent inhibitors since in the case of phosphate the Michaelis complex is more stable than the covalent intermediate (SO).The values of Ki for phosphate and arsenate a t various pH values have been published (SO,106, 110). 0-p-Nitrophenyl phosphorothioate is a t best a poor substrate. Breslow and Katz (111) gave a V,,, for 0-p-nitrophenyl phosphorothioate (pH 8.0, 1.0 M NaCl, 25"C), which is 100 times smaller than for p 102. S. M. Avaeva, S. N. Kara-Murza, G. L. Kogan, N. V. Raskova, and M. M. Botvinik, Dokl. Akad. Nauk SSSR 172, 1436 (1967). 103. S. Avaeva, S. Kara-Murza, N. Raskova, and M. Botvinik, Khim. Prirodn. Soedin. 3, 328 (1967); C A 68, 1049p (1968). 104. N. Okada, Osaka Daigaku Zgaku Zasshi 15, 211 (1963); C A 60, 1992f (1964). 105. I. B. Wilson and J. Dayan, Biochemistry 4, 645 (1965). 106. C.Lazdunski and M. Lazdunski, BBA 113, 551 (1966). 107. S. E. Halford, N. G. Bennett, D. R. Trentham, and H. Gutfreund, BJ 114, 243 (1969). 108. F.Eckstein and H. Sternback, BBA 146, 618 (1967). 109. H.Neumann, JBC 243, 4671 (1968). 110. H.N.Fernley and P. G . Walker, BJ 111, 187 (1969). 111. R.Breslow and I. Katz, JACS 90, 7376 (1968).

TABLE V I I COMPETITIVE INHIBITORS OF ALKALINE PHOSPHATASE Ki X 106 Compound Orthophosphate

(M)

Buffer

5.6 2.5 10

1.OM tris O.1M tris 0.01 M tiis 0.01 M tris 0 . 1 M tris 0.02 M Barbital 0.01M tris 1.OMtris 0.01 M tris 0.01 M tris 0.01 M tris 1.0 M tris 1.0 M t,ris 0.01 M tris 0.01 M tris 0 . 3 M tris 0.3 M tris 0.02 M Barbital

0.8

Thiophosphate Arsenate Phenyl phosphonate pNitrobenzy1 phosphonate 2-Hydroxy-5-nitrobemyl phosphonate Phenyl phosphonate pChloroanilidophosphonate Uridine 5'-0-phosphorothioate Thymidine 5'-0-phosphorotbioate pNitropheny1 phosphorothioate

0.6 17 2.5 5.6 20.0 950 5000 100 220 2800 40 120 92 < O . 01

Conditions pH 8.0 pH8.0 pH8.0 pH8.0 pH 8.0 pH9.0 pH8.0 pH 8.0 pH8.0 pH8.0 pH 8.0 pH 8.0 pH 8.0 pH7.5 pH8.5 pH 8.0 pH 8.0 pH8.7

1MNaCl 1MNaCl 0.4MNaCl 1.5MNaCl 0.4MNaCl 1.OMNaCl 1.OMNaCl

0.4MNaCl 0.4MNaCl

1.5MNaCl

Ref. 25" 25" 25" 25" 25" 25" 45" 25" 25" 25" 25" 25" 25" 45" 25" 25" 25" 25"

(4, 31) (105) (96)

(106) (106) (100) (106)

E

3 3 i!

2 M

(4) (96)

(106) (96) (107) (107) (106) (94)

(108) (108) (109)

E

396


nitrophenyl phosphate. I n contrast, they found that nonenzyme hydrolysis of 0-p-nitrophenyl phosphorothioate is much more rapid than hydrolysis of p-nitrophenyl phosphate. The enzyme hydrolysis of the phosphate and thiophosphate esters follows the same pattern as nonenzymic hydrolysis of triesters, where the hydrolysis of the thiophosphate ester is slower. Triester hydrolysis is presumed to follow an addition-elimination mechanism, in which the P=O (P=S) bond order decreases in the transition state and the oxygen (sulfur) increases in charge, while monoesters are presumed to use an elimination (to metaphosphate) -addition sequence, in which the P=O (P=S) bond order increases in the transition state and the charge on oxygen (sulfur) diminishes. These effects are thought to reflect the lesser electronegativity of sulfur compared with oxygen. Breslow and K a ts suggested that the enzymic hydrolysis of monoesters resembles the alkaline hydrolysis of triesters and points strongly to an addition-elimination sequence for alkaline phosphatase. I n contrast to the above, Neumann ( l o g ) , using an enzyme solution (pH 8.7, 0.1 M tris, 25°C) 3000 times more concentrated than that of Breslow and Katz, did not detect hydrolysis of 0-p-nitrophenyl phosphorothioate after 2 hr. She also reported the isolation of an extremely stable enzyme-nitrophenyl phosphorothioate complex. One might speculate that in the light of the extreme stability of the proposed enzymephosphorothioate complex, the complex may actually be a covalent species

sl

(M- -OH)

I

OH

which does not hydrolyze.

C. THEPHOSPHATE ENZYME Agren (112) and Engstrom (113) isolated serine phosphate from mammalian alkaline phosphatase that had been incubated with inorganic phosphate in acid p H ( < 6 ) . Engstrom (114) and Schwartz and Lipmann (35) later obtained similar results with E . coli alkaline phosphatase. They found that a large percentage of the enzyme is phosphorylated, that compounds like glucose 6-phosphate and sodium arsenate inhibit 112. G. Agren, 0. Zetterqvist, and M. Orjamae, Acta Chem. Scand. 13, 1047 (1959). 113. L. Engstrom and G. Agren, Acta Chem. Scand. 12, 357 (1958). 114. L. Engstrom, BBA 58, 606 (1962).

17. E. coli

397


phosphate incorporation, and that phosphorylation did not occur with enzyme that was denatured by heating. Engstrom (115) also showed that the enzyme could be labeled by rapidly quenching the enzyme substrate solution during its hydrolysis of glucose 6-phosphate ("P) . In a study of the amount of labeling vs. pH, Schwartz (116) showed that labeling is a maximum at about pH 4.0 and is essentially zero a t pH 7.0 and higher. This is in contrast to the hydrolytic activity of the enzyme, which is maximum above pH 8, 2% as great a t pH 6.0, and much less a t lower pH. He also found that the rate of exchange of '*O into inorganic phosphate is higher a t lower pH. As an explanation of the phosphoryltransferase activity of mammalian phosphatases, Morton (117) had earlier advanced the idea that phosphate ester hydrolysis catalyzed by the nonspecific phosphatases occurs in two catalytic steps similar to that shown by Wilson et al. (118) for cholinesterase. The first step is the formation of a phosphoryl enzyme with the splitting out of the alcohol group. The second step is the hydrolysis of the phosphoryl enzyme. While it might appear that the phosphoprotein obtained in the above labeling experiments confirms the Morton hypothesis, the p H relationships require some explanation, and various other considerations such as the potent inhibitory properties of phosphate a t p H 8.0, its poor inhibitory properties a t pH 5.0, the extreme thermodynamic stability of the phosphoprotein and other thermodynamic considerations require further demonstration of a phosphoryl enzyme intermediate (116, 119, 120).

I n order to test for the existence of a phosphoryl enzyme intermediate, Barrett et al. (121) carried out an experiment whose method is illustrated in Scheme I

/ E

+

Tris-P

+

ROH

1

lH,o E

+

Pi

+

ROH

trisl

\;.

Tris-P+ E

E

+ P,

115. L. Engstrom, Arkiv Kemi 19, 129 (1962). 116. J. H. Schwartz, Proc. Null. Acad. Sci. U . S. 49, 871 (1963). 117. R. K. Morton, Discussions Faraday SOC.20, 149 (1955). 118. I. B. Wilson, F. Bergmann, and D. Nachmansohn, JBC 186, 781 (1950). 119. C. Milstein, BJ 92, 410 (1964). 120. M. M. Pigretti and C. Milstein, BJ 94, 106 (1965). 121. H. Barrett, R. Butler, and I. B. Wilson, Biochemistry 8, 1042 (1969).

398


Scheme I shows the hydrolysis of a phosphate ester in the presence of tris, which can serve as a phosphate acceptor so that O-phosphoryltris is a product as well as Pi. It has been shown that in the presence of alcohols such as tris and ethanolamine the rate of substrate utilization is increased, that formation of alcohol exceeds that of phosphate, and that the difference is due to the formation of the O-phosphorylamino alcohol (122, 123). The question was: Does the reaction with water and with tris emanate from the Michaelis complex or from a phosphoryl enzyme intermediate (E-P)? If the reactions with tris and water stem from a phosphoryl enzyme, the ratio of products tris-phosphate and Pi would be independent of the leaving group RO, but if the reactions stem from the reversible complex containing the leaving group, the ratio of products would depend upon the structure of R. It was found that the ratio of free alcohol to phosphate was 2.39 f.0.02 for nine different substrates, including esters such as p-cresyl phosphate P-naphthyl phosphate, and phosphoenol pyruvate. This experiment established the occurrence of a phosphoryl enzyme intermediate. Similarly, Neumann (124) found the same percent transphosphorylation with a given acceptor and three substrates. Several acceptors were used. These results also prove the existence of a phosphoryl enzyme. With the establishment of the phosphoryl enzyme, the question was whether or not the phosphoryl enzyme was the same as the phosphoprotein found by incubating inorganic phosphate with alkaline phosphatase a t low pH (35, 114-116, 119, 120). Wilson and Dayan (105) pointed out that the phosphoprotein is thermodynamically very stable: I t is lo5 times more stable than O-phosphorylserine (125) and 0phosphoryl ethanolamine (105, 126). Alkaline phosphatase, as a true catalyst, must catalyze both the hydrolysis and the formation of phosphate esters. Therefore, if a serine residue existed which was capable of forming a thermodynamically stable phosphate ester, alkaline phosphatase as a nonspecific catalyst would catalyze its formation from both inorganic phosphate and phosphoester substrates. I n order to see if the phosphoryl enzyme is thermodynamically stable as compared to ordinary phosphate esters, Levine e t al. (30) carried out kinetic experiments which yielded information concerning the equilibria between Pi and alkaline phosphatase (E) (127). 122. J. Dayan and I. B. Wilson, BBA 81, 620 (1964). 123. I. B. Wilson, J. Dayan, and K. Cyr, JBC 239, 4182 (1964). 124. H.Neumann, European J . Biochem. 8, 164 (1969). 125. G. E. Vladimirova, A. I. Komkova, and N. A. Fedorova, Biokhimiya 26, 426 (1961); C A 55, 22427f (1961). 126. J. Dayan and I. B. Wilson, BBA 77, 446 (1963).

17. E . C d i

399


E

+ Pi=k-iki E *Pi Sk-2k t E-P + Hz0

The equilibrium constant for dissociation of the Michaelis complex, E . P i , i.e., the complex that does not involve the covalent bond is

and the equilibrium constant for the hydrolysis of the phosphoryl enzyme, E-P, to yield enzyme and Pi is

where k-, is defined to contain the concentration of water. Similarly,

at equilibrium. Phosphate competitively inhibits the hydrolysis of phosphate esters. The appropriate reaction is shown in (128). The steady state solution has the Michaelis-Menten form for competitive inhibition

where

127. Many papers use the older numbering system for rate constants, as used by Michaelis, but the newer system is now more prevalent and is used in this review. 128. Primed rate constants (k’) are used in schemes for substrate hydrolysis, and unprimed rate constants (k) in schemes for phosphate binding.

0 E

ki’

+ Sk’-iS E - S - 1 Ekr’

S

=

I

II I

-OH

R W P 4 H OH

+ ROH

400


K, =

The values obtained for the various constants (see Table VIII) show that the phosphoryl enzyme is thermodynamically stable, ie., KE-p is small. The same quantities that were evaluated by kinetic measurements were also evaluated by labeling measurements using 32P inorganic phosphate [Reid et al. (SI)].There is no labeling a t pH 8.0 and very little labeling a t pH 7.0, but enough to make the measurements. From Eqs. (1)-(3) for the equilibrium between inorganic phosphate and alkaline phosphatase, one can obtain

Using the equation for Ki [Eq. (7) 1, one can transform Eq. (8) to explicitly contain Ki.

A plot of (E")/(E-P) vs. l / ( P i ) should yield a straight line whose slope is KE-p and whose intercept is KE+/Ki. These results are seen in Table VIII, where the results from the two different approaches compare quite well, indicating that the phosphoryl enzyme and the phosphoprotein are the same. These data also show that the reason that little phosphoryl enzyme is detected by phosphate labeling a t pH 7.0 and above is not because the phosphoryl enzyme is unstable TABLE VIII VALUESFOR VARIOUSCONSTANTS

8.P

7.0 a 6.0 a 5.5 a 4

(1.5 X 1.4 X (2.1 X 6.0 X (1.4 x 1.2 x (1.8 x

lo-') lo-' lo-' lo-&) 10-6 10-6)

(2.5 X 3.8 X (2.3 X 3.1 X ( 5 . 1 x 10V) 1 . 0 x 10-5 (1.6 x 10-5)

Kinetically determined values.

( 2 . 5 X lo+) 4.1 X (2.3X 6.7 X (7.6 x 5 . 4 x 10-6 (1.1 x 10-4)

( 1 . 7 X 1W2) 2 . 9 X lo-* (1.2 X 1.1 (0.7) 4.4 (6.6)

(1.7) 2.7 (1.2) 51 (67) 81 (85)

17. E . Coli


401

a t high p H but because the Michaelis complex is even more stable. This is indicated by a low value of lc+/k2.

D. THEROLEOF ZINC There has been some uncertainty concerning the metal content of alkaline phosphatase and the role of zinc in the catalytic process. Early measurements by Plocke et al. (36, 50) showed that there were 2 gatoms per dimer. The zinc requirement for enzymic activity was demonstrated by the inhibition of the enzyme with metal binding agents in accord with the order of the stability constants of their zinc complexes. It appears that in some cases (EDTA) zinc is removed from the enzyme and in other cases (CN) the ligand adds to the metalloprotein. A zincfree inactive apoenzyme was formed by dialysis against 1,lO-phenanthroline. Complete activity was restored by zinc; only zinc, cobalt, and possibly mercury produce active enzyme. Several investigators now find that four zinc ions are bound by the dimer but only two are necessary for activity. Lazdunski et al. (62) showed that the rate of inactivation of the enzyme by EDTA is biphasic, corresponding to two different zinc binding sites associated with enzymic activity. Phosphate decreases the rate of inhibition by EDTA in a manner corresponding to the binding of phosphate with dissociation constants for the second. They of 1 X lo-&for the first zinc removal and 6 X propose that there are four zinc binding sites, of which the strongest and weakest are required for activity. If one site is occupied by zinc and three by C d ( I I ) , there is 11% activity. They concluded that the two essential zinc sites are the same as measured by Cohen and Wilson (see later). Simpson and Vallee (61) found that when alkaline phosphatase is exposed to 8-hydroxyquinoline-5-sulfonic acid, two zincs are rapidly removed and the enzyme is inactivated to within 10%. The two remaining zincs are removed more slowly, presumably with the loss of the remaining activity. When zinc is added to the apoenzyme, the first two ions produce 85% activity. Thus it would appear that there are two binding sites that must be occupied by zinc ions for activity (51). Two classes of binding sites are also indicated by studies with the cobalt enzyme. The first two cobalt ions bound by the apoprotein do not produce an active enzyme. I n the absence of evidence to the contrary, it is assumed that these two sites are the same as the “unnecessary” zinc sites. The absorption spectrum of the two-cobalt enzyme is similar

402


to the spectrum of octahedral cobalt complexes. Addition of two more cobalt ions generates enzymic activity and a more complex spectrum develops containing four maxima in the visible wavelength range which corresponds to neither octahedral nor tetrahedral spectra. The spectrum suggests an unusual coordination environment ( 5 1 ) . Applebury and Coleman (48) found that biosynthetic enzyme and apoenzyme labeled with 65Zn bound 2-3 zinc ions per dimer at neutral pH but as many as 7 a t pH 10.0. Dialysis for 24 hr removes “extra” zinc, and after 20 days only two remain and the enzyme is active. Thus it appears that the two zinc binding sites that are most readily occupied in the apoenzyme and must be occupied for activity are the ones that most strongly retain zinc ions during dialysis but, on the other hand, most readily lose zinc ions by reaction with 8-hydroxyquinoline-5sulfonic acid. I n other studies (129) Applebury and Coleman found that the Co(I1) enzyme is active and has a multibanded visible absorption spectrum indicating an unusual and probably distorted geometry. They showed that the absorption bands reach maximum intensity a t 2.2 Co(I1) per dimer (compare Simpson and Vallee). Phosphate induces major changes in the magnitude of the oscillator strengths and optical activity of the visible absorption bands but little change in their energy. Thus phosphate appears t o produce a major change in the dissymmetry of the local environment of the two cobalt ions but little change in the d-orbital splitting. These workers showed (54) that Mn (11) , Co (II), Zn (II), and Cd (11) induce binding of phosphate to alkaline phosphatase, while N i ( I I ) , C u ( I I ) , and Hg(I1) do not. Cohen and Wilson (130) measured the activity of the enzyme as a function of pZn over a broad range of zinc ion concentration using zinc ion buffers. The curve indicated that the activity depends upon two ionizations with constants and 10-10.22in 1 M NaCl a t pH 8.0, 25°C. The activity of the enzyme after the first ionization of zinc ion was 12% of the original. Using equilibrium dialysis with 65Zn(II) and 1,lO-phenanthroline, Csopak (131) found that the first two zinc ions bound to the apoenzyme had a dissociation constant of in 0.1 M tris a t pH 8.5 and 25°C. At the time Cohen and Wilson did their work, the enzyme was believed to contain only two zinc ions (36, 5 0 ) . In the light of later developments described above, it would appear that the two ionizations might refer to pairs of zinc ions. Reynolds and Schlesinger (53) differed from others in their finding 129. M. L. Applebury and J. E. Coleman, JBC 244, 709 (1969). 130. S. R. Cohen and I. B. Wilson, Biochemistry 5, 904 (1966) 131. A. Csopak, European J . Biochem. 7, 186 (1969).

17. E . coli


403

that the enzymic activity of alkaline phosphatase increases linearly with the number of zinc ions bound up to four. Spectrophotometric titration shows that approximately 6 tyrosine residues are not exposed to solvent in the case of the dimer containing three Zn(I1). They also found (49) that a t pH values between 7.0 and 8.0 and Zn(1I) concentration > M , alkaline phosphatase rapidly and reversibly forms a tetramer, as shown by osmotic pressure and sedimentation studies. At pH 8.0 and an equilibrium concentration of Zn(I1) = M , they found 16 Zn(II)/ tetramer. In summary, all workers except Reynolds and Schlesinger (63) found that only two zinc ions are important for activity. However, Lazdunski et al. (62) found that the first and last of four zincs to bind to the apoenzyme are important for activity, Simpson and Vallee (61) found that the first two of four !zincs and the last two of four cobalts are important for activity, while Applebury and Coleman (48) found that the first two zincs and the first two cobalts are necessary. Cohen and Wilson (130) found two different dissociation constants for the essential zincs, and Csopak (131) reported one dissociation constant for binding two zincs. Some of the disagreement in the metal binding work may arise from uncertainties as to whether binding was controlled by kinetics or thermodynamics. To illustrate what is meant, consider the addition of one zinc ion to apoenzyme. Is the site where this zinc ion is bound the most stable site or the one most accessible? Evidently this might depend upon the time, and whether or not the concentration of zinc is controlled by a chelating agent. Recently, Cottam and Ward (132) found that with the titration of apo-alkaline phosphatase with Zn(I1) up to a mole ratio of four Zn(II/ dimer results in no increase in the W l NMR linewidth, ". . . while in previous studies of zinc activated biological reactions, a large increase in the chloride linewidth was observed with zinc bound to macromolecules." However, an increase in the chloride linewidth is observed when the pH is decreased below 5.0. This was interpreted as showing that Zn(I1) in alkaline phosphatase is not exposed to solvent at p H > 5.0. In an ESR study of Cu(I1) binding to alkaline phosphatase, Csopak and Falk (133) reported that two Cu(I1) binds to the same specific sites as the two Zn (11), that the ESR spectrum for the one copper enzyme is different from the two copper enzymes, and that phosphate binding causes a shift of the spectral lines, A discussion of the kinetic studies of the Co(I1) alkaline phosphatase 132. G.Cottam and R. Ward, Federation Proc. 29, 868 (1970). 133. H.Csopak and K. E. Falk, FEBS Letters, 7, 147 (1970).

404


(94, 95) is covered in the kinetics portion of this chapter, and a more detailed discussion of the role of metals in alkaline phosphatase can be found in Chapter 18 by Fernley and in a review by Spiro (133a).

E. NUMBEROF ACTIVESITES The data relevant to the number of active sites of alkaline phosphatase can be divided into two groups: One group derived from studies a t low substrate concentrations (S 2 lo-’ M ) indicates one active site per dimer, and the other group derived from studies a t high substrate conM ) indicate two sites. centrations (S 2 Heppel et al. (28) found that the Michaelis-Menten form held a t low substrate concentrations, but a t higher substrate Concentrations ( > lo-’ M ) there was substrate activation, i.e., the activity exceeded the extrapolated V,,,. A double reciprocal plot over the entire range fell along two distinct straight lines. Thus if the substrate concentrations are kept in the high range where the Michaelis form also holds, two V,,,, values (one roughly 30% larger than the other) and two K , values can be determined. This effect was described as substrate activation and was not taken as evidence for a second site, although, of course, it could be so taken. At even higher substrate concentration, there is substrate inhibition (134). Substrate activation does not occur in solutions of high ionic strength (96, 106, l34), but it has been suggested that it might be masked by substrate inhibition (134). “Active site burst titrations” of the Zn(I1) enzyme and the Co(I1) enzyme a t acidic pH and low substrate concentrations [ (70-1) X MI indicated one active site (96, 98, 99, 110,135), while similar experiM ) yielded a ments (136) a t high substrate concentration (2 x value of 2.7 sites per dimer. I n the latter experiment, error in the number of active sites might arise from the fact that the results were obtained by extrapolation from steady state measurements without direct observaion of the pre-steady-state phase. In the low substrate experiments, the pre-steady-state phase was observed directly. A burst was obtained a t intermediate substrate concentrations (4 x lo-‘ M ) , but the “size” was not reported (137). The amount of phosphoryl enzyme formed by the addition of phos133a. T. G. Spiro, in “Inorganic Biochemistry” (G. Eichorn, ed.). Elsevier, Amsterdam, 1971. 134. R. T. Simpson and B. L. Vallee, Biochemistry 9, 953 (1970). 135. S. H. KO and F. KBzdy, JACS 89, 7139 (1967). 136. W. K. Fife, BBRC, 28, 309 (1967). 137. A. Williams, Chem. Commun. No. 19, p. 676 (1966).

17.

E. C O l i


405

phate to an enzyme solution has been studied by acid precipitation techniques. I t was found that covalently bound phosphate approaches 1.0 equivalent a t low pH in the presence of +lo-' M phosphate (31, 35, 54, 114, 116, ldO), and approaches 2.0 equivalents a t low pH in the M phosphate (138). It was also found that the formapresence of tion of the phosphoryl enzyme is dependent on the metal ion ( 5 4 ) . I n contrast to the Zn(I1) enzyme, the Cd(I1) protein (inactive) in equilibrium with phosphate, forms a significant concentration of the phosphoryl enzyme in the alkaline pH range, reaching a maximum of 1 equivalent per dimer of enzyme a t pH 7.0. Mn(I1) and Co(I1) proteins also form significant amounts of phosphoryl enzyme in the alkaline pH region but less than the Cd (11) protein ( 5 4 ) . Equilibrium dialysis studies with alkaline phosphatase determine the sum of the covalently and noncovalently bound phosphate. Phosphate binding determined by this technique is found to remain relatively constant between pH 8 and 5 ( 5 4 ) .The results of the dialysis method show that one phosphate is bound with a binding constant of + lo6M (53, 54, 134) , while a second phosphate is bound with a binding constant of < lo3M ( 1 3 4 ) . It is also found that when the ionic strength is decreased from 1.0 to 0.05, the first phosphate binding constant increases by a factor of two and the second binding constant decreases by a factor of ten. I n other studies it was concluded that the binding of the first phosphate is Zn(I1) dependent because half as much phosphate is bound when the enzyme contains only one zinc atom, while the binding of the second phosphate is Zn(I1) independent since a t high phosphate concentrations the apoenzyme will bind phosphate ( 5 4 ) . Circular dichroic titrations of Co (11) alkaline phosphatase with both phosphate and arsenate also confirm the presence of a single unique anion binding site a t alkaline pH and low anion concentration (54, 1.29). By quenching enzyme solutions during the hydrolysis of 32P-ATPand T - A M P a t high substrate concentrations ( > M ) , Lazdunski et a2. (138) found that two moles of phosphate are bound per mole of enzyme a t low pH and one mole per mole of enzyme a t alkaline pH. I n contrast, by means of a rapid sampling apparatus, Reid and Wilson found only 0.1 mole of phosphate bound per mole of enzyme during the hydrolysis of 32P-ATP, 32P-PPi,and 32P-p-nitrophenyl phosphate a t low substrate concentrations (7 the rate determining step is phosphorylation. Several workers have found “burst” kinetics for various substances a t low pH for both Zn(I1) and Co(I1) enzymes (96, 98,135-137,147). The rate of phosphorylation of the Co(I1) enzyme is much faster than the Zn(I1) enzyme; however, the rate of steady state hydrolysis by the Zn(I1) enzyme is twice as fast as by the Co(I1) enzyme ( 9 6 ) . “Burst” 147. H. N. Fernley and P. G . Walker, BJ 110, l l p (1968).

17. E.

411


kinetics have also been found for the Co(I1) enzyme a t pH 8.0 (96). Thus the point of disagreement seems to be the rate determining step for the Zn(I1) enzyme a t high pH. In order to account for the fact that almost all substrates are hydrolyzed at the same rate a t high pH, even though dephosphorylation is not the sole rate determining step, Trentham and Gutfreund (98, 148) proposed that the mechanism involves a first-order rearrangement of the enzyme-substrate complex. This step is slow compared with the subsequent transfer of phosphate from substrate, S, to enzyme: the final step being the liberation of phosphate from a phosphoryl enzyme intermediate : E

+

S

E .S

slow

E*.S

--

Phosphorylation, etc.

(El)

In order to test Scheme 111, the kinetics of the combination of the reversible competitive inhibitor, 2-hydroxy-5-nitrobenzylphosphonate with the enzyme were studied by the stopped-flow and temperature-jump techniques (107).A relatively slow change in optical absorption was found. This was interpreted as indicating that a first complex between inhibitor and enzyme occurs rapidly but leaves the absorption unchanged. This silent complexation step is very rapid and is followed by a slow conformational change in the complex which is reported by a change in optical absorption. The rate of this conformational change is the rate controlling step in the hydrolysis of substrates. This is a reasonable interpretation of the changes in absorption, but the new hydrolytic scheme has some shortcomings. It does not automatically explain why all substrates are hydrolyzed a t about the same rate. Trentham and Gutfreund assumed that the conformational change of the E - S complex occurs a t a fixed rate independent of the nature of S. First, this assumption, while better, is not too different from assuming that the change E * S+ E-P ROH is independent of S. Second, the increased rate of production of ROH in the presence of an acceptor such as ethanolamine is not explained nor is the observation that there is little or no change in the rate of formation of phosphate. Indeed, this mechanism predicts that the rate of formation of phosphate would be decreased by an acceptor. The slow change in the absorption of 2-hydroxy-5-nitrobenzylphosphoric acid in the presence of enzyme observed by Trentham and Gutfreund can be interpreted in a somewhat different way. Assume there are

+

148. H. Gutfreund, BJ 110, 2p (1908).

412


two enzyme conformations possible, E, and Ep, and further assume that substrates combine much more readily with Ep than E,: E,

Ep

+S

slow

+ Ep Eo. S

-

etc.

The result is that E, is pulled to Ep to Ep*Son addition of S; there is no “silent” complex and the rate is automatically independent of S. This interpretation is thermodynamically equivalent to the interpretation of Trentham and Gutfreund but kinetically different and leads to different kinetic predictions. The full scheme based on Trentham and Gutfreund’s suggestion that a rate controlling conformational change is involved, but interpreted in a somewhat different manner, is

Scheme IV is presented for “zero” concentrations of ROH, S, and Pi corresponding to initial rates of hydrolysis and transphosphorylation, thereby enabling the scheme to be simplified in that single arrows (unidirectional steps) can be employed in appropriate places. I n this scheme the leaving group of any substrate serving as an acceptor must be able to react with Ep-P; it may or may not be able to react with E,-P. If an acceptor can react readily with E,P it would imply that the conjugate substrate could react readily with E, [unless Em-P were a high energy form and S were a normal ester (I@)] ; and the distinction bet,ween E, and Ep would disappear. It is better for the time being to postulate that E,P reacts rapidly only with water and Ep-P only with other leaving groups. The increase in rate of formation of p-nitrophenol from p-nitrophenyl phosphate as a substrate, in the presence of an acceptor, tris, is explained by the scheme which shows that the enzyme form produced by transphosphorylation is the active Ep. The slow conversion of E, to Ep is thus 149. If we allow E,P to be a high energy form, then PI could be an acceptor and PPi a substrate acting on E..

17. E.

413


bypassed and ED-P can be quickly reformed; the result is that the rate of formation of Pi can remain nearly the same. This scheme leads to the Michaelis form with

For purposes of simplification the concentrations of acceptor, R O H , and water are included in their respective rate constants k,' and k l . To meet the original aim of the scheme, namely, to have the rate of hydrolysis independent of the substrate and yet not have dephosphorylation rate controlling, it is necessary to suppose that k,' 6. Thus the bigger k,' is relative to the rate of hydrolysis, the smaller 6 would be; hence, the ratio of maximum velocities would be larger than the percent phosphorylation.


Mammalian Alkaline Phosphatases H . N . FERNLEY I . Introduction . . . . . . . . A . General Survey . . . . . . B . Distribution . . . . . . . C . Function . . . . . . . I1. Molecular Properties . . . . . . A . Purification Procedures . . . . B. Physical Properties . . . . . C . Chemical Modification of Phosphatases I11. Catalytic Properties . . . . . . A . Substrate Specificity . . . . . B. Reaction Catalyzed . . . . . C . Assay Techniques . . . . . D. Kinetic Studies . . . . . . I V . Mechanism of Enzymic Action . . . .

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. . . . .

. . . . .

.

.

.

.

.

417 417 420 421 422 422 423 427 428 428 430 432 434 443

.

1 Introduction

A . GENERALSURVEY Alkaline phosphatases catalyze the hydrolysis of almost any phosphomonoester to give Pi and the corresponding alcohol. phenol. or sugar. etc. They are distinguished from a similar group of enzymes. the acid phosphatases. primarily by the pH dependence of their catalytic activity which is higher in the alkaline range than a t neutral or acid pH . The earliest literature references to (alkaline) phosphatases established that extracts from many mammalian tissues. particularly kidney and in417

418

H. N. FEBNLEY

testinal mucosa ( I ) , could hydrolyze nucleotides, hexosephosphates, and glycerophosphate (2). Robison’s demonstration that ossifying cartilage was a rich source of phosphatase (3) marked the beginning of a phase of intensive investigation, developing the hypothesis that phosphatase was directly involved in the process of calcification. Bone phosphatase was found to have an optimum activity at pH 8.4-9.4 ( 4 ) similar in this respect to the enzymes from kidney and intestinal mucosa and to that present in blood plasma ( 5 ) . Other tissues with relatively high phosphatase activity, such as liver and spleen, were shown to contain a n additional enzyme with optimum activity a t pH 5 ( 6 ) .Davies proposed they be termed alkaline and acid phosphatase, respectively, by which names they are presently accepted. Erdtman made the important observation that kidney phosphatase, after dialysis, required Mg2+ for maximum This was confirmed and extended to include all mammalian activity (7). alkaline phosphatases whether dialyzed or not (8) and has led to the general practice of including Mg2+in the assay medium. Clinical interest in phosphatases developed with the recognition that certain pathological conditions, notably obstructive jaundice (9), rickets (lo),and various bone diseases (11) , were characterized by large increases in blood plasma alkaline phosphatase levels. Accurate measurements were required for diagnosis, and this led to the development of sensitive methods for estimating phosphatase activity (12, IS). Folley and Kay established that hydrolysis of phenyl phosphate by milk phosphatase obeyed Michaelis-Menten kinetics provided initial rate measurements were employed (14). Two complicating factors had been recognized, one a strong inhibition by product, Pi (15, 1 6 ) , the other an inhibition a t high substrate concentration (16, 17). 1. P. A. Levene and F. Medigreceanu, JBC 9, 65 (1911); P. Grosser and J. Husler, Bbchem. 2. 39, 1 (1912). 2. H. von Euler, 2. Physiol. Chem. 79, 375 (1912); R. H. A. Plimmer, BJ 7, 43 (1913). 3. R. Robison, BJ 17, 286 (1923). 4. R. Robison and K. M. Soames, BJ 18, 740 (1924). 5. M. Martland and R. Robison, BJ 20, 847 (1926). 6. D. R. Davies, BJ 28, 529 (1934). 7. H. Erdtman, 2. Phgsiol. Chem. 172, 182 (1927); 177, 211 and 231 (1928). 8. H. D. Jenner and H. D. Kay, JBC 93, 733 (1931). 9. W. M. Roberts, Bn’t. Med. J. I, 734 (1933). 10. H. D. Kay, Brit. J . Ezptl. Pathol. 10, 253 (1929). 11. D. Hunter, Lancet i, 897, 947, and 999 (1930) ; H. D. Kay, JBC 89, 235 (1930). 12. H. D. Jenner and H. D. Kay, Bn’t. J. Exptl. Pathol. 13, 22 (1932); E. J. King and A. R. Armstrong, Can. M e d . Assoc. J . 31, 376 (1934). 13. A. Bodansky, JBC 101, 93 (1933).

18.

MAMMALIAN ALKALINE PHOSPHATASES

419

More recently, isotopic labeling experiments have assumed a major role in establishing the detailed mechanism of enzymic action. It was shown that alkaline phosphatase possesses transferase activity whereby a phosphoryl residue is transferred directly from a phosphate ester to an acceptor alcohol (18). Later it was found that the enzyme could be specifically labeled a t a serine residue with 32P-Pi(19) and that szPphosphoserine could also be isolated after incubation with 32P-glucose 6phosphate (2O),providing strong evidence that a phosphoryl enzyme is an intermediate in the hydrolysis of phosphomonoesters. The metal-ion status of alkaline phosphatase is now reasonably well resolved (21-23). Like E . coli phosphatase it is a zinc metalloenzyme with 2-3 g-atom of Zn2+per mole of enzyme. The metal is essential for catalytic activity and possibly also for maintenance of native enzyme structure. The relationship between the various tissue alkaline phosphatases has been under discussion for many years (24).Bodansky established that inhibition by bile acids could be used to distinguish between intestinal and bone or kidney isoenzymes ( 2 6 ) .The organ-specific behavior of rat tissue phosphatases toward a variety of compounds was investigated by Fishman (26).Of particular importance was the observation that Lphenylalanine is a stereospecific inhibitor for the intestinal isoenzyme (27).Immunochemical (28,29) and electrophoretic techniques (SO, 31) have shown that there are also physical differences between the tissue phosphatases. It is not yet clear what the precise nature of these differences is (%), although in part it results from a variability in sialic acid content. S. J. Folley and H. D. Kay, BJ 29, 1837 (1935). M. Martland and R. Robison, BJ 21, 665 (1927). E. Jacobson, Biochem. Z . 249, 21 (1932). E. Bamann and E. Riedel, Z . Physwl. Chem. 229, 125 (1934). 0. Meyerhof and H. Green, JBC 183, 377 (1950). L. Engstrom and G. Agren, Acta Chem. Scand. 12, 357 (1958). 20. L. Engstrom, Arkiv Kemi 19, 129 (1962). 21. J. C. Mathies, JBC 233, 1121 (1958). 22. L. Engstrom, BBA 52, 36 (1961). 23. D. R. Harkness, ABB 126, 503 (1968). 24. S. Belfanti, A. Contardi, and A. Ercoli, BJ 29, 842 and 1491 (1935). 25. 0. Bodansky, JBC 118, 341 (1937). 26. W. H. Fishman, S. Green, and N. I. Inglis, BBA 62, 363 (1962). 27. W. H. Fishman, S. Green, and N. I. Inglis, Nature 198,685 (1963). 28. M. Schlamowitz and 0. Bodansky, JBC 234, 1433 (1959). 29. H. H. Sussman, P. A. Small, and E. Cotlove, JBC 243, 160 (1968). 30. D. W. Moss, D. M. Campbell, E. Anagnostou-Kakaras, and E. J. King, BJ 81, 441 (1961). 31. A. W. Hodson, A. L. Latner, and L. Raine, Clin. Chim. Acta 7, 255 (1962). 14. 15. 16. 17. 18. 19.

420

€ N. I.FERNLEY

For a more detailed account of phosphatase history and properties the reader is referred to earlier reviews (SS-SS),the more clinical aspects being covered by Gutman (@), Posen (41), and Fishman and Ghosh (42).

B . DISTRIBUTION Alkaline phosphatase is found in bacteria and fungi, although often it is repressed by Pi (43).There is little information concerning its distribution in invertebrate tissues; it is present in the Indian leech (4.4) and in developing Drosophila (45) and the surf clam (46‘). It is relatively abundant in fishes (47) and mammals; it is absent from higher plants (36‘).For mammals, tables of activities have been published (34, 35,&), but some of the values recorded may be lower than the true values because of the long incubation periods, high substrate concentrations, and often high product concentrations involved in the assays. The order of activities for different tissues is reasonably well established with intestinal mucosa + placenta > kidney + bone > liver +lung c spleen. In rat intestinal mucosa the alkaline phosphatase activity is about 0.2 pmole substrate hydrolyzed per minute per milligram of protein, similar to the total ATPase activity (49). Comparable values for rat kidney (50) and human placenta (23)are 0.06 t 0.05 pmole/min/mg of protein and 10.2 pmole/min/g of tissue, respectively. Another organ rich in alkaline 32. J. C. Robinson and J. E. Pierce, Nature 204, 427 (1964); D. W. Moss, R. H. Eaton, J. K. Smith, and L. G. Whitby, BJ 98, 32C (1966). 33. R. Robison, Ergeb. Enzymforsch. 1, 280 (1932). 34. H. D. Kay, Physiol. Rev. 12, 384 (1932). 35. S. J. Folley and H. D. Kay, Ergeb. Enzymforsch. 5, 159 (1936). 36. J. Roche, “The E’nzymes,” 1st ed., Vol. 1, p. 473, 1950. 37. J. Roche and Nguyen-van-Thoai, Advan. Enzymol. 10, 83 (1950). 38. T. C. Stadtman, “The Enzymes,” 2nd ed., Vol. 4, p. 55, 1960. 39. R. K. Morton, Comp. Biochem. 16, 55 (1965). 40. A. B. Gutman, Am. J . Med. 27, 875 (1959). 41. S . Posen, Ann, Internal Med. 67, 183 (1967). 42. W. H. Fishman and N. K. Ghosh, Advan. Clin. Chem. 10, 256 (1957). 43. T. Horiuchi, S. Horiuchi, and D. Mizuno, Nature 183, 1529 (1959); J. F. Nyc, R. J. Kadner, and B. J. Crocken, JBC 241, 1468 (1966). 44. D. Bhoomittra, J . HCtochem. Cytochem. 12, 311 (1964). 45. H. Schneiderman, W. J. Young, and B. Childs, Science 151, 461 (1966). 46. P. Strittmatter, H. B. Burch, and L. Laster, BBA 100, 304 (1965). 47. 0. Bodansky, R. M. Bakwin, and H. Bakwin, JBC 94, 551 (1931). 48. M. G. Macfarlane, L. M. B. Patterson, and R. Robison, BJ 28, 720 (1934). 49. J. P. Quigley and G. S. Gotterer, BBA 173, 456 (1969). 50. R. Kinne and E. KinneSaffran, European J . Physiol. 308, 1 (1969).

18.


421

phosphatase is guinea pig mammary gland with an activity of 8.5 k 5.2 pmole/min/g of moist tissue (14). The distribution within a particular tissue is, of course, not homogeneous. Kay noted that kidney cortex was much richer in phosphatase than the medulla (61) and also that activity along the intestinal tract was variable (66).An extensive study of intestinal phosphatase activities during development has been made by Moog (63).Histochemical studies indicate that the intestinal enzyme is localized predominantly a t the surface membrane of the epithelial cell microvilli (54, 66),a feature confirmed by centrifugal fractionation of intestinal epithelial cells (66). Similarly, kidney alkaline phosphatase is found in the brush border of the proximal tubule epithelial cells (60). I n placental tissue the enzyme is located a t the surface of the trophoblastic syncytium (67),in liver it is adjacent to the bile canaliculi (68),and in bone it is found in hypertrophic cartilage cells, osteoblasts, and osteocytes (69). Generalizing one can say that alkaline phosphatase is abundant in those tissues concerned with transport of nutrients; it is often present in secretory organs and developing tissues ; it is almost absent from muscle, mature connective tissue, nonossifying cartilage, and red blood cells.

C. FUNCTION At the present time it is not possible to assign a precise function for any alkaline phosphatase. Undoubtedly bone phosphatase is concerned in ossification and two alternative roles have been proposed: (1) Precipitation of calcium phosphate is induced by the localized production of high concentrations of Pi owing to phosphatase activity (33) ; (2) the enzyme permits crystal growth a t nucleation sites in the matrix by ensuring the removal and continued absence of PPi which is known to be a crystal “poison” (60). Other factors must be involved (36)because tissues with high concentrations of alkaline phosphatase (e.g., gut, kidney, and 51. H. D. Kay, BJ 20, 791 (1928). 52. H. D. Kay, BJ 22, 856 (1928). 53. F. Moog, Federation Pmc. 21, 51 (1962). 54. S. L. Clark, Am. J. Anat. 109,57 (1961). 55. S. Ito, Federation Proc. 28, 12 (1969). 56. J. W. Porteous and B. Clark, BJ 96, 159 (1965). 57. M. Wachstein, J. G . Meagher, and J. Ortis, Am. J. Obstet. G y e c o l . 87, 13 (1963). 58. M. Wachstein and E. Meisel, A m . J. Clin. Pathol. 27, 13 (1957). 59. E.Borghese, Intern. R e v . Cytol. 6, 289 (1957). 60. H. Fleisch and W. F. Neumann, A m . J. Physiol. 200, 1296 (1961); H. Fleisch, R. G. G. Russell, and F. Straumann, Nature 212, 901 (1966).

422

H. N. FERNLEY

placenta) do not normally calcify, while tissues such as aorta, in which phosphatase is absent, can be made to calcify. Furthermore, rachitic cartilage has a high phosphatase activity yet it will not ossify. In the other tissues which are major sources of phosphatase it is perhaps significant that the enzyme is localized at the absorptive surface, suggesting a direct role in the transport of nutrients across the epithelial membrane. Bodansky found an increase in plasma phosphatase following ingestion of carbohydrate ( 6 1 ) . Ingestion of fat by rats gave an increased synthesis of intestinal phosphatase which later appeared in the blood plasma and lymph ducts ( 6 2 ) .This may correlate with the histochemical finding that intestinal epithelial cells from rats on a high fat diet showed marked phosphatase activity in the Golgi region ( 6 3 ) .A careful investigation by Langman and collaborators (64) indicated that ingested fat (not carbohydrate) leads to the appearance of intestinal phosphatase in the plasma of certain groups of individuals. Regulation of alkaline phosphatase activity was discussed by Cox and Griffin (66). Steroids such as hydrocortisone or prednisolone (1 pg/ml of medium) can induce a 3- to 20-fold rise in certain HeLa cell cultures, while other cell lines are induced by 15 mM phenyl phosphate. Recently, it has been shown that fibroblast cell cultures are also stimulated by prednisolone (66). According to one report, Pi may have a control function; here a decrease in the level of rat kidney Pi produced by a low phosphate diet was accompanied by an increase in alkaline phosphatase. Nine other enzymes monitored were unaffected (67).


A. PURIFICATION PROCEDURIB Methods for the extraction and purification of many tissue phosphatases have been published. The most highly purified preparations are 61. A. Bodansky, JBC 104, 473 (1934). 62. R. M. Glickman, D. H. Alpers, G . D. Drummey, and K. J. Isselbacher, BBA 201, 228 (1970). 63. K. Watanabe and W. H. Fishman, J. Hktochem. C y t o c h m . 12, 252 (1964). 64. M. J. S. Langman, E. Leuthold, E. B. Robson, J. Harris, J. E. Luffman, and H. Harris, Nature 212, 41 (19ss). 65. R. P. Cox and M. J. Griffin, ABB 122, 552 (1967). 66. M. D. Waters and G. K. Summer, BBA 177, 650 (1969). 67. F. Melani, G. Ramponi, M. Farnararo, E. Cocucci, and A. Guerritore, BBA 138, 411 (1967).

18.


423

those from calf intestinal mucosa (68, 6 9 ) , horse or pig kidney (21,69a, 7 0 ) , bovine liver ( 7 l ) , and human placenta (29, 7 2 ) . The enzyme from the last-named source has been obtained in crystalline form (23, 73). Other preparations described include those from calf bone (74) bovine brain ( 7 5 ) , bovine synovial fluid ( 7 6 ) , milk (77), dog feces (78), and human leukocytes ( 7 9 ) . The enzyme is normally firmly attached to lipoprotein membranes but can be solubilized either by autolysis [mincing or grinding the tissue followed by incubation for several days in the presence of 25% acetone (80)] or by treatment with n-butanol (mincing followed by stirring with excess butanol for about 30 min). After extraction the enzyme is reasonably stable and purification is straightforward. Crude extracts are usually subjected to a series of fractionations with ammonium sulfate and acetone or ethanol. More recently, supplementary techniques such as DEAE-cellulose chromatography and gel filtration have been used to achieve a high degree of purification. Quoted recoveries are good, averaging about 25% of the activity in the initial extract.

B. PHYSICAL PROPERTIES 1. Composition

Some properties of those enzymes that have been sufficiently purified to allow a detailed evaluation of their physical parameters are listed in Table I (22, 23, 69, 69a, 71, 73, 81, 8 2 ) . Molecular weights have been 68. R. K. Morton, BJ 57, 595 (1954). 69. P. Portmann, 2.Physwl. Chem. 309, 87 (1957). 69a. P. Portmann and G. Gerfaux, Chimia (Aarau) 15, 428 (1961). 70. F. Binkley, JBC 236, 735 (1961). 71. L. Engstrom, BBA 92, 71 (1964). 72. J. G. Georgatsos, ABB 121, 619 (1967). 73. N. K. Ghosh and W. H. Fishman, BJ 108, 779 (1968). 74. G. Agren, 0.Zetterqvist, and M. Ojamb, Acta Chem. Scand. 13, 1047 (1959). 75. C. Brunel, G. Cathala, and M. Saintot, BBA 191, 621 (1969). 76. D. Dabich and 0 . W. Neuhaus, JBC 241, 415 (1966). 77. R. K. Morton, BJ 55, 795 (1953). 78. M. A. M. Abul-Fadl and E. J. King, BJ 44, 431 (1949). 79. S. Trubowitz, D. Feldman, S. W. Morgenstern, and V. M. Hunt, BJ 80, 369 (1961).

80. M. A. M. Abul-Fadl, E. J. King, J. Roche, and Nguyen-van-Thoai, BJ 44, 428 (1949). 81. N. K. Ghosh, 5. S. Goldman, and W. H. Fishman, Enzymologiu, 33, 113 (1967). 82. E. B. Robson and H. Harris, Ann. Hum. Genet. 30, 219 (1967).

424

H. N. FERNLEY

TABLE I PROPERTIES OF MAMMALIAN ALXALINEPHOSPWTASES ~

Tissue source Calf intestinal mucow Horse kidney Bovine liver Human placenta 4

Ref.

Specific activity'

(88)

2070

100,000

(69a) (71)

1990 1290 70CP

150,000? 125,000

(23)

~~

Zns+ Molecular content Sialic weight (%) acid 0.2

0.15

0

+'

+

Hexose hexosamine

Approx. tissue mnc:

Variable, about 20%

60J

26%

15 1.5 80

+d

As micromoles of substrate hydrolyzed per minute per milligram of protein a t 37"

unless otherwise recorded.

* Value a t room temperature. From Ghosh et al. (81) and Robson and Harris (82). From Ghosh and Fishman (73). As milligrams of protein per kilogram of moist tissue. From Portmann (69).

reported for some other preparations, e.g., milk, 190,OOO (83) and synovial fluid, 72,000 (76).Turnovers per active site have been recorded for enzymes from several sources: milk, 2700 sec-' a t 25" (83); calf intestine, 1450 sec-l a t 20" (Fig. 2 ) ; and human liver and intestine, 5030 and 6550 sec-' a t 37" (84). The placental enzyme is thought to be a dimer of equal weight subunits (86) which are not necessarily identical: Genetic studies have indicated that there are three common types of subunit which can combine to give six electrophoretically distinguishable variants (82). No similar genetic variation has been observed with other tissue alkaline phosphatases. Higher molecular weight forms of placental phosphatase have been observed (73,89),while at pH values above 10.5 reversible dissociation into monomers has been found to occur (86). Amino acid compositions of placental phosphatase have been published (23,73, 86).Two are in good agreement and are given in Table 11. The overall composition is remarkably similar to that of E . coli phosphatase (87)-the relative percentages of amino acid residues on a molar basis are ( E . coli values in parentheses): acidic, 21 (21) ; basic, 14 (12); hydrophilic, 15 (17) ; and nonpolar, 51 (51). The amino acid sequence around the reactive serine group of calf intestinal phosphatase is A s p 83. 84. 85. 86. 87.

T. E. Barman and H. Gutfreund, BJ 101, 460 (1966). D. W. Moss, R. H. Eaton, and P. B. Scutt, BBA 154,609 (1968). A. J. Gottlieb and H. H. Sussman, BBA 160, 167 (1968). H. H. Sussman and A. J. Gottlieb, BBA 194, 170 (1969). F. Rothman and R. Byrne, JMB 6, 330 (1963).

18.

425


Amino acid Ala 4-%

ASP CYS Glu GlY His Ile Leu

Residuesa 123 65 109 14* 112 100 31 38 87

104 60 100 108 100 28 36 88

Amino acid LYS Met Phe Pro Ser Thr TrP TYr Val

Residues 49 24 39 56 52 68

52 24 36 60 64 68

C

37 70

36 80

Data are from Harkness (B), Table 11, and Sussmann and Gottlieb (86), Table I, and are given as moles per 100 moles of glycine. b Measured as cysteic acid. c Titration with N-bromosuccinimide gave 0.45 g tryptophan per 100 g of protein. 0

Ser-Ala (88), identical with the corresponding sequence in the E . coli enzyme (89). Ultraviolet absorption spectra have been published for enzymes from the following sources: calf intestine (N), horse kidney (69a),and human placenta ( 2 3 ) . For crystalline placental phosphatase El$ nm = 7.8 (in 0.05 M phosphate buffer pH 7.0). Titration curves for calf intestinal phosphatase (range pH 4-10) indicate an isoelectric point of 5.7 which is invariant with respect to temperature (15"-25") and ionic strength (0.02-0.5) (91). 2. Stability

At room temperature alkaline phosphatases are generally stable in neutral or mildly alkaline solution but are sensitive to inactivation by acid. Unfortunately, most stability data refer to impure preparations and some of the following statements may need modifying when further information is available. Scutt and Moss investigated the denaturation of human liver and intestinal enzymes a t pH 2.1 and 0" ( 9 2 ) .The liver enzyme was significantly more labile, and both enzymes could be par88. L. Engstrom, BBA 92, 79 (1964). 89. J. A. Schwartz, A. M. Crestfield, and F. Lipmann, Proc. Natl. Acad. Sci. U.S.49, 722 (1963); C. Milstein, BBA 67, 171 (1963). 90. R. K. Morton, BJ 80, 573 (1955). 91. M. Lazdunski, J. Brouillard, and L. Ouellet, Can. J . Chem. 43, 2222 (1965). 92. P.B. Scutt and D. W. Mom, Enzymologia 35, 157 (1968).

426

H. N. FERNLEY

tially reactivated by adjusting the pH to neutrality. Placental phosphatase was irreversibly inactivated after dialysis at pH 2.3 and 4" (86). Inactivation of human tissue phosphatases by urea was extensively studied by Posen and colleagues (9.3). At 37" placental phosphatase had a half-life of 3 hr in 8 M urea while the bone enzyme in 3 M urea had a half-life of only 7 min. Intestinal phosphatase was intermediate in stability. It was also reported that the catalytic activity of placental phosphatase is substantially lower in 8 M urea. There is a similar differential effect of heat on the human isoenzymes which is currently of clinical interest because of a possible application in determining the tissue origins of plasma phosphatase (94). Placental phosphatase is by far the most heat resistant, withstanding 70" for 30 min (95) (in the absence of Mg2+however a partly purified preparation was 63% inactivated). With regard to the other phosphatases, Fishman and Ghosh concluded that human liver and intestinal isoenzymes had similar heat stabilities while bone phosphatase was significantly more labile ( 4 2 ) .HeLa cell cultures produce a phosphatase with a heat stability approaching that of the placental enzyme (66). Milk phosphatase after almost complete heat inactivation was found to undergo a slow reactivation under certain conditions (96). An analogous phenomenon is shown by E . coli phosphatase (97) suggesting that heat inactivation is not necessarily an all-or-nothing effect, 3. Effect of Chelating Agents

Cloetens (98) dialyzed pig kidney phosphatase against 0.01 M KCN for 6 days and found a considerable loss in activity. However, several minutes preincubation with Mgz+before assay gave up to 40% recovery of activity. Of a series of metal i p s tested, Zn2+was the most effective giving 70% recovery. Hofstee investigated the effects of glycine, EDTA, and metal ions on calf intestinal phosphatase (99) and concluded that dialysis against EDTA produced an inactive enzyme. Addition of Zn2+ 93. D. J. Birkett, R. A. J. Conyen, F. C. Neale, S. Posen, and J. BrudenellWoods, ABB 121, 470 (1967). 94. S. Posen, F. C. Neale, and J. S. Clubb, Ann. Internal M e d . 62, 1234 (1965); J. F. Kerkhoff, Clin. Chim. Acta 22, 231 (1968); C. W. Small, ibid. 23, 347 (1969) ; M. X. Fitzgerald, J. J. Fennelly, and K. McGeeney, A m . J. Clin. Pathol. 51,

194 (1969). 95. F. C. Neale, J. S. Clubb, D. Hotchkis, and S. Posen, J. Clin. Pathol. 18, 359 (1965). 96. R. L. J. Lyster and R. Aschaffenburg, J . D a i v Res. 29, 21 (1962). 97. L. A. Heppel, D. R. Harkness, and R. J. Hilmoe, JBC 237, 841 (1962). 98. R. Cloetens, Biochem. Z . 307, 352 (1941) ; 308, 37 (1941) ; 310, 42 (1941). 99. B. H. J. Hofstee, ABB 59, 352 (1956).

18.


427

led to reactivation while addition of Mg2+did not. Likewise preincubation with glycine reduced the activity, and reactivation was observed with Znz+but not with Mg2+.Morton confirmed that the purified enzyme was inactivated by EDTA and obtained partial recovery with Mg2+ or Mn2+ (100). In these experiments complexed EDTA was probably still present in the assay mixture since exhaustive dialysis is apparently necessary to remove such chelating agents (101). Mathies found that dialysis of pig kidney phosphatase against EDTA or Mg-EDTA produced a considerable loss of activity and a parallel reduction in the enzyme Zn content (21).A similar correlation between activity and Zn content was found with calf intestinal phosphatase which had been dialyzed a t pH 5 ( 2 2 ) . In this instance activity was not restored by addition of Znz+. A recent paper by Harkness (101a) has done much to clarify the field. It was found that Zn2+ chelating agents such as cysteine, EDTA, and o-phenanthroline (each 1 mM) are all potent inactivators of crystalline placental phosphatase ; for instance, 15 min preincubation with 10 p.M EDTA at pH 10.5 gave 957%inhibition. Full activity was immediately restored by addition of 100 pM Zn2+ compared to only 25% recovery with Mgz+.Preincubation of apophosphatase with 500 p M Zn2+ gave a 30-fold increase in activity while the corresponding values for Mg2+and Co2+were 0.5 and 5-fold, respectively. One can conclude from these results that Zn2+ is an essential metal ion for alkaline phosphatase (possibly replaceable by Coz+) and that Mg2+is much less effective. Nevertheless, there is a large body of evidence demonstrating that Mg2+ effects are specific, large, and reproducible. This will be discussed later (Section 111,D15).

C. CHEMICAL MODIFICATION OF PHOSPHATASES Anagnostopoulos found that the amino group reagents ketene, nitrous acid, formaldehyde, and phenyl isocyanate all inactivated bovine liver and kidney phosphatases (10.2). On the other hand, acetylation of chicken intestinal phosphatase with acetic anhydride gave an active product with optimum activity more alkaline than normal (103).The enzyme preparation was impure and acetylation only 70% complete 100. R. K. Morton, BJ 65, 674 (1957). 101. M. L. Applebury and J. E. Coleman, JBC 244, 709 (1969). 101a. D. R. Harkness, ABB 126, 513 (1968). 102. C. Anagnostopoulos. BBA 4, 584 (1950). 103. W. Cohen, M. Bier, and F. F. Nord, ABB 67, 479 (1957).

428

€ N. I.FERNLEY

so that one does not know how many of the enzyme amino groups were acetylated. Fishman and Ghosh found extensive inactivation of rat intestinal phosphatase by this reagent with little change in the kinetic parameters (10.4). Similarly, acetylation of human tissue phosphatases with acetic anhydride gave low yields of active enzyme with altered electrophoretic mobilities but with little change in Km or p H optimum (105).Carbamoylation of human and pig kidney phosphatases with 0.6 M cyanate gave a product with increased Km and decreased activity (106). I n the writer’s opinion such studies should be related to active site determinations; however, it is clear that some amino groups can be modified without the binding of substrate being affected. Iodoacetamide, iodosobenzoate p-hydroxymercuribenzoate, and N-ethylmaleimide have all been found to inhibit intestinal phosphatases but high concentrations are required (104, 107).It is doubtful therefore whether alkaline phosphatase contains essential thiol groups or indeed possesses any free thiol groups. Recently, chicken intestinal and E . coli phosphatases have been coupled to human IgG with glutaraldehyde for use as antigen detectors (108).


A. SUBSTRATESPECIFICITY In an early review, Kay listed a number of phosphate derivatives that were hydrolyzed by bone phosphatase (34).These included hexose phosphates, glycerophosphates, ethyl phosphate, adenylate, and phenyl had a much lower pH phosphate. It was thought that PP,-which optimum-was also hydrolyzed by the same enzyme. Phosphodiesters were regarded a t best as poor substrates and possibly were not substrates at all. By 1936 it was evident that the situation was more complex than had originally been thought (35).The existence of different classes of phosphatases was recognized ; some, such as hexosediphosphatase (109), were much more specific than others, and it appeared that one tissue could contain several diff erent kinds of phosphatase. Kidney extracts, 104. W. H. Fishman and N. K. Ghosh, BJ 105, 1163 (1967). 105. D.W.Mom, BJ 118, 17P (1970). 106. M.J. Carey and P.J. Butterworth, BJ 111, 745 (1969). 107. M. Lazdunski and L. Ouellet, Can. J . Biochem. Physiol. 40, 1619 (1962). 108. S. Avrameas, Zmmunochemistry 6, 43 (1969). 109. W.Heymann, Monutsschr. Kinderheilk. 48, 14 (1930).

18.


429

for instance, hydrolyzed phosphomonoesters, diesters (110), metaphosphate (111), and phosphoroamidate (112). The last-named activity could be separated from the monoesterase activity. Roche (36) classified nonspecific phosphatases into five groups, the first of which characteristically hydrolyzed phosphomonoesters and included acid and alkaline phosphatases. An authoritative paper by Morton (113) defined the substrate specificity of purified calf intestinal phosphatase: As well as monoesters, phosphocreatine was found to be a substrate (giving creatine and P i ) , but it had a much broader pH-activity curve than either pglycerophosphate or phenyl phosphate. Pyrophosphate derivatives such as PPi, ADP and ATP diesters (e.g., diphenyl phosphate) or triesters (e.g., trimethyl phosphate) were not hydrolyzed. Portmann confirmed that PPi, ADP, and diphenyl phosphate were not substrates (69). The whole question of the specificity was reopened with the discovery that E. coli phosphatase, contrary to an earlier statement (114), hydrolyzed a variety of polyphosphates including metaphosphate of average chain length 8 (97). It was subsequently reported that partially purified phosphatases from several mammalian tissues had appreciable PPr-ase activity a t pH 8.5 (116).This was confirmed (116) and extended to include ATPase and fluorophosphatase activities (117). Proof that the same enzyme is responsible for the monoesterase and PP1-ase activities was afforded by heat inactivation studies, cross inhibition experiments, and inhibition of PPi-ase activity by L-phenylalanine, a specific inhibitor of intestinal phosphatase. It was also found that calf intestinal phosphatase couid be phosphorylated by 32P-PPiand the number of sites so labeled agreed with the number of active sites determined with a monoester substrate using a stopped-flow technique (118). It would seem that the main reason for the confusion with regard to the PPi-ase activity results from the inclusion of Mg2+in the assay. This stimulates the monoesterase activity but almost completely inhibits PPi-ase activity (117). The substrate specificity of human placental phosphatase has recently been determined by Harkness, and a selection of these data is given in Table 111. Compounds not hydrolyzed included trirnetaphos110. K.Asakawa, J . Biochem. ( T o k y o ) 10, 157 (1928); 11, 143 (1929). 111. T.Kitasato, Bwchem. 2. 197, 257 (1928);201, 206 (1928). 112. M.Ichihara, J. Biochem. (Tokyo) 18,87 (1933). 113. R. K. Morton, BJ 61, 232 (1955). 114. A. Garen and C. Levinthal, BBA 38, 470 (1WO). 115. R.P.Cox and M. J. Griffin, Lancet ii, 1018 (1965). 116. D.W. Moss, R. H. Elaton, J. K. Smith, and L. G. Whitby, BJ 102, 53 (1967). 117. H.N. Fernley and P. G. Walker, BJ 104, 1011 (1967). 118. H.N. Fernley and S. Bisaz, BJ 107, 279 (1968).

430

H. N. FERNLEY

TABLE I11 HUMANPLACENTAL ALKALINEPHOSPHATASE RELATIVEREACTIONRATESAND MICHAELISCONSTANTS FOR VARIOUS SUBSTRATES' Substrateb 5'-AMP ADP ATP 5'-UMP UDP UTP PPi PPPi a

Relative rateC 1.00 1.13 0.37 0.79 1.00

0.41 0.18

Relative rate

Km (mM)

Glucose-1-P a-Gly cerophosphate 8-Glycerophosphate

1.00 1.09 1.31

4.8

2-Phosphogly cerate 3-Phosphogly cerate

0.53

Km (mM)d

3.0 4.4 5.7

4.0

0.20

Substrate

Phosphoserine Phosphoethanolamine p-Nitrophenyl-P

1.4

0.63 0.87

1.04

1 .oo

2.9 0.8

From Harkness (IOla).

a Each 10 mM. c

d

In 0.1 M glycine buffer pH 10.5 a t 30". Based on reaction rates at six substrate concentrations spanning the range 3 4 0 mM.

phate, bis-p-nitrophenyl phosphate, phosphocreatine, and aminophosphonates. Observations in the author's laboratory indicate that crude placental phosphatase does hydrolyze phosphocreatine, the reaction being inhibited by L-phenylalanine. Phosphatases from chicken intestine and E. coli have also been reported to hydrolyze cysteamine S-phosphate and other S-phosphates (but not p-nitrophenyl 0-thiophosphate) giving Pi and the corresponding thiol (119). €3.

REACTION CATALYZED

From the preceding data one may conclude that alkaline phosphatases are capable of hydrolyzing compounds containing P- - -F, P- - -0- -C, P- - -0- -P, P- - 4, and P- - -N bonds but not the phosphonate P- - -C bond (120). It has been established that hydrolysis of glucose l-phosphate (121) and other 0-phosphates (122) proceeds entirely by P---0 fission. Stein and Koshland (122) also concluded that a pentacovalent intermediate of type (I) is not likely t o form during the hydrolysis since with H,lsO and phenyl phosphate there was no incorporation of l8O into the substrate. 119. H. Neumann, JBC 243, 4671 (1968). 120. M. Kochman, P. Mastarlerz, and E. Wolna, Arch. Immunol. Therap. Ezptl. 12, 106 (1964). 121. M. Cohn, JBC 180, 771 (1949). 122. S. S. Stein and D. E. Koshland, ABB 39, 229 (1952).

18.

43 1


-0, +,oHO+OH OR

(I)

HO\ $0 HO/'\XR (11)

Studies on the transferase action of milk and intestinal phosphatases have shown that compounds such as glucose, glycerol, and propanediol can accept a phosphoryl residue from a wide variety of donors (123). The overall reaction is therefore transfer of a phosphoryl group from n donor of type (11) where X is F, 0, S, or N and R is H or an alkyl substituent, etc., or may even be absent, to an acceptor of type R--OH where R' is H or an alkyl substituent, with fission of the P---X bond. Since the enzyme must also catalyze the reverse reaction the acceptor specificity should extend to all compounds of the type R--XH. In contrast to the lack of specificity with respect to the nonphosphoryl part of the substrate or acceptor is the strict specificity for the phosphoryl residue. Phosphodiesters and triesters are not hydrolyzed nor are mixed esters of types (111) and (IV) (124).

(111)

(IV)

It appears that only a terminal phosphoryl group is transferred by alkaline phosphatase, and recently it has been stated that ATP is indeed hydrolyzed by stepwise production of P I (125). It follows that metal-ion complexes of phosphomonoesters and polyphosphates should not be substrates if the terminal phosphoryl group is involved in metal binding. The inhibition of PPl-ase activity by excess Mgzt is now well established (117,126); however, it is not certain which of the MgZt-PPI complexes are substrates, inhibitors, or perhaps neither. In the writer's opinion (for the reasons given above) only the uncomplexed species should be substrates. The strong inhibition observed with excess Mg2+ does not necessarily mean that complexes such as MgPPI*- or MgzPPl are inhibitory; it may be attributable to the almost complete removal of free PP, and to a relatively large inhibitory effect from released PI. Little evidence is available as to which is the preferred ionic form 123. R. K. Morton, BJ 70, 139 (1958). 124. H. N. Fernley, unpublished experiment (1988). 125. D. W. Moss and A. K. Walli, BBA 191, 476 (1969). 126. R. H. Eaton and D. W. Moss, BJ 102, 917 (1967); P. J. Butterworth, hid. 110, 671 (1988); P. R. V. Nayudu and P. L. Miles, ibid. 115, 29 (1969).

432

H. N. FERNLEY

of the substrate. One attempt to settle this point has been made (127), but the arguments presented by the authors in favor of un-ionized phosphomonoesters as the true substrate species are questionable.

C. ASSAYTECHNIQUES 1. In Vitro

Early methods for determining phosphatase activity involved gravimetric measurement of the Pi content before and after incubation of substrate with enzyme. In the decade 1920-1930 a number of colorimetric procedures were introduced, based on the reduction of phosphomolybdate (35). This principle is of course still the basis of most PI determinations, though many variations exist. For instance, if the substrate is acid stable, the method of Fiske and Subbarow (188) is suitable, but not very sensitive, and modifications such as that of Chen et a2. (129) may be employed to improve the color yield. With labile substrates milder conditions must be adopted (130, 131), and trouble may also be encountered with a catalyzed hydrolysis of certain esters by molybdate (132). I n such cases i t is preferable to transfer the phosphomolybdic acid to an organic solvent, leaving the ester in the aqueous phase (133). Two recent sensitive methods involve complexing phosphomolybdic acid with malachite green (134) or methyl green (155). Alternatively an enzymic method which will estimate 30 ng PI directly and can be adapted (by cycling the TPNH formed) to measure amounts of Pi down to the limit set by impurities in the reagents and glassware may be employed (136). Trace amounts of 32P-Picaxi be recovered by precipitation with ammonium molybdate and triethylamine after adding carrier Pi (137).The triethylammonium phosphomolybdate is washed free of contaminating ester and then dissolved in acetone or aqueous 127. A. F. Reid and J. H. Copenhaver, BBA 24, 14 (1957). 128. C. H. Fiske and Y. Subbarow, JBC 66, 375 (1925). 129. P. S. Chen, T. Y. Toribara, and H. Warner, Anal. Chem. 28, 1756 (1956). 130. 0. H. Lowry and J. A. Lopez, JBC 162, 421 (1946). 131. J.-L. Delsal and H. Manhouri, Bull. SOC.Chim. BbZ. 40,1623 (1958). 132. H. Weil-Malherbe and R. H. Green, BJ 49, 286 (1951). 133. I. Berenblum and E. Chain, BJ 32, 295 (1938); R. H. Dreisbach, Anal. Biochem. 10, 169 (1965). 134. K. Itaya and M. Ui, Clin. Chim. Acta 14, 361 (1966). 135. H. Van Belle, Anal. Bbchem. 33, 132 (1970). 136. D. W. Schulr, J. V. Passonneau, and 0. H. Lowry, Anal. Biochem. 19, 300 (1967). 137. Y. Sugino and Y. Miyoshi, JBC 239, 2360 (1964).

18.


433

ammonia. As little as 3 ng labeled Pi has been measured by this method (118).

I n practice it is often more convenient to measure the release of a phenol from an aryl phosphomonoester. Standard serum phosphatase methods employ phenyl phosphate (I%?),p-nitrophenyl phosphate (1S9), phenolphthalein monophosphate (IN), or thymolphthalein monophosphate (1.41) where the phenol released can be determined spectrophotometrically [only the Bodansky method (IS) uses a Pi determination]. A number of fluorogenic substrates have been used for phosphatase studies, e.g., P-naphthyl phosphate ($0, I@), 4-methylumbelliferyl phosphate (143), and 3-O-methylfluorescein phosphate (144). The main advantage here is the much greater sensitivity of fluorescence as compared with spectrophotometric assays: as little as 1 pmole of 4-methylumbelliferone can be detected in continuous assay. 2 . Histochemical and Gel Localization

The histochemical localization of alkaline phosphatase has been very extensively investigated; for a survey of this field see Burstone (145). The original method of Gomori involves the in situ precipitation o f released Pi by Ca2+present in the reaction mixture, followed by exchange of Ca2+for Coz+ and subsequent formation of CoS with ammonium sulfide (146). More recent methods are based on the coupling of a phenol, released from a suitable aryl phosphomonoester, to a diazonium salt present in the medium (147) or added later (148). A further development eliminating the need for coupling a t all is achieved by the use of phosphate esters which on hydrolysis give highly insoluble fluorescent (148) or indigogenic ( 1 4 8 ~ phenols. ) 138. P. R. N. Kind and E. J. King, J . Clin. Puthol. 7, 322 (1954). 139. 0. A. Bessey, 0. H. Lowry, and M. J . Brock, JBC 164, 321 (1946). 140. A. L. Babson, S. J. Greeley, C. M. Coleman, and G. E. Phillips, Clin. Chem. 12, 482 (1966). 141. C. M. Coleman, Clin. Chim. Actu 13, 401 (1966). 142. L. J. Greenberg, BBRC 9, 430 (1962). 143. H. N. Fernley and P. G . Walker, BJ 97, 95 (1965). 144. H. D. Hill, G. K. Summer, and M. D. Waters, Anal. Biochem. 24, 9 (1968). 145. M. S. Burstone, “Enzyme Histochemistry,” p. 160. Academic Press. New York, 1962. 146. G. Gomori, Proc. SOC.Ezptl. Biol. M e d . 42, 23 (1939). 147. M. L. Menten, J. Junge, and M. H. Green, Proc. SOC.Exptl. B i d . M e d . 97, 82 (1944). 148. M. S. Burstone, J . Natl. Cancer Znst. 24, 1199 (1960). 148a. P. L. Wolf, J. P. Horwite, J. Vazquez, and E. von der Muehll, Enzymologia 35, 154 (1968).

434

H. N. FERNLEY

Location of alkaline phosphatase on starch or polyacrylamide gels has been achieved using CY- or P-naphthyl phosphate in conjunction with a stabilized diazonium salt, e.g., fast blue RR (149) or fast blue BB (160).

D. KINETICSTUDIES 1. Factors Affecting Activity

Perhaps the most characteristic feature of alkaline phosphatase is the way in which the pH optimum changes with increasing substrate concentration. A typical set of curves for calf intestinal phosphatase and phenyl phosphate is given in Fig. 1. Other examples of this type of behavior are found with P-glycerophosphate and chicken intestinal 1600 -

1200 H c

0

p

800

-

._ 0 c C

J

400 -

Y

6

008.5

I

I

I

I

I

I

9.5

10.0 1 00

9.0 9 0

pH at 38'

FIG. 1. Hydrolysis of phenyl phosphate by calf intestinal alkaline phosphatase. The curves refer to the following substrate concentrations: A, 25 p M ; B, 50 B M ; C, 100 p M ; D,500 p M ; E, 750 p M ; F, 2.5 m M ; G, 25 m M ; and H, 75 mM. Initial velocities are expressed as micromoles of product per milligram of enzyme per minute. From Morton (100).

149. S. H.Boyer, Science 134, 1002 (1961). 150. I. Smith, P.J. Lightstone, and J. D. Perry, Clin. Chim. Acta 19, 499 (1968).

18.


435

(151, 152) or rat intestinal (153) phosphatases, phenyl phosphate, and dog intestinal phosphatase (154), P-naphthyl phosphate and human tissue phosphatases (SO),and p-nitrophenyl phosphate and bovine synovial phosphatase ( 7 6 ) . With different aryl phosphates a t a fixed substrate concentration an analogous set of curves is generated (155). A correlation between the second dissociation constant of the ester and the p H optimum, rate of hydrolysis, and K, value was noted: With increasing pK of the substrate, the pH optimum became more neutral, the rate of hydrolysis decreased, and K,,, increased. With rat intestinal phosphatase and pglycerophosphate a linear relationship was found between the logarithm of the substrate concentration and the pH optimum (153) and has since been confirmed for tissue phosphatases of several mammalian species and of poultry (166). The significance of all these observations is at present far from clear. Any rationalization is difficult because many variables are involved. As Stadtrnan has pointed out (38)the nature of the buffering ions, the presence of activating cations, the different assay techniques, substrates, and enzyme preparations can each influence the overall activity. Attempts have been made to isolate particular factors and these will be briefly discussed. With regard to ionic strength, results from two laboratories using calf intestinal phosphatase suggest the following effects: (1) a t p H 7.5 an increase in ionic strength is associated with a decrease in K , and an increase in V,,, (91); (2) in the p H range 8-10 an increase in ionic strength is associated with a shift to lower pH of the whole activity curve, a decrease in maximum turnover, and a possible decrease in K , (143). The last-named value is difficult to isolate because there is no region of p H in which K m is unchanging. In this instance K , values were compared at pH values for which Vmax was half its maximum value. It should perhaps be emphasized that such effects may complicate p H activity studies unless constant ionic strength buffers are employed. Furthermore, addition of ions such as Mg2+may make a significant contribution to the total ionic strength thus modifying any specific effect. I . Motzok and A. M. Wynne, BJ 47, 187 (1950). I. Motzok, BJ 72, 169 (1959). M. H. Ross, J . 0. Ely, and J. G. Archer, JBC 192, 561 (1951). N. I. Rzhekhina, Biokhimiya 28, 321 (1963). G. E. Delory and E. J. King, BJ 37, 547 (1943); P. G. Walker and E. J. King, ibid. 47, 93 (1950). 156. I. Motzok and H. D. Branion, BJ 72, 177 (1959) ; 80, 5 (1961) ; I. Motzok, ibid. 87, 172 (1963). 151. 152. 153. 154. 155.

436

H. N . FERNLEY

With regard to buffering ions the following are commonly employed in phosphatase assays (4.2, 76, 149), the approximate pK values a t 25" being given in parentheses: ammediol (8.8), borate (9.3), carbonate (10.3), diethylbarbiturate (8.0), ethanolamine (9.5), and tris (8.1). Amine buffers with hydroxyl groups are capable of acting as phosphoryl acceptors (60) and a t high concentration can enhance alkaline phosphatase activity (157). One report states that borate and carbonate are inhibitory for calf intestinal phosphatase (168). I n a detailed study of buffer effects it was shown that pH-activity curves were different for each buffer, for each substrate, and for each tissue phosphatase employed (159). Even altering the buffer concentration may have a marked effect on the activity (160). One can conclude that changing from one buffer system to another generally produces a shift in the pH-activity curve, possibly because of associated changes in ionic strength, while inhibitory effects may be an additional complication.

2. Studies on Km and V,,, Morton (100) recorded plots of pK, against pH for milk and intestinal phosphatases with phenyl phosphate which fitted a theoretical pattern, of zero slope a t lower pH changing to -1 a t higher values with a discontinuity a t pH 9.2, as described by Dixon (161). With p-nitrophenyl phosphate and calf intestinal phosphatase, Lazdunski and Ouellet found a minimum in K,,, a t p H 8.5, while on the alkaline side pK, fell linearly with increasing pH but with a nonintegral slope (166). Dabich and Neuhaus using p-nitrophenyl phosphate and bovine synovial phosphatase reported essentially the same result ( 7 6 ) . Fernley and Walker, using 4-methylumbelliferyl phosphate and calf intestinal phosphatase, also found a nonintegral relationship but were unable to confirm a minimum in K, (143). Other workers have recorded nonintegral negative slopes (156), slopes changing from -1 to 0 a t pH 9 (163), and even slopes changing from 0 to -2 to 0 with discontinuities a t pH S.6 and 9.6 (4.2). A diagram illustrating these various patterns is given in Fig. 2. The 157. T.-U. Hausamen, R. Helger, W. Rick, and W. Gross, Clin. Chim. Acta

15, 241 (1967). 158. C. A. Zittle and E. S. Della Monica, ABB 26, 112 (1950). 159. Z. Ahmed and E. J . King, BBA 45, 581 (1960). 160. N . V. Novikova, V. A. Novitskaya, E. G . Prokof'eva, and N. I. Rzhekhina, Biokhimiya 34, 273 (1969). 161. M. Dixon, BJ 55, 161 (1953). 162. M. Lazdunski and L. Ouellet, Can. J. Chem. 39, 1298 (1961). 163. E. F. Alvarea, M. D. Penalver, and M. Lora-Tamayo, Anales Real SOC. Espan. Fis. Quim. (Madrid) B61, 1039 (1965).

18.

437

MAMMALIAN ALKALINE PHOSPHATASES 6 r

r

:x A

31

I

I I

1

I

I

I

I

FIG.2. Diagram of pH effects on K,. The plots are taken from the following references: A, Fishman and Ghosh (42);B. Morton (100); C, Fernley and Walker (14.9) ; D, Ladzunski and Ouellet (16'8) ; E, Alvarez et al. (163).

main area of agreement here is that K , generally increases with increasing pH and that plots of pK, against p H have linear regions, often with nonintegral slopes. As with K,, the effect of pH on V,,,,, cannot be described by a simple ionization curve, With calf intestinal phosphatase, the log V,,,, curve for a monoester substrate is sigmoid (145, 162) or, in the case of synovial phosphatase, extremely shallow ( 7 6 ) .Both curves approach a maximum value at alkaline pH. Barman and Gutfreund, however, found that milk phosphatase had an optimum a t pH 10 with only 60% activity a t pH 11 (85). This is by no means typical since placental phosphatase has been shown to be fully active with the same substrate, p-nitrophenyl phosphate a t pH 11.5 (86).With PPi as substrate there is evidence that an optimum in V,,,,, is reached a t considerably lower p H values (8.59.2) (116, 117, 164). A pH-activity curve for calf intestinal phosphatase is given in Fig. 3. Features to note are the plateau in activity around pH 7, corresponding to a minimum in the phosphorylation rate constant, and a change in rate determining step a t about p H 6 (166). Taking into account all the above observations, one can formulate the following conclusions. For a given substrate the fact that K,,, inapproaches a limiting value is creases with increasing pH while V,, sufficient to explain, at least qualitatively, the change in pH optimum with substrate concentration. Also changing the buffer or ionic strength 164. H. H. Sussman and E. Laga, BBA 151, 281 (1968). 185. H.N.Fernley and P. G. Walker, BJ 102, 48P (1967).

438

H. N. FERNLEY

3 1

04

I

I

I

I

1

I

5

6

7

8

9

10

PH

FIG.3. Hydrolysis of 4-methylumbelliferyl phosphate by calf intestinal alkaline phosphatase. Activities are recorded as turnovers per site per second a t 20" and I = 0.02, using tris-acetic acid (pH 8) buffers.

may be expected, by virtue of the sensitivity of K , and V,,, to the environment of the enzyme, to produce shifts in the pH dependencies of these parameters and hence in the pH-activity curves. Bodansky estimated the energy of activation for the hydrolysis of ,8-glycerophosphate by bone phosphatase at pH 9 to be 9940 cal/mole (166). For the hydrolysis of p-nitrophenyl phosphate by placental phosphatase a t pH 10.5 the corresponding figure is 10,380 cal/mole ( 1 0 1 ~ ) . Taking into account changes in ionization of the enzyme, a value of 9800 cal/mole for 4-methylumbelliferyl phosphate and calf intestinal phosphatase was derived (143). The comparable values for nonenzymic hydrolysis of monoanions of aryl phosphates are 27,000-31,000 cal/mole (167). With regard to heats of ionization, Lazdunski and Ouellet derived values of 6000-7000 cal/mole for each of three groups a t the active center of calf intestinal phosphatase (162), while an overall figure of 9600 cal/mole, reflecting the shift of log V,,,, curves with temperature along the pH axis was recorded by Fernley and Walker (143). As with changing ionic strength the effect of temperature on K , is difficult to evaluate. If one accepts that K , values for which the corresponding V,,, values are half-maximum are comparable, there is a decrease in K , with increasing temperature (1.43, 162), indicating an increase in entropy associated with Michaelis complex formation. A detailed study of the effects of dioxane and ethanol on calf intestinal phosphatase showed that V,,, for p-nitrophenyl phosphate decreased 166. 0. Bodansky, JBC 129, 197 (1939). 167. C. A. Bunton, E. J. Fender, E. Humeres, and K.-U. Yang, 32, 2806 (1967).

J. Org. Chem.

18.


439

with increasing solvent concentration such that plots of log V,,, against 1/D, where D is the dielectric constant of the assay medium, are linear over the pH range 8.0-10.5 and solvent concentration range 0-30% (91). Morton also observed a decrease in activity with increasing glycerol, glucose, or sucrose concentration (12.3). Lazdunski et al. interpreted their findings in terms of a model whereby activation of the ES complex is accompanied by an increase in molecular diameter. A correlation between the change in diameter (decreasing with increasing charge on the enzyme) and catalytic activity was suggested.

3. Phosphoryl Enzyme Formation This feature has been extensively investigated by Engstrom (20, 71, 88, 168, 169;see also Sections I,A and I1,B) whose results may be summarized as follows: (1) incubation of alkaline phosphatase with 32P-P, at pH 6 6 and O”, followed by acid inactivation, leads to the appearance of the label in the enzyme protein; (2) after acid hydrolysis the only labeled amino acid found is phosphoserine; (3) one mole of Pi is incorporated per mole of enzyme; (4) the presence of Znz+in the enzyme is essential for phosphorylation; (5) bound Pi can be displaced by addition of glucose 6-phosphate to the phosphorylation medium; and (6) very little phosphoryl enzyme is formed under alkaline conditions. Most of these observations have since been verified. Phosphorylation by substrate has been shown to occur under acid conditions by using a stopped-flow technique (118, 165) as illustrated in Fig. 4. Under alkaline conditions the phosphoryl enzyme cannot normally be observed or isolated because the rate of dephosphorylation exceeds the maximum rate of phosphorylation (170).One interesting aspect is that the pH-rate profiles for phosphorylation and dephosphorylation are quite different, as is the case for E . coli alkaline phosphatase (171).Barman and Gutfreund studied the formation and breakdown of milk phosphoryl phosphatase using a rapid-quenching technique and concluded that dephosphorylation could not be rate limiting for the hydrolysis of p-nitrophenyl phosphate at pH 7 (83).

4. Transferase Activity Transfer of a phosphoryl group-mentioned in Sections I,A and 111,Ahas been shown to be independent of the “free energy” of the donor 188. 169. 170. 171.

L. Engstrom, BBA 52, 49 (1961). L. Engstrom, BBA 54, 179 (1961). H. N. Fernley and P. G. Walker, BJ 116, 543 (1970). H. N. Fernley and P. G . Walker, BJ 111, 187 (1969).

440

H. N. FERNLEY

Time ( s e c )

FIG.4. Phosphorylation of calf intestinal alkaline phosphatase by 0.55 p M 4methylumbelliferyl phosphate at pH 4.3 and 20". In terms of scheme (1) (Section IV) the phosphorylation rate constant (haPp) was estimated to be 10.8 sec-', the dephosphorylation rate constant ( h ) ,0.9 wc-'and the active site concentration (EJ, 70 nM. A signal of 10 nA corresponds to the release of 9.75 n M methylumbelliferone. From Fernley and Bisaz (118).

molecule, while the percentage transfer is related to the acceptor alcohol concentration by an equation of the Michaelis form (123).The maximum rate of transfer was however only 8-50% of the total activity, depending upon the particular acceptor, while half-saturation concentrations were in the range 0.44.9M . Where there is a choice of acceptor groups, a8 with glucose or glycerol, transfer occurs preferentially to a primary hydroxyl group [Kay, when demonstrating the equilibration between P-glycerophosphate, Pi, and glycerol catalyzed by intestinal phosphatase ( 5 1 ) , later found that the ester product was in fact mainly a-glycerophosphate ( 1 7 2 ) ] . Morton postulated that there may be one or more acceptor sites on the enzyme: One site would be the more attractive hypothesis, but it must first be explained why saturation effects are observed with only partial transfer. 5 . Effect of Metal Ions on Activity

A most perplexing aspect of alkaline phosphatase behavior is the bewildering variety of effects that have been demonstrated with the addition of divalent metal ions. Activation of various tissue phosphatases is observed with Co2+,Mg2+,and Mn2+ (69, 90,169); other ions, notably Be2+and Zn2+,are inhibitory while Ca2+,Ni2+,and CdZ+probably have little effect. Inhibition by Zn2+is not observed in the presence of glycine 172. H.D.Kay and E. R. Lee, JBC 91, 135 (1931).

18.

441


(93, 99) and that resulting from Be2+is very sensitive to Pi (173).The inhibition of Be2+and Zn2+can be reversed by Mg2+,but the effects are complex (90). Few details are available on the kinetics of Zn2+inhibition, which is surprising in view of the absolute requirement for this ion [one report (174) states that it is noncompetitive or mixed]. The kinetics of Mg2+ activation have been repeatedly investigated though not usually in depth. An illustration is given in Fig. 5. The activation approximately follows a titration curve with a metal ionenzyme dissociation constant of about 500 f l .The slight fall in activity at high Mgz+ concentration may be a nonspecific ionic strength effect. An important feature of the activation is that Mg2+does not affect the affinity for substrate while the affinity for Mg2+has a different p H dependence from that for substrate (163). Unpublished results from this laboratory though differing in some details support the above findings. A recent observation with phenylalanine-inhibited placental phosphatase indicates that Mgz+ may act by increasing the number of active sites rather than enhancing a particular rate (170). It is possible that Mg2+ brings additional sites on the same molecule into action. This cannot be the only mechanism however because of the very large increases sometimes observed (176).Furthermore, one might expect the additional sites to have different properties and apparently they do not. A second

6

5

4

3

2

I

P Mg

FIQ.5. Activation of calf intestinal alkaline phosphatase by Mg". Assays were performed at 38" in 0.05 M ethanolamine-HC1 pH 9.9 with 2.5 m M phenyl phosphate. From Morton (90). 173. W. N . Aldridge, Nature 165, 772 (1950). 174. D. W. MOM,BJ 112, 699 (1969). 175. F. Binkley, BBRC 6, 67 t1961).

442

H. N. FERNLEY

possibility is that Mgz+(and certain other cations) facilitate a redistribution of Zn2+either by restoring it to an essential site or perhaps by removing it from an inhibitory site.

6. Inhibition Studies An important inhibitor of alkaline phosphatase is Pi, normally a very effective competitor with an affinity comparable to that for good substrates (107,117). I n consequence the kinetics of hydrolysis are often approximately first order whatever the substrate concentration, and for this reason initial rate measurements should be limited to 10% hydrolysis. With poor substrates, e.g., phosphocreatine (113) or o-carboxyphenyl phosphate (176), additional care is required. Arsenate is an even more powerful competitive inhibitor (101a, 113) and can prevent the incorporation of P, by intestinal phosphatase a t pH 5 (169).Phosphonates have been reported as weak inhibitors of intestinal phosphatase (l20),and one has been used recently as a chromophoric probe for E . coli phosphatase (177). Many amino acids are weak inhibitors of the various tissue phosphatases (%), and where investigated in more detail the inhibition has been found to be noncompetitive or mixed (178).The effects appear to vary considerably with the nature of the particular enzyme. Cysteine and histidine probably inhibit by virtue of their Znz+ chelating ability (107, 179). Other compounds in this category include iodosobenzoate, iodoacetamide (lor),and ZnZ+( 1 7 4 ) . One component of urea inactivation of human tissue phosphatases has been shown to be a noncompetitive inhibition, reversible on dilution (93). So far the only well-characterized uncompetitive inhibitor is L-phenylalanine, shown to be uncompetitive for intestinal (180) and placental By contrast, D-phenylalanine has no inhibitory effect. phosphatases (4%’). The L isomer apparently acts by preventing the breakdown of phosphoryl phosphatase (170),possibly by blocking the acceptor site mentioned in Section 111,D,4. A list of concentrations required to produce 50% inhibition of a wide variety of phosphatases showed that all the enzymes were affected though there was a spread in susceptibilities from 0.8 m M for HeLa cell culture phosphatase to 26 mM for mouse intestinal phosphatase 176. C. A . Zittle and E. W. Bingham, ABB 86, 25 (1960). 177. S. E. Halford, N. G. Bennett, D. R. Trentham, and H. Gutfreund, BJ 114, 243 (1969). 178. 0. Bodansky, JBC 185, 605 (1946); 0. Bodansky and N. Strachman, ibid. 174, 465 (1948). 179. S. G. Agus, R. P. Cox, and M. J. Griffin,BBA 118, 363 (1966). 180. N. K. Ghosh and W. H. Fishman, JBC 241, 2516 (1966).

18.


443

(65). Fishman et al. recorded that a-phenyl-a-alanine and N-formylphenylalanine do not inhibit rat intestinal phosphatase while 8-phenyl-Palanine, p-fluorophenylalanine, and tyrosine do, although details of the inhibition were not given (26). Another type of inhibition is that resulting from excess substrate. This feature generally determines the shape of the acid limb of the pH-activity curves a t high substrate concentration (see Fig. 1 ) and has been studied in detail (14, 100, 154, 181). A particularly striking example is given by Schmidt and Thannhauser where, with PPi as substrate, the lower the substrate concentration the higher the activity (182). Kay considered that their results could be explained on the basis that an inactive ES, complex was formed, while Rzhckhina postulated the formation of ES, and ES, complexes. There does not appear to be enough data available to settle this issue or to establish whether additional substrate molecules combine a t the same site, a t identical but different sites, or perhaps with a phosphoryl-enzyme intermediate. The latter possibility should be timenable to experimental verification using a rapid-quenching technique. The kinetics of EDTA inhibition are complex (101u, 183), being both time dependent and substrate dependent; however, it is clear that the presence of substrate offers limited protection against inactivation. It has been postulated ( 1 0 1 ~ )that EDTA is initially bound to the enzyme and only prolonged incubation leads to dissociation of Zn2+.The kinetics of reactivation differed for these two cases, being much slower for apophosphatase. I t is also possible that a slow reactivation reflects a change i n protein structure subsequent to Zn2+binding. Diisopropylfluorophosphate, in contrast to its action on other serine Iiydrolases, has only a slight inhibitory effect on alkaline phosphatase in the range 1-10 mM (76, 113).

IV. Mechanism of Enzymic Action

Thc available evidence is consistent with the phosphoryl transfer reaction proceeding via a phosphoryl-enzyme intermediate; of particular relevance is the observation that the rate of decay of this compound is compatible with the overall rate of hydrolysis (83, 118).The sequence of steps for hydrolysis of an 0-ester may be written as in ( l ) ,where ROP 181. A . P. Brestkin and N. I. Rzhekhina, Biokhimiya 30, 471 (1965). 182. G. Schmidt and S. J. Thannhauser, JBC 149, 369 (1943). 183. R. A. J. Conyers, D. J. Birkett, F. C. Neale, S. Posen, and J. Brudenell-Woods, BBA 139, 363 (1967).

444

H. N. FERNLEY

represents the substrate, E H and EP are free and phosphoryl enzyme, ki

EH

+ ROP Sk-iE H . ROP-

ka

\

ROH

ka

EP

EH

(1)

n

H20

Pi

.

respectively, and EH ROP is the Michaelis complex. The Michaelis parameters in terms of these rate constants and the concentration of active sites, Eo, are given in Eqs. (2) and (3).

The early time course of first-product liberation under conditions where the substrate concentration does not change appreciably during the transient state, as in Fig. 5 , is described by Eqs. (4) and ( 5 ) . Use of

kz (5) 1 {@-I kz)l/{ki[ROPI] (1), and the related equations allows evaluation of E,, k , and k,, while k , may, in favorable circumstances, be determined (indirectly) from K, or by varying [ROP] (184). The exponential term describing the first-order formation or decay of phosphoryl phosphatase is equal to [ k , app k3] with [ P i ] replacing [ROP] in ( 5 ) for certain rapid quenching experiments (83).It should be borne in mind that the above rate constants do not necessarily refer to single reaction steps; for instance, Ic, may involve binding of water, transfer of phosphate, and release of Pi as well as possible isomerization steps. With regard to substrate binding, a plot of log V,,JK, against pH may be used to demonstrate ionizations which affect the association of enzyme and substrate (185). The negative slopes observed above pH 9 with bovine phosphatases (7'6,162) suggest that a group with an apparent pK in this region is involved in the binding. By analogy with E . coli phosphatase, where there is good evidence that the metal ion (in this case Co2+) is situated at, or very close to, the binding site for Pi (101), it is probable that the ionizing group involved in the binding of substrate to mammalian alkaline phosphatases is in fact Znz+hydrate. The nonintegral

kz app

=

+

+

+

184. F. Kexdy and M. L. Bender, Biochemistry 1, 1097 (1962). 185. K. J. Laidler, Trans. Faraday SOC.51, 540 (1956).

18.


445

slopes often observed in pK, against p H plots could result from electrostatic effects because of the increasing negative charge on the molecule with increasing pH (91).Possibly two Zn2+groups are associated with each active site since the above plots can have negative slopes greater than unity, the difference not being accountable for by changes in V,,, (143, 162). It is evident from Table I11 that V,,,,, values for P-0-C esters are grouped fairly closely in spite of rather large differences in the p K values of the leaving groups, suggesting that there is a common rate determining step which is independent of the nature of the substrate (within limits). Postulating such a step a t any point after phosphoryl transfer leads to kinetic schemes that generally are implausible in one way or another; for instance, the absence of an initial burst and the inability to isolate phosphoryl enzyme under alkaline conditions are strong arguments against a rate limiting dephosphorylation or later isomerization step. A rate determining conformational change prior to phosphoryl transfer has been proposed for E. coli phosphatase (177,186). For this mechanism one assumes that the initial Michaelis complex is catalytically inert and that an isomerization step is required to generate an “active site.” With regard to the mammalian phosphatases, the dissimilar p H dependencies of k , and k , support the idea that E H - R O P and EP have different conformations. Also, it is otherwise difficult to account for the differential action of L-phenylalanine on lc, and k,. An alternative possibility is for a conformational change and phosphoryl transfer to occur simultaneously. From this viewpoint the transition state for the conformational change is the same as for phosphoryl transfer and Michaelis complex formation, with the further possibility that an “active site” only exists during the conformational transition. The driving force for the whole process could arise through a distortion of the protein induced by substrate binding (187). The serine group which becomes phosphorylated does not appear to possess any marked nucleophilic reactivity, nor is there any evidence that a histidine group participates as a general acid-general base catalyst. Rate constants for the nonenzymic hydrolysis of alkyl and aryl phosto phate monoanions a t 25” are in the range sec-l (167),while the comparable alkaline phosphatasc-catalyzed values (in this case they refer to dianions) are in the range lo2 to lo3 sec-’. Thus one has to account for a rate enhancement factor of lo0 to 10l2. Moreover, the 186. D. R. Trentham and H. Gutfreund, BJ 106, 455 (1968). 187. D. E. Koshland and K. E. Neet, Ann. Rev. Biochem. 37, 359 (1968); W. P. Jencks, “Catalysis in Chemistry and Enzymology,” p. 282. McGraw-Hill, New York,

1969.

446

H. N . FERNLEY

former values are related to the pK values of the leaving groups, while the latter are not. The mechanism of nonenzymic hydrolysis is thought to involve an unstable metaphosphate intermediate with elimination of phenol rather than phenoxide ion (167).An alternative proposal, involving the formation of a pentavalent intermediate (188), may be more important in relation to the enzyme-catalyzed reaction. This is depicted in (6) where A is an unspecified group or groups allowing proton transfer

x:

R

H

across the active site and the two wedges denote phosphoryl binding groups. The transition state would be favored if the conformationally preferred phosphoryl to serine P - - 0 distance were intermediate between the covalent bond length of 1 . 6 A (189) and the van der Waals separation of 3.3 A (190). With regard to the acceptor site several points might be made. ( 1 ) It is difficult to visualize a site which could cope with the enormous range of potential acceptors whose only common feature appears to be the ability to form a hydrogen bond. (2) There is some evidence of discrimination against water as an acceptor in that rate of transfer to glycerol or glucose compared with that to water greatly exceeds the mole ratio, which may mean that the protein surface a t the acceptor site is not very hydrophilic. (3) The site should also be involved in the forward reaction (by the principle of microscopic reversibility) . An interesting phenomenon in this connection is the stereoselectivity observed in the preferential hydrolysis of L (+) -phosphomandelate by liver and kidney alkaline phosphatases (191). It is not stated whether the selectivity originates in the binding or in the rate of hydrolysis, but whatever the mechanism it seems there must be a direct interaction between mandelate and the enzyme.

Finally there is the general question, so far unanswered, of why differ188. J. 0.Chanley and E. Feageson, JACS 77, 4002 (1955); E. B. Herr and D. E. Koshland, BBA 25, 219 (1957). 189. G. H. McCallum, J. M. Robertson, and G. A. Sim, Nature 184, 1863 (1959). 190. L. Pauling, “The Nature of the Chemical Bond,” 2nd ed., p. 189. Cornell Univ. Preas, Ithaca, New York, 1948. 191. E. Bamann and P. Schwarze, 2.Physiol. Chem. 349, 192 (1968).

18.


4.47

ences in free energy of hydrolysis are reflected more in K, than in Vmax values (there are exceptions to this generalization, notably Pi and PP,, where binding is good yet hydrolysis is slow). For the group of closely related aryl esters studied by King and colleagues (155) there probably is a good correlation between affinity and the pK of the phosphate ester. However, this is not the usual case and it may be concluded that other modes of interaction are involved in the binding, the contribution from each mode depending upon the nature of the particular R and X groups.


Acid Phosphatases VINCENT P . HOLLANDER I . Introduction . . . . . . . . . . A . Distribution . . . . . . . . . B . Historical . . . . . . . . . C . Specificity of Acid and Alkaline Phosphatase . D . Problems with Assay . . . . . . E . Electrophoretic Behavior of Acid Phosphatases I1. Prostatic Acid Phosphatase . . . . . . . A . General . . . . . . . . . . B . Assay . . . . . . . . . . C . Kinetics . . . . . . . . . . D . Preparation . . . . . . . . . E . Electrophoresis . . . . . . . . F . Functional Groups and Effect of Group Reagents G . Transphosphorylation . . . . . . H . Use as a Reagent for Structural Studies . . I . Physical Properties . . . . . . . I11. Red Cell Acid Phosphatase . . . . . . . A . General Properties . . . . . . . B . Purification and Separation of Genetic Types . IV . Liver Acid Phosphatase . . . . . . . . A . Rat Liver . . . . . . . . . B . Mouse Liver . . . . . . . . C . Bovine Liver . . . . . . . . V . Spleen Acid Phosphatase . . . . . . . VI . Acid Phosphatnse in Serum . . . . . . . VII . Miscellaneous Sources . . . . . . . . A . Gaucher Acid Phosphatase . . . . . B . Bone Acid Phosphatase . . . . . . C . Plant Acid Phosphatase . . . . . . D . Neurospora cms'sa . . . . . . . E . Saccharomyces Phosphatase . . . . . 449

. . . . . . . . .

. . . . . . . . .

. . . . . .

. . .

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

. . . . . . . . .

. . . . . . . . .

. . . . . . . .

. . . . . . . .

. . . . . . . .

. .

. .

. .

450 450 450 450 454 454 455 455 457 457 466 468 469 472 473 476 477 477 477 484 484 489 491 493 495 496 496 496 497 497 497

450

VINCENT P. HOLLANDER

F. Staphylococcal Acid Phosphatase G. Amebic Phosphatase . . . H . E . coli Acid Phosphatase . . I. Melanogaster . . . . .

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

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

498 498 498 498

1. Introduction

A. DISTRIBUTION Acid phosphatase or orthophosphoric monoester phosphohydrolase

(EC 3.1 3.2) activity is widespread throughout nature. Hydrolysis of a variety of orthophosphate esters as well as transphosphorylation reactions are catalyzed by enzymes from many sources. Table I illustrates their ubiquitous nature.

B. HISTORICAL I n 1924, Martland et al. (1) reported on phosphatase activity in red blood cells. Roche later differentiated between the phosphatase of the red cells with pH optimum 6.0-6.2 and the phosphatase from white cells with optimum 8.8-9.0. Roche also showed that a-glycerophosphate was split more rapidly than P-glycerophosphate by red cell extracts while the reverse was true of acid phosphatase activity in plasma ( 2 ) . While studying the source of acid phosphatase activity in male urine, Kutscher and Wolberg discovered the very high activity of acid phosphatase in human prostate (3).This tissue was shown by Woodard to have onethousand times the activity of extracts from bone, liver, and kidney (3a). Igarashi and Hollander crystallized the acid phosphatase of rat liver and showed that under certain conditions allosteric control of the activity could be demonstrated ( 4 ) .

C. SPECIFICITY OF ACIDAND ALKALINE PHOSPHATASE Historically, the difference between acid and alkaline phosphatase rested on the simple observation that enzymes could be separated which had optimal rates in completely different p H ranges. Since addition or 1. M. Martland, F. S. Hansman, and R. Robison, BJ 18, 1152 (1924). 2. M. J. Roche, Bull. SOC.Chim. B w l . 13,841 (1931).

3. W. Kutscher and H. Wolberg, 2.Physiol. Chem. 236, 237 (1935). 3a. H. Q. Woodard, Cancer Res. 2, 497 (1942). 4. M. Igarashi and V. P. Hollander, JBC 243, 6084 (1968).

19.

451

ACID PHOSPHATASES

TABLE I SITES OF DISTRIBUTION OF NONSPECIFIC ACID PHOSPHOMONOESTERASES Distribution

Referen ces Nonanimal

Molds

Uzawa (6) Bamann and Meisenheimer (6)

Yeasts

Albers and Albers (7) Luers and Malseh (8) Schaeffner and Krumey (9)

Seeds (almonds, barley)

WaldschmidbLeitz arid Mayer (10) Joyce and Grisolia (11)

Citrus fruits Bacteria (staphylococcal, E. mli)

Axelrod ( I d ) Malveaux and San Clemente (Ida) Rothschild (I%, 1%) Animal.

Prostate gland (man and monkey)

Kutscher and Wolberg (3) Kutscher and Pany ( I S ) Siebert et al. (14)

Kidney

Perlmann and Ferry (16)

Liver

Bamann and Diederichs (16)

Spleen

Bamanri and Diederichs (17)

Erythrocytes

Abul-Fad1 and King (18)

Blood plasma

Fishman and Davidson (19)

Minute amounts of the acid phosphomonoest>eraseshave also been found to occur in the pancreas, in skeletal and heart muscle, and in the mucosa of the small intestines. 5. S. Uzawa, J . Bwchem. ( T o k y o ) 15, 19 (1932). 6. E. Bamann and M. Meisenheimer, Biochem. 2. 287, 380 (1936). 7. H. Albers and E. Albers, 2.Physwl. Chem. 234, 47 (1935). 8. H. Luers and L. Malseh, Wochschr. B T ~ u46, . 143 (1929). 9. A. Schaeffner and F. Krumey, 2.Physiol. Chem. 255, 145 (1938). 10. E. Waldschmidt-Leitz and K. Mayer, 2.Physiol. Chem. 236, 168 (1935). 11. B. K. Joyce and S. Grisolia, JBC 235, 2278 (1960). 12. B. Axelrod, JBC 167, 57 (1947). 12a. F. J . Malveaux and C. L. San Clemente, J. Bacteriol. 97, 1209 (1969). 12b. J. Rothschild, Compt. Rend. l'mv. Lab., Ser. Chim. 35, 457 (1965-1967). 12c. J. Rothschild, Compt. Rend. Trav. Lab., Ser. Chim. 35, 391 (1966). 13. W. Kutscher and J. Pany, 2. Physiol. Chem. 255, 169 (1938). 14. G. Siebert, G. Jung, and K. Lang, Biochem. 2.326, 464 (1955). 15. G. E. Perlmann and R. Ferry, JBC 142, 513 (1942). 16. E. Bamann and K. Diederichs, Chem. Ber. 68, 6 (1935). 17. E. Bamann and K . Diederichs, Chem. Ber. 67, 2019 (1934). 18. M. A. M . Abul-Fad1 and E. J. King, BJ 45, 51 (1949). 19. W. H. Fishman and H. M. Davidson, Methods Bwchem. Anal. 4, 257 (1957).

452


deletion of various peptide groups with alteration of a common enzymic core might be expected to alter the effect of p H on optimal rate, the enzymes should be examined for more fundamental differences in reaction mechanism. Table I1 (20) shows that alkaline phosphatases hydrolyze S-substituted monoesters of phosphorothioic acid (RSPO, Naz; R = -CH, CH, NH,, -CH, COO-, etc.), and 0-substituted monoesters of orthophosphoric acid. Table I11 (20) shows that the K , and maximal velocities of hydrolysis for these substrates are the same. The hydrolysis of the 0-substituted monoesters of phosphorothioic acid (ROPO, SKH ; R = -CHa,-nitrophenyl) not only does not occur but also O-p-nitrophenol thiophosphate was a potent inhibitor of alkaline phosphatase. Acid phosphatase (wheat germ, potato, and prostate) did not hydrolyze S-substituted monoesters of phosphorothioic acid (Table 11) but did hydrolyze 0-substituted monoesters of phosphorothioic acid under identical conditions. The similarities in rates a t which S-substituted monoesters of phosphorobhioic acid and of 0-substituted monoesters of TABLE I1 HYDROLYSIS PRODUCTS OF VARIOUS 0- AND 8-SUBSTITUTED MONOESTERS OF PHOSPHOROTHIOIC ACIDBY ALKALINEAND ACID PHOSPHATASESQ Substrate Cysteamine 8-phosphate N-Acetylcysteamine 8phosphate 8-(Carboxymethyl) phosphorothioate 8-[2-(Methoxycarboxyl)ethyl] phosphorothioate 0-Methyl phosphorothioate

Alkaline phosphatase, products identifiedb

Acid phosphatase, products identifiedc

Cysteamine, 3aPi N-Acetylcysteamine,

No cleavage No cleavage

32Pi

d, azPi

No cleavage

a, 3ZPj

No cleavage

No cleavage

36S-Phosphorothioate, d 36SPhosphorothioate, d W-Phosphorothioate, p-nitrophenol Pi, p-nitrophenol

0-Ethyl phosphorothioate

No cleavage

0-p-Nitrophenyl thiophosphate

No cleavage

p-Nitrophenyl phosphate

p-Nitrophenol,

Pi

From Neumann (90). Identical products were obtained with alkaline phosphatases from the various sources. c Identical products were obtained with acid phosphatases from the various sources. The alcohol liberated was not identified. 0

b

20.

H. Neumann, JBC 243, 4671 (1968).

19.

453

ACID PHOSPHATASES

TABLE I11 ENZYMIC HYDROLYSIS OF THREE TYPESOF SUBSTRATES BY ALKALINE AND ACID PHOBPHATASEB" Cleavage shown by high voltage paper, electrophoresis [Ref. (S)]

Enzyme and substrate Alkaline phosphatase (E. coli) Cysteamine S-phosphate p-Nitrophenyl phosphate p-Nitrophenyl thiophosphate Alkaline phosphatase (intestinal) Cysteamine S-phosphate p-Nitrophenyl phosphate 0-p-Nitrophenyl thiophosphate Acid phosphatase (potato)c Cysteamine S-phosphate p-Nitrophenyl phosphate 0-p-Nitrophenyl thiophosphate

Yea Yes No

4 . 2 X 1W8 5 . 8 X 10-8

9.4 X 10-' 9.4 X lo-'

Yes YeS No

0 . 7 X lov8 1.1 X lW8

2 . 5 X 10-4 2 . 5 X 1k4

5.2 X 3.2 x

2.5 X lO-' 2.2 x 10-4

No Yes Yes

10-10

From Neumann (20).

* Maximum rates of hydrolysis,

V,,,, were expressed as moles of substrate hydrolyzed per ml per min per pg of enzyme per ml of reaction mixture. The reactions were follow'ed by spectrophotometrk determination of the products. Essentially identical results were obtained with acid phosphatase preparations from wheat germ and from bovine prostate gland.

orthophosphoric acid were split by alkaline phosphatase suggests that this enzyme does not bind to either 0 or S linking atoms in these compounds. The lack of hydrolysis of 0-substituted monoesters of phosphorothioic acid by alkaline phosphatase suggests that the two hydroxyl groups are essential for enzymic activity. The inability of acid phosphatase to hydrolyze S-substituted monoesters of phosphorothioic acid suggests that an oxygen linkage is required and cannot be substituted by sulfur. Acid phosphatases are not inhibited by metal complexing agents in contrast to alkaline phosphatases. These results imply that there are basic differences in the two enzymes which transcend consideration of pH optimal rates. Acid phosphatase catalyzes an apparent transition state displacement and P-0 cleavage according to the following: 0-

R-0-A-0II 0

+

HlBOH

-

ROH

0-

+

I H'*O-P-OI1 c,

(1)

One atom of lsO is introduced for every mole of phosphate ion liberated

454


(21). During the nonenzymic acid-catalyzed hydrolysis of phosphomonoesters ( 2 2 ) ,P-0 cleavage also occurs.

D. PROBLEMS WITH ASSAY Two major difficulties must be considered in any assay for acid phosphatase. The enzyme is subject to surface inactivation (23, 2.4). Accordingly, reproducible initial hydrolytic rates are not always obtained, and the kinetic behavior should be checked in any new assay developed. Discrepancies between the amount of inorganic phosphate produced and phenol liberated from phenolic phosphates may be substantial if extensive phosphotransferase activity occurs because of phosphoryl acceptor action on the part of hydroxylic buffers or other constituents of the incubation mixture (25, 2 6 ) . Fluorogenic assays have been developed with very high sensitivity (27). Reference will be made to particular assays in the discussion of the specific enzymes.

E. ELECTROPHORETIC BEHAVIOR OF ACID PHOSPHATASES Beckman et al. (28) have studied the electrophoretic separation of the acid phosphatase activity in tissue extracts on starch gel a t pH 8. They described four electrophoretic bands: A, B, C, and D. Table I V (28) shows the distribution of activity in different organ extracts. The ABD pattern predominated in kidney; BD in liver, intestine, heart, and skeletal muscle; B in skin and D in pancreas. The C component was present in a large number of placentae but not in other adult organs. All four electrophoretic components were inhibited by D- (+)-tartrate; A contained sialic acid, D had a lower pH optimum and was more heat resistant than A, 3,and C. Components C and D showed parallel electrophoretic behavior. In human skin fibroblasts grown in tissue culture, the acid phosphatase was generally high and the most common pattern was BD. Almost every culture showed some activity. The BD 21. S. S. Stein and D. E. Koshland, Jr., ABB 39, 229 (1952). 22. W. W. Butcher and F. H. Westheimer, JACS 77, 2420 (1955). 23. K. K. Tsuboi and P. B. Hudson, ABB 55, 191 (1955). 24. F. Sch@nheyder,BJ 50, 378 (1952). 25. V. N. Nigam and W. H. Fishman, JBC 234, 2394 (1959). 26. H. M. Davidson and W. H. Fishman, JBC 234, 526 (1959). 27. B. Rotman, J. A. Zderic, and M. Edelstein, Proc. N a t l . A m d . Sci. U . S. 50, 1 (1963). 28. L. Beckman, G. Beckman, S. Bergman, and E. Lundgren, Actu Genet. Statist. M e d . 18, 409 (1968).

19.

455

ACID PHOSPHATASES

TABLE IV DISTRIBUTION OF ACID PHOSPHATASE COMPONENTS IN THE TISSUES OF 14 DIFFERENT INDIVIDUALS~~~

Heart Skeletal muscle Kidney Skinc Liver Pancreas4 Intestine

ABD

BD

AB

B

D

1 3 13 0 1 0 1

13 10 1 0 13 1

0 0 0 2 0 0 0

0 0

0 1 0 0 0 13

13

0

12 0 0 0

0

~~~

From L. Beckman, G. Beckman, S. Bergman, and E. Lundgren, Isozyme variations in human cells grown in vitro. 11. Acid phosphatase. Aeta Genet. Statist. Med. 18, 409-415 (1968). Karger, Basel/New York. b Numbers refer to the number of individuals demonstrating a particular type of enzyme. A faint trace of D may be found. d The D component is slightly faster than in other tissues. 0

pattern is the principal activity in most human tissues with the exception of skin, kidney, and pancreas. It is of considerable interest that cultured cells show a different pattern from skin homogenates; D is missing, or present only as a faint trace in skin homogenates from adult individuals. Many cultures from fetal tissues show the C component. The presence of the C component in four out of seven representative cell cultures from adult sources suggests dedifferentiation to a primitive state. Prostate, red cell, and liver acid phosphatase will be discussed in detail because considerable data are available for these enzymes. II. Prostatic Acid Phosphatase

A. GENERAL Study of intermittent effluxes of acid phosphatase activity in the urine of mature human males (99) led to the discovery of the enzyme in semen and prostate by Kutscher and Wolberg (3).The enzyme is very active in human prostatic tissue and the caudal lobe of the rhesus monkey. Dog prostate contains much less enzyme than human tissue. Cat, guinea pig, rabbit, and rat prostates contain little (30).Synthesis 29. A. Dmochowski and D. Assenhajm, Naturwksenschaften 23, 501 (1935). 30. A. B. Gutman and E. B. Gutman, Proc. SOC.Expptl. BWZ. Med. 39, 529 (1938).

TABLE V THE RELATIVEEASE OF HYDROLYSIS OF VARIOUS SUBSTRATES BY PROSTATIC AND ERYTHROCYTIC ACIDPHOSPHATASESO*~ _____

~

~~

Units of acid phosphatase/100 ml Normal range (units/100 ml)

Group

Substrate

1

Phenyl phosphate Phenolphthalein phosphate pNitropheny1 phosphate 8-Naphthyl phosphate &Glycerophosphate a-Naphthyl phosphate

~~

2 3

~

~

Serum plus prostatic acid phosphatase

Serum plus erythrocytic acid phosphatase

19.6 96 2.05 3.3 6.3 32.8

78.6 1120 17.W 15.1 1.3 3.9

Heated serum

Relative specificity for prostatic acid phosphatase

~

1-4 2-10 1-2.3" 0.7-1.6 0-1 0-5

1.9 0

0

0.5

2.3 0.9 1.2 1.9

0

48

0.6

98

.From A. L. Babson, P. A. Read, and G. E. Phillips, Am. J . Clin. Puthol. 32, 83 (1959). Copyright (1959), The Williams & Wilkins Co., Baltimore, Maryland. 8 All activities are in the respective assay units as noted by Babson et al. (36). Units per liter.

3

z H

2 'd

X

8

r. El

19.

ACID PHOSPHATASES

457

is under endocrine control since administration of testosterone to immature rhesus monkeys promptly increased enzymic activity (30).The small amount of acid phosphatase activity in the serum of normal human males derives not from the prostate but largely from the blood platelets (31-33), or through hemolysis of red cells with subsequent release of erythrocyte acid phosphatase (vide injra) . Prostatic acid phosphatase is present in the serum of individuals with prostatic cancer which has invaded the capsule.

B . ASSAY Most investigators utilize p-nitrophenyl or a-naphthyl phosphate as substrate. The determination of serum prostatic acid phosphatase was developed by Fishman and Lerner (34) based on the tartrate inhibition of prostatic enzyme discussed below. Babson et al. (36, 36) demonstrated that a-naphthyl phosphate was much more easily split by prostatic than red cell phosphatase. Table V (36) shows the results obtained when prostatic or red cell phosphatase was added to human serum which had been incubated a t pH 8.6 for 1 hr at 37” to destroy all endogeneous phosphatase activity. The table shows the superiority of a-naphthyl phosphate as substrate. A spectrofluorometric method for the estimation of acid phosphatase has been devised. It uses a-naphthyl phosphate as substrate; thus, it is somewhat more specific for prostatic acid phosphatase than most (37). C. KINETICS 1. Effect of p H and Substrate on Rate

Figure 1 (38)illustrates that maximal hydrolysis of phenyl phosphate occurred a t p H 4.9, 5.0, and 5.0 in acetate, citrate, and tris-HC1 buffers, respectively. For p-nitrophenyl phosphate, the corresponding values were 4.9, 4.7, and 5.5. For /3-glycerophosphate, the values were 5.5, 5.7, and 31. M. B. Zucker and J. Borrelli, J . Clin. Invest. 38, 148 (1959). 32. L. Chevillard, Compt. Rend. Soc. Biol. 139, 249 (1945). 33. T. Mann, “The Biochemistry of Semen and of the Male Reproductive Tract.” Wiley, New York, 1964. 34. W. H. Fishman and F. A. Lerner, JBC 200, 89 (1953).

35. A. L. Babson, P. A. Read, and G. E. Phillips, Am. J . Clin. Pathol. 32, 83 (1959). 36. A. L. Babson and P. A. Read, Am. J . Clin. Pathol. 32, 88 (1959). 37. D. M. Campbell and D. W. Moss, Clin. Chim.Acta 6, 307 (1961). 38. V. N. Nigam, H. M. Davidson, and W. H. Fishman, JBC 234, 1550 (1959).

458


PH

FIQ. 1. Prostatic acid phosphatase activity as a function of pH: ( 0 )phenyl phosphate ; ( 0 )p-nitrophenyl phosphate ; and ( A ) p-glycerophosphate. Buffers: Ac, acetate; Cit, citrate; and tris. From Nigam et al. (38).

a range of 5.0-6.0. A sharp optimum was obtained only with citrate buffer. Table VI (38) shows that there are differences in K,,, depending on the nature of the substrate and buffer. At reasonable experimental rates, hydrolysis was proportional to enzyme concentration although the curve for P-glycerophosphate was slightly concave down. Figure 2 (58) shows that the effect of temperature on reaction rate was quite similar for phenyl and p-nitrophenyl phosphate and altogether different for /I-glycerophosphate. The rate of hydrolysis increased strikingly from methyl to pentanyl TABLE VI EFFECTOF SUBSTRATE A N D BUFFER ON KINETIC CONSTANTS OF ACIDPHOSPHATASEO

a

Buffer

Acetate (MI

Citrate (M)

Phenyl phosphate Nitrophenyl phosphate Glycerophosphate

7 . 5 x 10-4 8.1 X lo-' 4.0x lo-*

9.1 x 10-6 3.1 x 10-4

From Nigam et al. (38).

2 . 0 x 10-3

19.

459

ACID PHOSPHATASES

Temp.

FIG.2. Effect of temperature on hydrolysis. Symbols are explained in Fig. 1. From Nigam e t al. (38).

phosphate and decreased with further lengthening of the carbon chain (39)*

2. Surface Inactivation

Solutions of acid phosphatase are particularly sensitive to surface inactivation. Figure 3 (23) shows the inactivation rate of the enzyme in the presence and absence of surface-active detergents. The inactivation process is temperature sensitive and the protection by detergent is total. Most of the enzyme inactivation proceeds with first-order kinetics. A variety of agents-gelatin, bovine serum albumin, egg albumin, and Tween-80-protect the enzyme against inactivation.

3. Fluoride Inhibition Reiner and his colleagues (40) demonstrated that fluoride inhibition of prostatic acid phosphatase showed interesting and unexpectedly complex kinetics. The unusual nature of the inhibition can readily be appreciated from Fig. 4 (40). As the fluoride concentration is increased over a 1000-fold range, the extent of inhibition rises and then subsequently falls with a further increase of inhibitor. At lower fluoride concentration, the inhibition is clearly competitive. Two possibilities were explored for an explanation of these unusual concentration effects of inhibition. There could be two forms of fluoride in the reaction mixtures; the inhibitory form and the second which predominates a t higher 39. S. J. Thannhauser and G. Schmidt, Federation Proc. 19, 332 (1960). 40. J. M.Reiner, K. K. Tsuboi, and P. B. Hudson, ABB 56, 165 (1955).

460

VINCENT P. HOLLANDER m

.c .0

$ 0.0y . ; .-> L

e

u

0 0 .-g

1.8 -

1.6

j 'c

0 m

-

-

14-

-0

---

"\,

0

h,

Kz0.14 min-I

K~0.33min-1 3

FIG. 3. Surface inactivation rate of prostatic acid phosphatase by shaking and protection by added surface-active agent. Shaking mixtures (20 ml) contained purified enzyme (0.36 pg of protein/ml) in 0.05 M acetate buffer at p H 5.5. The solutions were shaken in 50 ml volumetric flasks using a mechanical shaker (Burrell, model C C ) . Temperatures were maintained by immersion of the flasks in an appropriately set water bath. After specified intervals of shaking, duplicate 0.1 ml aliquots were removed into tubes containing Triton X-100.All tubes were a w y e d simultaneously, following the shaking procedure, with 0.05 M phenyl phosphate as substrate. Curve 1: Enzyme Triton X-100at 0°C and 29°C. Curve 2: Enzyme alone a t 0°C. Curve 3: Enzyme alone at 29°C. From Tsuboi and Hudson (23).

+

-log ( F 1

FIG.4. Relationship between fluoride concentration and enzyme inhibition. Renction mixtures contained in addition to substrate and fluoride, 0.1 M acetate, and 40-fold purified enzyme (in 0.0170 gelatin), all at pH 5.5 in a 1.0-ml reaction volume. Points designated by triangles and plus symbols (+) are calculated from theory. Curve 1: p-Glycerol-PO, (13 M ) Curve 2 : Yeast adenylic acid (0.044 M ) Curve 3 : Phenyl-PO, (0.14 M ) From Reiner et al. (40).

.

.

.

19. ACID

461

PHOSPHATASES

concentration and does not combine with the enzyme. On the other hand, two forms of fluoride could exist, one of which also predominates a t higher concentration and protects the active site from combination with the first or inhibitory form. The concentration dependence suggested that the second form was a polymer, and this was consistent with the well-known properties of fluoride. The first hypothesis is not really consistent with the observed maximum of fluoride concentration. Increase in total fluoride would result in an increase in the hypothetical inhibitory form. The inhibition would then steadily increase with fluoride concentration and a maximal degree of inhibition would not be found. Kinetic equations were developed to support the second hypothesis based on the underlying concept that substrate could displace the second polymeric form from the enzyme surface and thus relieve the degree of inhibition. Vescia and Chance (41) demonstrated that fluoride and tartrate inhibition (vide infra) of acid phosphatase showed completely different kinetics when the hydrolysis of phenyl phosphate was compared with transphosphorylation from this substrate to glucose. Figures 5 and 6 (41) show that fluoride inhibition is competitive when the data are plotted according to Lineweaver and Burk. However, the inhibition is noncompetitive with respect to transphosphorylation of the same substrate to glucose. The authors suggested that there are two distinct sites NaF 1/500 M

15-

NaF 1/1000M

> -

\

NoF V20W M

0

500

1000

I /s

FIQ.5. Action of different concentrations of NaF on phenol (pmole) liberated by human prostatic phosphatase in the presence of different amounts of phenyl phosphate. From Vescia and Chance (41).

41. A. Vescia and E.

K. Chance, BBA 30, 446 (1958).

462


100

-

I/S

F’Io. 0. Action of different concentrations of NaF on G-6-P (pmole) formation by human prostatic phosphatase in the presence of different amounts of phenyl phosphate and excem glucose. From Vescia and Chance (41).

on the enzyme surface for glucose or water binding. Fluoride and tartrate compete for the water site, but glucose must be bound a t a different, but close site. The inhibitors presumably cannot compete for substrate since one would expect both hydrolysis and transphosphorylation to show competitive kinetics under such circumstances. 4. Inhibition b y a-Hydroxycurboxylic Acid

Kilsheimer and Axelrod (4.2) showed that D- (+)-tartaric acid (4.2~~) , but neither meso- nor L-(-)-tartaric acid, was a potent inhibitor of prostatic acid phosphatase. This observation has been confirmed and extended by a number of investigators. The stereospecificity cannot be explained by any structural similarity to obvious substrates. Table VII (4.3) shows the inhibition of prostatic acid phosphatase by a variety of hydroxycarboxylic acids. The values in the legend represent the concentration of inhibitor giving 50% inhibition with 0.0067 M p-nitrophenyl phosphate as substrate. The effective inhibitors possess a hydroxyl group of the D configuration in the a position. If a SH group is substituted for the a-hydroxyl group, as in D,L-thiomalic acid, no inhibition occurs. Amino acids are not inhibitory. A p-hydroxy group alone (phydroxybutyric acid) is not inhibitory. The basis for inhibition is not B. Axelrod, JBC 227, 879 (1957). 42a. The literature is conflicting on the nomenclature of the steric forms of tartaric acid. I n this article, D-(+)- or merely (+I tartaric acid will designate the dextrorotatory form ; L-(-)- designates the levorotatory enantiomorph. Tables and discussion of articles have been altered to conform to this. The reader can avoid confusion by ascertaining the rotation of a described inhibitor. 42. G. S. Kilsheimer and

19. ACID

463

PHOSPHATASES

TABLE V I I INHIBITION OF PROSTATIC ACID PHOSPHATA~E BY VARIOUSHYDROXYCARBOXYLIC ACID AND RELATEDCOMPOUNDS"

Inhibitor

Concentratio for 50% inhibition (MI 0.001& 0 . 00265c

(+)-Tartaric acid Monoethyl ester of (+)-tartaric acid D-( +)-Glyceric acid b(+)-Threonic acid D,kGlyceric acid Diamide of (+)-tartaric acidd mesa-Tartaric acid Dihydroxytartaric acid Tartronic acid

0 . 00325c 0.0131 0 .O14lc 0.0172c 0.044% 0.055& 0.0608

D-Malic acid (unnatural)

0 . O76lc

L(-)-Arabonic acid

0.088P

Diethyl ester of (+)-tartaric acid Oxalic acid Hydroxypyruvic acid Ketomalonic acid Ammonium n-gluconate D-(+)-Xylonic acid 2,4-Dihydroxybenzoic acid D,kMalic acid D-Glucono-S-lactone D-( +)-Ribonic acid 3,5-Dihydroxybenzoic acid Malonic acid D-Glucuronolactone D-Saccharic acid D-Galacturonic acid

0.105 0.107 0.111 0.115 0.125 0.129 0.152 0.167 0.177' 0,324 0.420 0.420 0.516 0.543 0.545

Inhibitor D-(+)-Arabonic acid trans-Aconitic acid cis-Aconitic acid D,L-Aspartic acid Benzoic acid Citric acid L-Cysteine L-Glutamic acid Glycolic acid D,L-B-Hydroxybutyric acid m-Hydroxybenzoic acid p-Hydroxybenzoic acid a-Hydroxyisobutyric acid Isocitric acid D-(-)-Lactic acid L-(+)-Lactic acid D,bLactic acid LMalic acid (natural) Mercaptosuccinic acid Mucic acid D-Phosphoglyceric acid Pyruvic acid Salicylic acid D, L-Serine Succinic acid (-)-Tartaric acid D, L-Threonine

Concentration for 50% inhibition (M)b

No effect No effect No effect No effect No effect No effect No effect No effect No effect No effect No effect

No effect No effect No effect No effect No effect No effect No effect No effect No effect No effect No effect

No effect No effect No effect No effect No effect No effect

The reaction conditions are those described for the standard assay. Calculations were made by using Hunter and Downs' modification of the Michaelis-Menten equation. bThe inhibitors were present a t a concentration of 0.089M in the experiments in which no effect was recorded. c The values are based on six or more concentrations of inhibitor. d The tartramide may have contained up to 30% of the monoamide [Kilsheimer and Axelrod (&')I. (1

464


the intermediate phosphorylation of the a-hydroxyl group to form the actual inhibitor. No induction period was observed to allow for the formation of such an inhibitor. The inhibition of D-glyceric acid was lost upon phosphorylation of the /I-hydroxyl group. Kilsheimer and Axelrod found some small inhibitory power by the meso form of tartrate in contrast to the results of Abul-Fad1 and King. However, the minor inhibition observed could result from slight contamination of the meso with the /I form. On the other hand, meso-tartaric acid might actually be a weak inhibitor since one of the asymmetric carbon atoms has the required D configuration, although the other does not. The P-carbon must be part of a carboxyl group or be attached to a carboxyl or hydroxyl group. The carboxyl group is active when free, esterified, or amidated. The activity of monoethyl ester of D-tartrate is not a result of hydrolysis during reaction to the free acid since ester concentration determined by the hydroxamic method does not change during the reaction. When different substrates for prostatic acid phosphatase are used, the dissociation constant of the enzyme-inhibitor complex Ki is identical in spite of widely different K,,, values for substrate. Figure 7 (42) shows the linear relationship obtained when the data are plotted according to the method of Hunter and Downs ( 4 3 ) . The different substrates have the 1

1

1

1

1

1

1

I P-Nitrophenyl phosphate

S ( M Xlo3)

FIG.7. Inhibition of prostatic acid phosphatase by D-(+)-tartaric acid. The reaction mixtures all contained equivalent amounts of the enzyme preparation, the indicated concentration of substrate (pH 5.0), 0.05 M acetate buffer ( p H 5.01, and tartaric acid (pH 5.0) ; total volume, 4.5 ml. Each point represents average values of determinations made with 5 X lO-'M and 10 X 10dM tartaric acid except in the M and 2 X 10- M tartaric acid was case of p-glycerophosphate for which 1 X used. From Kilsheimer and Axelrod (48). 43. A. Hunter and C. E. Downs, JBC 157, 427 (1945).

19.

465

ACID PHOSPHATASES

same Ki. Table VIII (42) shows the relationship between p H and tartrate inhibition. The concentration of inhibitor, C , required to produce 50% inhibition is given by the convenient expression of Hunter and Downs ( 4 3 ) .

+

C = K i S X KJKm (2) It is noted that the effectiveness of the inhibitor increases with increasing pH, although little change occurs in the region of p H 4.0-5.5. The amount of inhibition increases with increasing pH in spite of an increasing dissociation of the enzyme-inhibitor complex because K,,, increases more rapidly than K i with pH. Kilsheimer and Axelrod have shown that the stereospecific inhibition by D- (+)-tartrate of certain acid phosphatases is widespread. I n a wide phylogenetic study, they found that “unequivocal” plants lack and “unequivocal” animals possess phosphatases inhibitable by D-tartrate. This rule seems to hold even though all animal acid phosphatases were not inhibited (44). TABLE VIII INHIBITION OF p-NITROPHENYL PHOSPHATE HYDROLYSIS BY (+)-TARTARIC ACID AND

K,

OF

p-NITROPHENYL PHOSPHATE

M

3.05 3.30 3.68 4.00 4.13 4.48 4.68 5.00 5.48 5.76 6.10 6.40 6.70

3.3 2.8 1.7 1.4 1.6 1.6 1.6 1.6 1.6 1.4 0.99 0.72 0.62

x

AT VARIOUS

pH

VALUES4*b

108

0.015 0.037 0.048

0.0073 0.015 0.012

3.3 2.8 1.7

0.023 0.025 0.033 0.034 0.047 0.075 0.175 0.325 0.726

0.0056 0.0058 0.0079 0.0085 0.0109 0.0154

1.6 1.6 1.6 1.7 1.6 1.4 0.99 0.69 0.57

0.0254 0.034 0.061

From Kilsheimer and Axelrod (42). Standard assay conditions were used except that pH was varied by means of acetate buffers; C values (the concentration required for 50% inhibition) were calculated according to Hunter and Downs and are averages of determinations made with 0.0005 M , 0.001 M , and 0.002 M (+)-tartaric acid a t a substrate concentration of 0.0067 M . a

44. G. S. Kilsheimer and B. Axelrod, Nature 182, 1733 (1958).

466


5. Effect of Ions Prostatic acid phosphatase is partially and reversibly inactivated by calcium ion (&). Anions such as chloride, bromide, and thiocyanate inhibit prostatic acid phosphatase competitively with regard to substrate as well as noncompetitively. A kinetic analysis by London et al. (46) indicates that the noncompetitive inhibition was related to changes in charge on the protein molecule. A variety of nonspecific anions accelerate thermal denaturation of the enzyme. The enzyme is quite sensitive to a number of electrolyte changes, but it is not clear whether these factors are involved in biological control.

D. PREPARATION London et al. (4?', 48) purified prostatic acid phosphatase by extraction of the frozen sliced gland followed by calcium phosphate or Fuller's earth adsorption with subsequent acetone and ammonium sulfate fractionation. Such fractions could be further purified by bubbling CO, into the dilute protein solution. The froth produced by this procedure contained a large amount of protein and very little acid phosphatase activity. Combinations of these purification procedures led to considerable purification. Chromatography on Amberlite XE-69 also proved an effective technique to purify ammonium sulfate fractionated material. Addition of ammonium sulfate to solutions of this purified protein kept a t -5" resulted in the formation of a partially crystalline solid phase. However, identity of enzymic activity and the crystalline product could not be proved and there was considerable loss of enzymic activity following formation of such precipitates. Davidson and Fishman (26) described a simple purification procedure leading to a stable preparation of high activity. Frozen human prostates were extracted in a Waring blendor with tris-citrate buffer. It was found that the enzyme was particularly stable in this buffer a t high pH in the presence of ammonium sulfate. Salt fractionation readily led to highly active preparations. Dowex-50 chromatography also affords considerable purification (49). Since acid phosphatase has proved to be useful in structural studies on phosphoproteins and polynucleotides, Ostrowski and Tsugita (60) found it neces45. A. Steens-Lievens and H. J. Tagnon, Nature 195, 400 (1962). 46. M. London, R. McHugh, and P. B. Hudson, J . Gen. Physiol. 46, 57 (1962). 47. M. London and P. B. Hudson, ABB 46, 141 (1953). 48. M. London, A. Sommer, and P. B. Hudson, JBC 216, 81 (1955). 49. H. G. Boman, BBA 16, 245 (1955). 50. W. Ostrowski and A. Tsugita, ABB 94, 68 (1961).

19.

ACID PHOSPHATASES

467

sary to remove diesterase from preparations of purified enzyme. Ammonium sulfate purified material was chromatographed on DEAEcellulose. The enzyme preparation gave a sharply defined homogenous band on electrophoresis in agar and starch gel separations; analytical ultracentrifugation gave szo = 5.5. This preparation not only lacked diesterase activity but also was free of marked absorption a t 260 nm which had characterized previous preparations. Further purification of DEAE-cellulose material could be achieved by chromatography on CMcellulose. Elution by pH gradient gave two peaks of active material ( 5 1 ) . The main fraction was almost entirely homogeneous by free boundary electrophoresis although a small asymmetry a t the cathode side was evident a t pH 5. This preparation had a molecular weight by Sephadex chromatography of 109,000, sz0 = 5.8. Ostrowski (52) described a method for bulk purification of acid phosphatase. Frozen human prostate glands were extracted with 0.01% Tween 80. Ammonium sulfate fractionation followed by chromatography on Sephadex G-100 and DEAE-cellulose and refiltration through Sephadex G-100 gave a 15% yield and 100-fold purification with respect to the crude extracted tissue. The purified monoesterase is stable for years at pH 6.0 when stored a t -25". It loses more than 90% of activity in 3 M urea a t 25", pH 7.0, and dialysis of the enzyme leads to only 15% restoration of activity. The product is homogeneous on chromatography on DEAE, Sephadex G-100, as well as by free-boundary electrophoresis, disc electrophoresis, immunoelectrophoretic analysis, and precipitin reactions, The purified enzyme was free of phosphodiesterase activity. By sucrose gradient chromatography, s20,w was 5.7, and molecular weight 89,100. The method used for separation from contaminating phosphodiesterase is much more rapid than by means of Dowex 50-X2. The immunochemical method for establishing purity of the enzyme is particularly applicable here because the enzyme-antibody complex retains full activity. Highly purified prostatic acid phosphatase labeled with 14C has been obtained by incubation of slices of hypertrophic human gland with labeled amino acids (63). Lavallee and Rosenkrantz (54) studied the purification of dog prostatic acid phosphatase from prostatic secretion obtained from pilocarpine-stimulated dogs with cystopreputiostomy prostatic fistulas. A 4551. 52. 53. 54.

W. Ostrowski and J. Rybarskri, BBA 105, 196 (1965). W. Ostrowski, Acta Biochim. Polon. 15, 213 (1968). J. Rybarska and W. Ostrowski, Actu Biochim. Polon. 13, 145 (1966). W. F. Lavallee and H. Rosenkrantz, A B B 112, 381 (1965).

468


fold purification was achieved by adsorption of other proteins on DEAE-Sephadex.

E. ELECTROPHORESIS Mattila (55) separated acid phosphatase in prostatic extracts by agar gel electrophoresis. He obtained a broad heavy band a t the aglobulin region and a narrow band a t the p-globulin area. By immunodiffusion, two precipitin lines were found with acid phosphatase activity when prostatic tissue extracts were reacted against antiprostate serum. By immunoelectrophoresis, two precipitin lines in the a- and P-globulin regions also showed acid phosphatase activity. Separation of the two molecular forms of the enzyme indicated by these studies was achieved by fractionation of prostatic extracts on Sephadex G-200. Figure 8 (65) shows the enzymic activity separated into easily separable peaks which correspond to the immunoelectrophoretic fractions seen on examination of crude prostatic extracts. The two fractions had molecular weights of

Fractions at 6 ml

FIG.8. Sephadex G-200 gel filtration diagram of the acid phosphatase activity. There are two peaks, corresponding to the two immunologically different molecular forms of the enzyme. From S. Mattila, Invest. Urol. 6, 337 (1969). Copyright (1969), The Williams & Wilkins Co., Baltimore, Maryland. 55. S. Mattila, Invest. Urol. 6, 337 (1969)

19. ACID

PHOSPHATASES

469

470,000 and 84,000 with 80% of the activity in the latter range. It is probable that the fraction of acid phosphatase studied by Ostrowski and Rybarska (51) with molecular weight 109,OOO represents the major component similar to the 84,000 molecular weight product in the Mattila study. However, the enzyme seems to be much more heterogeneous than this work would indicate. Kaschnitz (56) separated four isoenzymes by disc electrophoresis of prostatic extracts. Sur et al. (57, 58) demonstrated that extracts of the human prostate gland showed acid phosphatase activity in a t least 13 active zones on starch electrophoresis a t p H 6.2. The acid phosphatase activity of human semen moved under similar conditions in a broad band slightly ahead, but overlapping the fastest region of prostatic phosphatase. Treatment of the prostatic extracts with butanol, protease, ethylenediaminetetraacetate (EDTA) , and other similar agents failed to decrease the apparent heterogeneity.

F. FUNCTIONAL GROUPS AND EFFECT OF GROUPREAGENTS 1. Sulfhydryl Groups

Prostatic acid phosphatase is reversibly inactivated by p-mercuribenzoate and by Cuz+and FeS+ ( 5 9 ) . I n contrast to red cell acid phosphatase, prostatic acid phosphatase is only partially inactivated even after prolonged periods of incubation a t high concentrations of p-mercuribenzoate. Addition of cysteine to the p-mercuribenzoate-treated enzyme produces complete reactivation. Binding of SH groups by p-mercuribenzoate renders the enzyme more labile to thermal denaturation, but no difference is obtained with surface inactivation (23).Similar partial inactivation with Cu2+is also subject to reactivation. 2. Iodinatwn

Prostatic acid phosphatase is irreversibly inhibited by reaction with iodine monochloride a t p H 8.1. Figures 9 and 10 (60)show the effect of concentration of ICl and the time course of the reaction. Very rapid inactivation occurred a t concentrations of 0.05 mM IC1. Further increase in the concentration of the reagent produced further inactivation but 56. Von R.Kaschnitz, Z . Klin. Chem. Klin. Biochem. 5, 126 (1967). 57. B. K. Sur, D. W. Moss, and E. J. King, Proc. Assoc. Clin. Biochem. 2, 11 (1962). 58. B. K. Sur, D. W. Moss, and E. J. King, BJ 84, 55P (1962). 59. E. S. Barron, Advan. Enzymol. 11, 201 (1951). 60. K.Bobrzecka, W. Ostrowski, and J. Rybarska, Acta Biochim. Polon. 15, 369 (1968).

470


- l o g (ICtl(M)

FIG.9. The effect of ICl concentration on the inhibition of phosphomonocsteriisc. The enzyme, 0.03 mg/ml, waa incubated for 3 min a t room temperature with ICI a t pH 8.1. The reaction was stopped by the addition of 30 mg % solution of nlbumin in citrate buffer, pH 5.5 (200 &0.1 ml of the incubation mixture). Then the activity of the enzyme waa measured. From Bobrzecka et al. (60).

only with considerable increase in concentration. Tartrate protected the enzyme against the action of iodine monochloride. The ultraviolet spectrum of the iodinated product suggests that tyrosine was modified in the reaction. The tyrosine groups in prostatic acid phosphatase were

Time ( m i n )

FIG.10. Time course of phosphomonoesterase inactivation by 0.1 mM ICI a t pH 8.1.Other conditions are as described for Fig. 9. From Bobrzecka et al. (60).

19.

ACID PHOSPHATASES

471

easily available to low concentrations of IC1 but not tyrosinase. This is consistent with the virtual absence of spectrophotometric titration of tyrosine groups below pH 10.

3. Reagents for Tyrosine and Tryptophan Purified prostatic acid phosphatase is inhibited by reaction with pcliazobenzenesulfonic acid in a manner consistent with a role for tyrosine in the active center. Carboxymethylation under conditions which would derivatize histidine had little effect on the activity. In contrast, incubation with N-bromosuccinimide or 2-hydroxy-5-nitrobenzyl bromide produced extensive irreversible inhibition consistent with a role for tryptophan in enzymic activity. Tartrate protected the enzyme against the action of both p-diazobenzenesulfonic acid and 2-hydroxy-5-nitrobenzyl bromide. Apparently, both tyrosine and tryptophan are important for the catalytic function of prostatic acid phosphatase. They are also presumably involved in the mechanism of tartrate inhibition. Diisopropylphosphorofluoridate ( D F P ) is an irreversible inhibitor of many esterases and proteases (61). The inhibitor interacts with the enzyme at the substrate site (62). O-Phosphoryl serine is isolated from the liydrolysates of the irreversibly inhibited enzymes ( 6 3 ) .Purified prostatic acid phosphatase is also inhibited by DFP. Because alkaline phosphatase is inhibited, this would be an expected result if it is assumed that the acid monoesterase is similar to the alkaline enzyme in general specificity and behavior as a transferase. The alkaline enzyme affords the expected O-phosphoryl serine upon degradation of the inhibited enzyme. However, prostatic acid phosphatase shows a completely unexpected kind of inhibition with DFP. For a given concentration of DFP, the degree of inhibition increases with time until a characteristic maximum value is obtained. The kinetics are those of a pseudo-first-order reaction. Both substrate and the competitive inhibitor D-tartrate protect the enzyme against D F P inhibition. I n contrast to the action of DFP on proteases and esterases, the inhibition of prostatic acid phosphatase is reversible. Thus, when 5 pg of enzyme is incubated for 2 hr with 1 x M DFP in 10 ml 0.1 M acetate buffer containing 0.01% gelatin a t p H 5, the enzyme loses 97% of its activity. Following dialysis of the reaction mixture for 24 hr almost all of the activity is restored. When DFS2Pis removed by gel filtration, the enzyme activity, although initially lacking 61. H. Greenberg and D. Nachmansohn, JBC 240, 1639 (1965). 62. J. A. Cohen, R. A. Oosterbaan, H. S. Jansz, and F. Berends, J . Cellular Comp. Physiol. 54, Suppl. 1, 231 (1959). 63. D. E. Koshland, Jr., Advan. Enzymol. 22, 52 (1960).

472


after chromatography, is slowly recovered on standing. Less than 0.01 mole of labeled phosphate per mole of enzyme was found in the fractions containing the enzyme. Under similar experimental conditions, a proteolytic enzyme like chymotrypsin would bind 1 mole of phosphorus per mole of enzyme. When DF32P-inhibited acid phosphomonoesterase was denatured and dialyzed, no radioactivity could be detected in the protein. When prostatic acid phosphomonoesterase was incubated with inorganic 3zP,no 0-phosphoryl serine was isolated from the incubation mixtures, in contrast to what is found with alkaline phosphomonoesterase. These results indicate that the inhibition of acid phosphatase by DFP is different from the inhibition of esterases and a variety of proteolytic enzymes. Serine does not seem to be part of the active site of prostatic acid phosphatase. The reversible nature of the inhibition and the slow recovery of activity following complete removal of inhibitor suggest that inhibition depends on a conformational change of the enzyme or on some slow process a t a specific amino acid residue. Further work is needed to resolve the question. Greenberg and Nachmansohn (61) repeated the essential part of these experiments with wheat germ and potato acid phosphatase and obtained similar results.

G. TRANSPHOSPHORYLATION Appleyard (64) noted that addition of ethanol to incubation mixtures of sodium phenolphthalein diphosphate with prostatic extract increased the rate of free phenolphthalein formation. Phosphate ion failed to show a comparable increase, and this discrepancy was attributed to transphosphorylation. Phosphoryl transfer may be effected by prostatic phosphatase t o acceptors other than solvent (65-67). Nigam and Fishman (25) studied phosphoryl transfer under conditions of 6&80% transfer to an acceptor. In the case of 1,4-butanediol, the optimal concentration was 0.8 M . I n this experiment, water molecules outnumbered acceptor molecules by 55/0.8 or 70-fold. I n spite of this, transfer far exceeded hydrolysis. Phosphoryl transfer to aliphatic alcohols can be easily measured when phosphates are used as donor compounds. The difference between alcohol formation from the substrate and phosphate ion production is a measure of the transfer reaction. Table IX (25) shows that four different substrates can transfer phosphoryl to butanediol with high efficiency. Table X (25) shows that aliphatic alcohols are good acceptors 64. J. Appleyard, BJ 42, 596 (1948). 65. B. Axelrod, JBC 172, 1 (1948).

66. H. Green and 0. Meyerhof, JBC 197, 347 (1952). 67. B. Axelrod, Advan. Enzymol. 17, 159 (1956).

19.

473

ACID P H O S P H A T A S E S

TABLE IX DONOR SUBSTRATE SPECIFICITY"-* Phosphate ester of

Organic radical (pmole/ml)

Phosphate (pmolelml)

Ester product (pmole/ml)

Mole transfer

Phenol p-Nitrophenol Glycerol Propanediol

0.224 0.261 0.150 0.155

0.067 0.108 0.024 0.042

0.157 0.153 0.126 0.113

71.0 58.5 84.0 73.0

0

(%I

From Nigam and Fishman (26).

* The incubation digest (7.0 ml, final volume) contained 1 ml of substrate (0.022 M

for phenyl and p-nitrophenyl phosphates, 0.03 M for propanediol phosphate, and 0.02 M for 8-glycerophosphate), 2.5 ml of 0.1 M acetate buffer (pH 5.0 for phenyl and p-nitrophenyl phosphate and pH 5.5 for propanediol and glycerol phosphates), 0.5 ml of 1:500 dilution of stock enzyme solution in dilute albumin, 3.0 ml of 1,4-butanediol solution (1.0 ml of butanediol and 2.0 ml of water) giving a final concentration of 1620 pmoles of butanediol per milliliter of the digest. Incubation time, 30 min. The reaction was stopped by the addition of 3.0 ml of 10% trichloroacetic acid, and suitable aliquots were analyzed for inorganic phosphate, phenol, p-nitrophenol, and glycerol. Propanediol was determined by the method used for glycerol in which 1,2-propanediol provided the reference curve.

of phosphoryl groups liberated from donor substrates. There is a trend toward greater transfer with an increase in the number of carbon atoms in the acceptor alcohol. Branch chain alcohols are less efficient as acceptors than corresponding unbranched alcohols. Glycols with vicinal hydroxyl groups serve as acceptors with 4144% transfer. However, the glycols with hydroxyl separated by one or more carbon atoms (1,3and 1 ,bbutanediols, etc.) were more efficient acceptors than those with vicinal hydroxyls. Much less transfer was observed with cyclic alcohols or benzyl alcohol. Phosphoryl transfer from phenyl phosphate to 1,4butanediol was maximal between pH 4.4-6.0. D- (+)-Tartrate inhibits phosphoryl transfer as well as hydrolysis. There can be little doubt that the phosphoryl transfer is a function of the purified hydrolase. However, Tunis and Chargaff (68) have provided evidence for separation of hydrolytic and phosphoryl transfer activity in a carrot phosphatase. The physiological role, if any, of transphosphorylation has not been established.

H. USE AS

A

REAGENT M)R STRUCTURAL ST~IES

When prostatic acid phosphatase is purified so that it is completely free of diesterase activity, it can be used in a variety of structural 68. M. Tunis and

E. Chargaff, BBA 21, 204 (1956).

TABLE X ACCEPTOR SPECIFICITY~J

Experiment NO.

Acceptor

Methanol Ethanol n-Propanol 12-Butanol Isopropanol Isobutanol Tertiary butanol 8 9

10 11 12

Ethylene glycol (1,hthanediol) Propylene glycol (1,2-propanediol) 2J-Butanediol 1 3-Butanediol 1,4-Biitanediol

Acceptor concentration (pmole/ml)

Phenol liberated (A) (pmolelml)

Aliphatic Alcohols 3530 0.270 2450 0.152 1920 0.087 785 0.141 1870 0.186 770 0.152 765 0.308 Glycols 2570 1960 1660 1600 1620

0.315 0.208 0.208 0.254 0.240

Inorganic phosphate liberated (B) (pmole/ml)

Alkyl phosphate formed (A - B) (pmole/ml)

0.170 0.111 0.032 0.039 0.170 0.079 0.288

0.1 0.05 0.055 0.102 0.016 0.078 0.02

35.0 26.6 43.5 71 . O 8.6

0.145 0.108 0.124 0.117 0.065

0.17 0.10 0.084 0.137 0.175

54.0 48.0 40.5 54.0 72.5

Mole phosphoryl transfer

(%I

51.5 6.0

3

2 3 H

+d

3:

$

E

F

13 14 15 16 17 18

a

l15-Pentanediol 2,5-Hexanediol 1,2,6-Hexanetriol 1,6-Hexanediol Glycerol 1,9-NonanedioI

3200 3400 3600 605 1960 17

0.211 0.135 0.242 0.266 0.288 0.198

19 20

Cyclopentanol Cyclohexanol

Cyclic Alcohols 800 0.020 830 0.108

21

Benzyl alcohol

Aromatic Alcohols 820 0.028

0.040 0.080 0.060 0.063 0.131 0.124

0.17 0.055 0.182 0.204 0.16 0.074

80.0 40.0

*

76.0 77.0 54.5 37.4

l

0.018 0.104

0.002 0.004

10.0 0.9

0.021

0.007

2.8

From Nigam and Fishman (26).

* The incubation digest (7.0 ml) contained 1 ml of

9

0.022 M phenyl phosphate; 2.5 ml of 0.1 M acetate buffer, pH 5.0; 0.5 ml of test enzyme solution; and 3.0 ml of solutions of acceptors giving a final concentration as shown in the third column. Incubation time, 30 min. Digests were inactivated by 3.0 ml of 10% trichloroacetic acid solution and were analyzed for phenol and inorganic phosphate. I n the case of the standard acceptor, 1,4-butanediol, the expected transfer product, l,&butanediol phosphate, was isolated in a yield of 35% from a large-scale experiment. The hydrolysis of this phosphate ester by prostatic acid phosphatase liberated approximately equimolar amounts of 1,4-butanediol and inorganic phosphate.

8 $

+d

?

Eez

476


studies. Terminal phosphate can be removed from protein peptides and nucleotides. Perlmann (69) showed that stepwise dephosphorylation of ovalbumin was possible. Only one of the two phosphorus-containing groups is removed with this enzyme and further dephosphorylation is possible with intestinal phosphatase which then gives a phosphorus-free ovalbumin. Even large molecules like tobacco mosaic virus may be dephosphorylated (70). Dephosphorylation and consequent estimation of chain length of oligonucleotides can be accomplished with nucleic acid breakdown products (71, 72).

I. PHYSICAL PROPERTIES Table XI (73) shows the Stokes radii and frictional ratio obtained by the study of purified acid phosphatase. The preparations show molecular homogeneity during filtration on Sephadex G-100, in the analytical ultracentrifuge, and during immunolectrophoresis. These data obtained by chromatography on Sephadex G-200 indicate that human prostatic acid phosphatase has an effective Stokes radius of 47.1 A and a frictional ratio of 1.56, suggesting considerable molecular asymmetry. TABLE XI MOLECULAR P A R A M E T E R S O F PROSTATIC ACID PHOSPHATASEa Stoke's radius

Frictional ratio

Method used

(A)

(flfo)

Porathb Laurent and Killanderc Ackersd Mean value'

47.0 47.3 47.2 47.1

1.56 1.57 1.56 1.56

From Ostrowski and Wasyl (73). Curve constructed according to J. Porath [Pure A p p l . Chem. 6, 233 (1963)l. Curve constructed according to T. C. Laurent and J. Killander [ J . Chromatog. 14, 317 (1964)l. Curve constructed according to G. K. Ackers [Biochemistry 3, 723 (1964)l. 6 Each number represents a mean value of three determinations. 69. G. E. Perlmann, J . Gen. Physiol. 35, 711 (1951). 70. M. P. Gordon, B. Singer, and H. Fraenkel-Conrat, JBC 235, 1014 (1960). 71. R. Markham and J. D. Smith, BJ 52, 565 (1952). 72. M. Privat de Garilhe, L. Cunningham, U.-R. Laurila, and M. Laskowski, JBC 224, 751 (1957). 73. W. Ostrowski and Z. Wasyl, B B A 181, 479 (1969).

19.

ACID PHOSPHATASEE

477

111. Red Cell Acid Phosphatase

A. GENERALPROPERTIES

In 1924, Martland and Robison (74) demonstrated a rapid rise in inorganic phosphate when laked blood was incubated. They correctly interpreted the reaction as the splitting of endogenous organic phosphate by an enzyme acting in the acid range and different from bone phosphatase. Behrendt (75) showed that erythrocyte enzyme was much more active than serum acid phosphatase in splitting phenyl phosphate. Table XI1 (18) illustrates the differences between human red cell and prostatic enzyme toward three commonly used substrates. Red cell, in contrast to prostatic enzyme, splits a-glycerophosphate more rapidly than the p isomer. It is of interest that the red cell enzyme splits neither a- nor P-naphthyl phosphate ( 7 6 ) .Table XI11 (18) demonstrates that a variety of metal ions show differences in inhibition between red cell and prostatic enzyme. Table XIV (18) shows that arsenate and oxalate inhibit both enzymes to a similar degree but that fluoride demonstrates substantially greater inhibition of prostatic enzyme than that from erythrocytes. However, tartrate fails to inhibit red cell acid phosphatase a t all, and it almost completely inhibits prostatic enzyme. The stereospecificity of the inhibition is discussed in the section of prostatic enzyme. Formaldehyde (0.1%) (77) inhibits red cell acid phosphatase completely but has no effect on the prostatic enzyme.

B. PURIFICATION AND SEPARATION OF GENETICTYPES Preparation of pure red cell phosphatase, and assay of such fractions, is complicated because of the extensive heterogeneity of the enzyme. This was first observed during column chromatography by Angeletti and Gayle (78) . Hopkinson in 1963 (76) investigated the heterogeneity and demonstrated that it was on a genetic basis. He described five distinct red cell acid phosphatase patterns obtained after electrophoresis of hemolyzates in starch gel. Bands of enzymic activity were detected with phenolphthalein diphosphate. Differences in phenotypes were best 74. 75. 76. 77. 78.

M. Martland and R. Robison, BJ 18, 765 (1924). H. Behrendt, Am. J. Clin. Pathol. 19, 167 (1949). D. A. Hopkinson, N. Spencer, and H. Harris, Nature 199, 969 (1963). M. A . M. Abul-Fad1 and E. J. King, J. Clin. Pathol. 1, 80 (1948). P. U.Angeletti and R. Gayle, Blood 20, 51 (1962).

TABLE XI1 RELATIVE RATESOF HYDROLYSIS OF Q- AND B-GLYCEROPHOSPHATES AND PHENYL PHOSPHATE BY THE ACID PHOSPHATASES OF THE PROSTATE AND RED CELLS-~ ___

~

~

Hydrolysis (mg P/30 min/100 ml enzyme solution) 0.02 M 6-Glycerophosphate

Enzyme solution

Red cells (human) Red cells (human) Prostate in normal saline _____

0.005 M Phenyl phosphate

Without Mg

0.003 MMg

0.01 MMg

Without Mg

0.003 MMg

0.01 MMg

Without Mg

0.2 0.2 29

0.25 0.2 29

0.3 0.25 30

11 9.2 28

11 9.2 28

11 -

55 43 53

~

From Abul-Fad and King (18). b Acetate buffer pH 5; 30 min at 37". 0

0.02 M a-Glycerophosphate

0.003 MMg

0.01 MMg

-

48

37

-

50

_

_

EFFECT OF METALLIC IONS ~

~

_

_

ON THE

TABLE XI11 AciD PHOSPHATASES OF PLASMA, PROSTATE, AND RED CELLS’-*

m

~~

tLJ m

Percentage inhibition

Enzyme solution -

Control enzyme without inhibitor (units/100 ml)

(0.01 M )

80

14

47

17

88

10

Calcium

Chromium (0.01 M )

Cobalt (0.01 M )

Manganese (0.01 M ) ~

~~

Red cells (human) Red cells (human) Red cells (rat) Plasma (normal) Prostate in normal saline Seminal fluid Prostate in normal plasma

4.2 50 160

10.0

25 17

0

20 -

40 50 16

8 9 -

0 4 -

0 0 0

15 15

17

Nickel

Zinc

(0.01 M )

(0.001 M )

23

20

~~

-

23

-

10 4

28

0

11

7

21

40 52

I)

4

12

From Abul-Fad1 and King (18). Sodium phenyl phosphate substrate; acetate buffer, pH 5; 30 min at 37”. Activity expressed as “units” (i.e., mg P/30 min/100 ml enzyme solution). a

b

480


TABLE XIV EFFECT OF ACIDRADICALS ON THE ACID PHOSPHATASES OF PLASMA, PROSTATE, AND REDCELLS'.~ ~~

~~

~

Activation or inhibition

Acid radical (0.01 M ) Arsenate Citrate Cyanide Fluoride Formate Oxalate Salicylate Tartrate" Taurogly cocholate

Prostate in normal saline

(90)

(5%)

(%)

(%I

(96)

- 72

-80 +5

-66 +8 12 -96 0 -22 0 -94 - 76

-70 +5

-66

+lo

4-15 - 97

Normal plasma

0 0 -30 0 -30 0 0 0

+8

-8 0 -27 0 0 - 77

+

Prostate in normal plasma

Seminal fluid in normal saline

Human red cells in water

-96 0 -25 0 -95 -85

+6

0

-25

0 - 93 -80

From Abul-Fad1 and King (18). Acetate buffer, pH 5 ; phenyl phosphate substrate; liberated phosphate determined after 30 min a t 37". The figures represent the average percentage of activation (+) or inhibition (-) effected in several experiments. D-(+)-Tartrate. b

demonstrated when tris-succinic acid was used in gel and citric acidsodium hydroxide in the bridge buffer a t p H 6. Figure 11 (79) indicates the electrophoretic pattern obtained from the six most common phenotypes found in human blood. These are designated A, BA, B, CA, CB, and C, and each consists of two or more isozymes. From family studies, Hopkinson et al. (76) demonstrated that these phenotypes could be explained by assuming three allelic autosomal genes called Pa, Pb, and P". Homozygotes would be represented as Pa Pa, Pb Pb, P" P". The homozygotes are represented by two electrophoretic bands of enzymic activity. Heterozygotes demonstrate the expected combination of isozymes. Additional rare phenotypes occur because of the existence of the alleles P' and Pd. Scott (80) purified red cell acid phosphatase of homozygous types A and B by using ammonium sulfate and DEAE-cellulose chromatography. The relative activity of these isozyme preparations was the same when tested with a number of substrates. Type B enzyme showed small kinetic 79. E. R. Giblett, "Genetic Markers in Human Blood," Chapter 11, p. 426 (a). Davis, Philadelphia, Pennsylvania, 1969. 80. E. M. Scott, JBC 241, 3049 (1966).

19.

ACID PHOSPHATASES

481

FIG. 11. Starch gel clectrophoretic patterns of red cell hemolyzates with the six phenotypes representing homozygosity and heterozygosity for the three common genes a t the acid phosphatase locus, P', Pb,and P'. From Giblett (79).

differences from A enzyme, less inhibition by inorganic phosphate, and less dependence on substrate concentration. When DEAE Sephadex column chromatography was used to separate isozymes from B, BA, CA, and CB red cells, three fractions were separated with increasing clectrophoretic mobility (81). Table XV (81) shows that these three isoenzymes E,, El, and E, (for slow, intermediate, and fast electrophoretic mobility) varied with respect to a variety of inhibitors. The intermediate band was not inhibited by fluoride. When isozymes of the five common phenotypes were separated on DEAE, the order of elution was slow C, slow B, slow A, fast A, and fast B. This behavior is presumably a result of differences in net charge (82). Different phenotypes vary in thermal stability. Loss of activity after heating is more rapid in A, BA, and B than in types CA and CB (83).I n spite of the striking differences in thermal stability, the sensitivity to urea and guanidine denaturation is the same for all of the isoenzymes. Phosphotransferase activity studied earlier by Tsuboi and Hudson (84) was studied in further detail by Luffman and Harris (83). Phenyl phosphate was incubated a t pH 6 in an aqueous system to which methanol, glycol, propanol, or ethanol were added as acceptor. Transferase activity was estimated by the difference between inorganic phosphate and free p-nitrophenol production. Transfer of phosphate from substrate t o methanol increased i n a linear fashion with increasing methanol concentration until the reaction mixture consisted of 25% methanol a t which point the transferase 81. M. R. Fenton and K. E. Richardson, ABB 120, 332 (1967). 82. D.A. Hopkinson and H. Harris, Ann. Hum. Genet. 31, 29 (1967). 83. J. E. Luffman and H. Harris, Ann. Hum. Genet. 30, 387 (1967). 84. K.K.Tsuboi and P. B. Hudson, ABB 43, 339 (1953).

482


TABLE XV

EFFECT OF VARIOUS COMPOUNDS ON THE ACTIVITYOF

THE

ACID PHOSPHATASE ISOZYMES~ ~~

% Activityb ~~

Addition None EDTA EDTA MgClr MgClr NaF NaF Formaldehyde Formaldehyde Oxalate

oxalate

L-Tartrate tTartrate

Concentration (M)

3.3 X 10-* 6.6 x 6 . 6 X 10-* 1.3 X lo-’ 1 . 3 X lo-* 2 . 0 x 106.6 X 10-’ 1.3 X 6 . 6 X 10-* 1.3 X lo-’ 6 . 6 X lo-* 1 . 3 X lo-*

E, phosphatase

Ei phosphatase

Ef phosphatase

pH 4.75 pH 5.5 pH 5.25 pH 5.5 pH 5.75 pH 5.5 100 103

54 92 92 97 77 84 68 58 36 106 -

100 99 64

100 101 98 106

73

100 95

72

86 54

54

95 83 104 101

-

-

100

-

88

100 100

-

100 100

-

59

-

83

-

100 -

100 103 100 140 69 98 64 60 28 102 76 113 100

100 100

-

140

-

93

-

53 84 96

-

From Fenton and Richardson (81). Assays were performed at the pH indicated, as described by Fenton and Richardson under section entitled (Methods) with p-nitrophenyl phosphate as substrate. (I

b

rate was maximal and three times the rate of hydrolysis seen in the absence of methanol. No consistent differences were observed between isoeneymes for the ability to catalyze the transferase reaction. With glycerol the degree of transferase observed was greater than with methanol, but the observed difference could easily be attributed to enzyme inactivation by methanol. Figure 12 (86)illustrates the result obtained by Luffman and Harris (83) when red cell acid phosphatase was chromatographed on Biogel P 60. The column was equilibrated with trisphosphate buffer pH 8, and stabilization of the red cell acid phosphatase was achieved by adding 2-mercaptoethanol (0.1%) and Tween 80 (0.08%). Elution was performed with a concave exponential gradient between 0.005 M tris phosphate buffer pH 8, and the same buffer 0.5 M in sodium chloride. The degree of retardation is surprising and consistent with a molecular weight of 7,000-10,000. The heritable nature of red cell acid phosphatase isozymes was discovered with the aid of the high separation power of starch gel chromatography. However, if the physical differences found in the isoeymes 85. D. A. Hopkinson and H. Harris, in “Biochemical Methods in Red Cell Genetics” (J. J. Yunis, ed.), Chapter 13, p. 353. Academic Press, New York, 1969.

19.

ACID PHOSPHATASES

35

45

55

65

75

55

95

105 115

125 135 145

155 165 175

IS5

Tube number

FIG. 12. Diagram of elution pattern of red cell acid phosphatase and various markers on Biogel P 60. The position of the various protein markers was determined both by optical density determination and by starch gel electrophoresis of the individual fractions (83). The experiment was carried out using a polyacrylamide gel (Biogel P 60, 50-150 mesh; exclusion limit >sO,OOO; Bio-Rad Laboratories, California) in 0.05 M tris buffer, pH 8.0, containing 0.08% (v/v) Tween 80 and 0.1% (v/v) 2-mercaptoethanol to stabilize the enzyme. Column 60 X 4 cm. Flow rate 20 ml/hr, 4 ml fractions. (A)OD at 280 nm, ( 0 )OD at 540 nm, ( B ) LDH; assay with p-nitrophenyl phosphate for AcP. From Hopkinson and Harris (86).

are to be translated into sequence differences, methods capable of largescale preparation must be developed. Fisher and Harris (86) have effected a 1300-fold purification of B acid phosphatase from human erythrocytes with a yield of 27%. The method involves adsorption on calcium phosphate gel, ammonium sulfate precipitation, and gel filtration. This type of procedure should allow the preparation of enzymes in sufficient quantity to do the type of studies that are now needed. In addition to differences in electrophoretic mobility between the various heritable types of red cell acid phosphatase, there are striking quantitative differences. Figure 13 (85) demonstrates that the mean level of enzymic activity of type B acid phosphatase is considerably greater than the mean level of type A. Type BA lies almost exactly between the two curves. The genetic relationships of the human isoenzymes have been reviewed by Giblett ( 7 9 ) . 86. R. A. Fisher and H. Harris, Biotechnol. Bioeng. 10, 829 (1968).

484


Red cell acid phosphatase activity

FIG.13. The distribution of red cell acid phosphatase activities in a randomly selected English population (dotted line) and in the separate phenotypes. From Hopkinson and Harris (86).

IV. liver Acid Phosphatase

-4.RATLIVER 1. Cellular Location

The lysosomal nature of liver acid phosphatase has been demonstrated by morphological and centrifugal techniques (87,88). Considerable interest has been shown in the purification of this enzyme because of its possible role in physiological and pathological catabolic processes (4,89, 90). The pericanalicular bodies of rat liver characteristically show activity for acid phosphatase when either glycerophosphate or naphthyl phosphate is used as substrate a t pH 5.0-5.5. Adult rabbit liver does not generally react with glycerophosphate as substrate, though acid phosphatase can be visualized with naphthyl phosphate. Livers of newborn and young rabbits show activity with both substrates. Since enzymic activity can be localized a t different sites in the same section by employing one and then the other substrate, it seems clear that the same enzyme is not involved (91). I n formalin-fixed rat liver, acid phosphatase activity is localized in peribiliary granules in the hepatic cells and in the Kupffer cells. The 87. J. Berthet and C. De Duve, BJ 50, 174 (1951). 88. J. Berthet, L. Berthet, F. Appelmans, and C. De Duve, BJ 50, 182 (1951). 89. B. W. Moore and P. U. Angeletti, Ann. N . Y . Acad. Sci. 94, 659 (1961). 90. R. Brightwell and A. L. Tappel, A B B 124, 333 (1968). 91. R. M.Rosenbaum and C. I. Rolon, Histochemie 3, 1 (1962).

19. ACID

PHOSPHATASEB

485

enzyme-containing granules correspond to the lysosomes and phagosomes as described by De Duve and co-workers (92,93). 2. Isolation and Purification

Barka (94) studied the acrylamide gel electrophoretic behavior of distilled water homogenates of rat liver prepared by high-speed centrifugation after a number of alternate freeze-thaw procedures. The soluble enzyme represented 70% of the total enzymic activity in crude homogenates of rat liver. In order that satisfactory electrophoretic separations be obtained, it was necessary that the gel contain a minimal amount of persulfate and be subjected to electrophoresis for some time prior to addition of the sample to the gel. Under these conditions, three separate bands of enzymic activity were obtained a t pH 8.7,while a minimum of 20 protein bands were separated. From these studies, it would appear that hepatic acid phosphatase occurs in rat liver in at least four forms; the last being the particle-bound activity not easily extracted by water homogenization. About 60% of the total acid phosphatase can be released from the mitochondrial-lysosomal fraction by freezethaw, sonication, or blender treatment. This material contains a single electrophoretic band of enzymic activity, while Triton X-100released almost all of the activity from the particles but gave two separate bands. These two fractions may represent material bound in different ways to lysosomes (96). The relationship of these two fractions and the soluble enzymes to the two fractions obtained by more gentle extraction in the procedure of Igarashi and Hollander ( 4 ) is not yet known. Two highly purified enzymes, one crystalline, have been obtained from rat liver. Table XVI ( 4 ) illustrates the general method of purification. Rat livers were homogenized in 50% glycerol and a pH 5 supernatant from this extract subjected to ammonium sulfate fractionation. Chromatography on Sephadex G-75gave a single peak of activity which was then fractionated on DEAE-cellulose which gave two peaks. One was not absorbed and the other was eluted at 0.095 M sodium chloride. Peak I material which failed to absorb t o DEAE-cellulose was chromatographed on hydroxylapatite. The eluate from this column after concentration and solution in 5 mM imidazole-glycine buffer, pH 7.1,gave a crystalline enzyme when this solution was brought slowly to 0.55 saturation with ammonium sulfate. Peak I1 enzyme eluted from the 92. C. De Duve, A m . Physiol. Soc. (Washington) p. 128 (1959). 93. A. B. Novikoff and E. Essner, A m . J . M e d . 29, 102 (1960). 94. T.Barka, J . Histochem. Cytochem. 9, 542 (1961). 95. J. M . Allen and J. Gockerman, Ann. N . Y . Acad. Sci. 121, 616 (1964).

486


TABLE XVI OF PURIFICATION PROCEDURE" SUMMARY

Fraction ~~

~~

Total activity (unit)

Specific activity (unit/ mg)

Purification (-fold)

Recovery

4320 2420 2296 500 404 398 130 91.2 92.4 78.4

0.020 0.112 0.124 0.268 1.12 10.6 18 18 0.64 2.68

1.0 5.6 6.8 13.4 56 532 885 890 32 134

100 56 53 12 9.4 9.2 3.0 2.1 2.1 1.8

(%I ~

~

Homogenate pH 5 Supernatant Ammonium sulfate Sephadex G-75 DEAE-cellulose peak I Hydroxylapatite Crystallization I Crystallization I1 DEAE-cellulose peak I1 Sephadex G-200

1612 211,000 2881 21,700 278 18,500 1,860 98.6 360 38.4 37 388 7.3 0.8 5.1 0.5 140 242 29 266

____

a

Total protein (mg)

Total volume (ml)

~

From Igarashi and Hollander (4).

DEAE-cellulose column was further purified on Sephadex G-200 but resisted all efforts a t crystallization. Figure 14 ( 4 ) shows the polyacrylamide gel electrophoretic pattern of crude rat liver extract with two bands, the resolution of these bands by DEAE-cellulose, and the crystalline peak I preparation. The crystalline enzyme had an approximate molecular weight of 100,OOO estimated by sucrose density gradient

1

1

2

3

4

5

FIG.14. Polyacrylamide gel electrophoresis at various purification steps. Gels 1 to 4 were stained for acid phosphatase activity; gel 5 was stained for protein. A current of 4 mA/gel was applied to gels 1 3 for 2 hr and to gels 4 and 5 for 6 hr. Gel 1, homogenate; 2, DEAE-cellulose peak 11; 3, DEAE-cellulose peak I ; 4 and 5, crystalline enzyme. From Igarashi and Hollander ( 4 ) .

19.

487

ACID PHOSPHATASES

centrifugation and Sephadex G-200 chromatography. Peak I1 enzyme had the same molecular weight. Mixtures of crystalline and peak I1 enzyme could not be separated by either G-200or sucrose density gradient centrifugation. The isoelectric points of crystalline and peak I1 enzymes were pH 7.7 and 4.5, respectively, when measured by the method of isoelectric focusing. Table XVII ( 4 ) shows that p-nitrophenyl phosphate (NP), P-glycerol phosphate, and fructose 1,6-diphosphate were hydrolyzed rapidly and in that order. Both the crystalline enzyme and peak I1 were free from measurable diphosphatase, pyrophosphatase, or alkaline phosphatase activity. D- ( f ) -Tartrate and fluoride were powerful inhibitors of the hepatic enzyme. Adenine nucleotides and oxalate were also effective inhibitors. No metal requirement or effect of EDTA on enzymic activity was found. Table XVIII ( 4 ) shows the effect of dioxane and oxalate on K , and TABLE XVII SUBSTRATE SPECIFICITY AND INHIBITORS" Inhibition with N P as substrate

Relative rate of hydrolysis* Compound

Crystalline

NP ATP

ADP AMP p-Gly cerol-P CX-D-G~UCOS~-~-P D-G~UCOS~~-P Fructose-1,6-di-P PP, Cyclic 3',5'-AMP Cyclic 2',3'-AMP Cyclic 2',3'-UMP Cyclic 2',3'-GMP Oxalate Pi L(+)-Tartrate Fluoride a

P I1

1.00

1 .00

0 0

0 0

0.20 0.33 0.07 0.05 0.15 0 0

0.27 0.35 0.05 0.02 0.14 0

0 0 0

0 0 0 0

Crystalline

P I1

0.68 0.67 0.58 0.08 0.21 0.05 0.24 0.22

0.66 0.61 0.52 0.07 0.17 0.03 0.21 0.22

0 0 0 0 0.37 0.16 0.97 0.94

0 0 0 0 0.37 0.17 0.97 0.96

From Igarashi and Hollander ( 4 ) .

* Relative rates of hydrolysis were determined with 0.5 ml reaction mixtures in 0.1 M sodium acetate buffer, pH 5.0, a t 37". Liberated phosphate was measured by the method of C. H. Fiske and Y. SubbaRow [JBC 66, 375 (1925)]. The amounts of enzyme used were 0.22 unit of crystalline enzyme and 0.24 unit of peak I1 enzyme. The concentration of substrate and inhibitor was 1.0 mM. For inhibitor study, 1.0 mM p-nitrophenyl phosphate was used as substrate. Inhibition was calculated from the amount of p-nitrophenol released and expressed as fractional inhibition.

488


TABLE XVIII EFFECTOF DIOXANE AND OXALATE ON KINETIC CONSTANTS"-*

K , f S.E. Addition to reaction mixture None Dioxane (8%) Dioxane (8%) and oxalate

Crystalline (mM)

Vmaxf S.E. P I1 (mM)

0.091 f 0.007 0.047 f 0.004 0.099 f 0.008 0.042 f 0,004 0.20 5 0.02 0.25 f 0.04

Crystalline

P I1

(1@ X AA/min)

(102 X AA/min)

1.88 f 0 . 1 2 2.01 k 0.12 1.28 f 0 . 1 3

2 . 3 1 f 0.08 2 . 6 4 f 0.13 1.76 f 0 . 1 3

1.29f0.15

1.77f0.14

(1.5 d )

Oxalate (0.6 mM)

0.24f0.05

0.19fO.04

From Igarashi and Hollander ( 4 ) . The reaction mixtures consisted of varying concentration of p-nitrophenyl phosphate in the range of 0.0075 to 0.19 mM and the indicated amounts of dioxane and oxalate, in a total volume of 1.0 ml, in 0.1 M sodium acetate buffer, pH 5.0. Triplicate measurements were done at each substrate concentration. Eight determinations were made under each experimental condition. a

b

V,,,,, of the two enzymes. Oxalate inhibition is of the mixed type; the inhibitor affects both K,,, and VlllilX.Neither variable is affected by dioxane. Figure 15a ( 4 ) shows that in the absence of dioxane, oxalate inhibition gives the expected hyperbolic curve. However, addition of dioxane to a final concentration of 8% converted the hyperbolic curve to the sigmoidal type of inhibition. This unexpected relationship is brought out in Fig. 15b ( 4 ) which contrasts plots of the reciprocal fractional inhibition against reciprocal oxalate concentration. The departure from linearity produced by the addition of dioxane to the reaction mixture is striking. The value of n in the figure, the slope of the Hill equation (96),is one in the absence of dioxane and greater than one in the presence of 8% organic solvent. The figure also shows that addition of dioxane to reaction mixtures inhibited by trypan blue gave very similar curves. Both the crystalline and peak I1 enzyme showed this behavior. Dioxane did not act by changing the enzyme in any permanent way, since if the organic solvent were removed by Sephadex chromatography, the original enzymic behavior was restored. Neither oxalate nor dioxane changed the apparent molecular weight of the enzyme. Table XIX ( 4 ) shows that treatment of the enzyme with fluorodinitrobenzene desensitized the crystalline enzyme to oxalate inhibition. Such dinitrophenylation did not appreciably alter the pH-activity curves of the 96. K. Taketa and B. M. Pogell, JBC 240, 651 (1965).

19.

489

ACID PHOSPHATASES

c

0.8

.s $ -

loo

.02 Trypan blue(pg/rnl) (0)

06

04

08

I

I /Trypan blue

(b)

FIG.15. ( a and b) Inhibition of crystalline acid phosphatase by varying concentrations of oxalate or trypan blue in the presence and in the absence of dioxane. Initial velocity was determined as in Table XVIII with 0.185 mM p-nitrophenyl phosphate, varying concentrations of oxalate or trypan blue, and 1.20 absorbance units of crystalline enzyme in the presence or absence of 8% dioxane. Peak I1 enzyme gave similar results. From Igarashi and Hollander (4).

enzyme. Desensitization was best achieved in the presence of phosphate, presumably because this anion protects the active site. Brightwell and Tappel (90)purified rat liver acid phosphatase from a lysosomal fraction by DEAE and CM-cellulose chromatography. Table XX (90)shows the specificity of the lysosomal enzyme. B. MOUSELIVER Mouse liver acid phosphatase is localized in the Kupffer cells in contrast to the alkaline phosphatase activity which is largely confined to the endothelial linings of the sinusoids. Under the conditions in which the activity of the reticuloendothelial system is enhanced, both enzymic activities are increased (97, 98). 97. G. J. Thorbecke, L. J. Old, B. Benacerraf, and D. A. Clarke, J. Hidochem. Cytochem. 9, 392 (1961). 98. P. Van Duijn, R. G. J. Willighagen, and A. E. F. H. Meijer, Biochem. Pharmacol. 2, 177 (1959).

490


TABLE XIX OF CRYSTALLINE ACID PHOSPHATASE TO OXALATE DESENSITIZATION

INHIBITION WITH FLUORODINITROBENZENE~-~ Inhibition with oxalate Incubation period (hr)

Buffer Phosphate (pH 8.1, r/2 = 0.1)

2 4

Tris-acetate (pH 8.1, r/2 = 0.1)

f 3

Tris-acetate, with 30 mM oxalate

t

4 Tris-acetate, with 20 mM N P

4

Control 0.50 0.50 0.50 0.50 0.50 0.50 0.50

Activity" Fluorodirecovered nitroafter dinitrobenzene phenylation 0.30 0.09 0.49

0.57 0.35 0.35 0 0.48 0.09 0.37

0.46 0.23

From Igarashi and Hollander (4). For each experiment, 0.5 ml of crystalline enzyme (0.01 mg) in the indicated buffer was stirred with 40 pmoles of fluorodinitrobenzene in the dark at room temperature. After the indicated period, the mixture was passed through a column, 1 x 15 cm, of Sephadex G-25. The control was prepared by an identical procedure but without fluorodinitrobenzene. c The activity of native enzyme is 1.00. 0

b

TABLE XX OF ACID PHOSPHATASE' SUBSTRATE SPECIFICITIES

Substrate

Conc. (mM)

Activity

Enzymeb Acid phosphatase

P-Gly cerophosphate p-Nitrophenyl phosphate Phosphoenol pyruvate ADP 5'-AMPd 3'-AMP 2'-AMP G-1-P G-6-P

25 8 1.0 1.5 1.5 1.5 1.5 1 .o 1.0

100 130 59 9 63 74 37 13 11

(%I

From Brightwell and Tappel (90). Enzymes were measured in fractions from the CM-cellulose eluate. c Inhibition by 1.4 m M D-(+)-tartrate was for 20 min of incubation. d No MnClz added. 4

b

Inhibition by tartrate' ( %)

97 80 87 92 88 98 98 81 79

19.

ACID PHOSPHATASES

491

MacDonald (99) showed that mouse liver acid phosphatase required active sulfhydryl groups for activity and that malonate buffer, pH 5.9, was useful for the assay of this enzyme because it stabilized the enzyme during the period of the assay. Verity and Reith (100) studied the nature of activation of lysosomal acid phosphatase in a large granule fraction from adult mouse liver. Such preparations have less than 10% of the acid phosphatase activity which they can exhibit following solubilization with Triton X-100. It should not be assumed that the additional activation resulted simply from particle disruption. Exposure of such lysosomal preparations to Hg" or organic mercurials produced maximal activation and an irreversible loss of structure-linked latency. Activation of three lysosomal enzymes-acid phosphatase, p-glucuronidase, and N-acetyl-p-o-glucosaminidase-were not comparable under these conditions. These findings suggest that acid phosphatase activation by destruction of structurelinked latency involves more specific reactions than mere membrane rupture.

C. BOVINELIVER Heinrikson (101) has purified a low molecular weight acid phosphatase from bovine liver (acid phosphatase 111) to apparent homogeneity. Bovine liver was extracted with 0.3 M sodium acetate, pH 5.0 containing 1 mM EDTA. Ammonium sulfate fractionation and acid precipitation of inactive material gave material which could be fractionated on Sephadex G-75. This step gave three peaks of activity. The third peak (fraction 111) had a low molecular weight and was extensively purified by chromatography on sulfoethyl Sephadex phosphate. An overall 5000-fold purification could be achieved by this procedure. The final product was homogeneous by electrophoresis on polyacrylamide gel and showed constant specific activity during rechromatography on sulfoethyl Sephadex. End group analysis using dimethylaminonaphthalene-5-sulfonylchloride gave a single product, the arginine derivative. No other amino-terminal residue could be detected. The molecular weight estimated from a carefully calibrated Sephadex G-75 column was 14,000. Sedimentation equilibrium gave a value of 16,590. Table XXI (101) shows the amino acid composition of this purified bovine liver acid phosphatase. A Lineweaver-Burk plot of the hydrolysis of p-nitrophenyl phosphate 99. K. MacDonald, BJ 80, 154 (1961). 100. M. A. Verity and A. Reith, BJ 105, 685 (1967) 101. R . L. Heinrikson, JBC 244, 299 (1969).

492


TABLE XXI AMINOACIDCOMPOSITION OF ACIDPHOSPHATASE 111.

Amino acid Lysine Histidine Arginine Aspartic acid asparagine Threonine Serine Glutamic acid glutamine Proline Glycine Alanine Half-cystine cysteine Valine Methionineb Isoleucinec Leucine Tyrosine Phenylalanine Tryptophand

+

+

+

4

No. of residues per molecule (MW 16,296) 13

2

7 15

7 6 16 9 10 10 2 13 4 8 11 4 6 2

From Heinrikson (101).

* Methionine plus methionine sulfoxide. c

Isoleucine plus alloisoleucine. Determined spectrophotometrically.

gave an excellent straight line with a K , of 7.5 x lo4 M . Optimal hydrolysis was achieved a t pH 5.5. The presence of Mg2+or mercaptoethanol in incubation mixtures led to rapid inactivation. Ethylenediaminetetraacetate exhibits a stabilizing effect. Table XXII (101) summarizes the activity of the purified acid phosphatase I11 toward a number of substrates. The most active hydrolysis is directed toward p-nitrophenyl phosphate, but a number of other physioIogically important phosphates are hydrolyzed a t significant rates. Very slight hydrolysis occurs with a-glycerophosphate and none whatever with the /3 form. Bovine liver acid phosphatase 111 should prove extremely useful for further studies on the physiological role of this enzyme. It can be obtained in good yield: 500 g of bovine liver yielded approximately 2 mg of apparently homogeneous enzyme. Yields of this order from starting material available in bulk will allow considerable further study of this purified enzyme. The amino acid composition of bovine acid phosphatase

19.

493

ACID PHOSPHATASES

TABLE XXII SUBSTRATE SPECIFICITY OF ACID PHOSPHATASE IIIo~* ~~

Activity relative to pnitrophenyl phosphate Substrate

(%I

p-Nitrophenyl phosphate Flavin mononucleotide Galactose &phosphate Glucose 1-phosphate 5'-UMP Pyridoxal 5-phosphate Fructose 6-phosphate Glucose 6-phosphate a-Gly cerophosphate Pyridoxamine 5-phosphate Mannose 6-phosphate D-Ribose 5-phosphate AMP,CMP, G M P B-Gly cerophosphate Pyrophosphate Bis-pnitrophenyl phosphate Nucleoside diphosphates

100

68 39 14 10 5 4 4

3 3 2 2 0 0 0 0 0 ~~

From Heinrikson (101). * Assays were carried out at 37" and pH 5.5. Each mixture contained 500 pmoles of sodium acetate, 5 pmoles of EDTA, 1.7 pg of step VII phosphatase, and 20 pmolea of substrate in a volume of 5.0 ml. Controls lacking enzyme were run together with each of the substrates tested. At 1, 2, 5, 10, and 20 min, 500 pl portions were removed and added to 4.5 ml of 0.6 N H4S04. Inorganic phosphatase was determined colorimetrically, and the initial rates were compared to that obtained with p-nitrophenyl phosphate. 4

111 is quite similar to a low molecular weight acid phosphatase purified by von Hofsten and Porath (109).

V. Spleen Acid Phosphatase

Chersi et al. (103)have carried out extensive purification of spleen acid phosphatase. Spleen was fractionated to yield crude spleen nuclease I1 (104). This preparation was found to contain large quantities of non102. B. von Hofsten and J. Porath, BBA 84, 1 (1962). 103. A. Chersi, A. Bernardi, and G. Bernardi, BBA 129, 12 (1966). 104. G. Bernardi, A. Bernardi, and A. Chersi, BBA 129, 1 (1966).

494


TABLE XXIII RELATIVE RATE OF HYDROLYSIS OF SEVERAL SUBSTRATES BY SPLEEN ACID PHOSPHOMONOESTERASE~~~ Relative rate of hydrolysis

(%I

Substrates p-Nitrophenyl phosphate 2',3'-AMP 5'-AMP &Glycerophosphate Glucose 1-phosphate Riboflavin phosphate Thiamin phosphate Phosphothreonine Phosphoserine ATP bis(p-Nitrophenyl) phosphate

100 96 63 60 9 29 21 6 4 0 0

From Chersi et al. (103). The substrate concentration was 1.5 mM. The liberation of inorganic phosphate at 37" was determined a t different incubation times. The initial rates were used. 0

b

TABLE XXIV EFFECTSOF SOME ACTIVATORSAND INHIBITORS ON SPLEEN ACIDPHOSPHOMONOESTERASE~.~ Activators or inhibitors

2',3'-AMP

p N i trophenyl phosphate

(%I

(%I

0.01 M cysteine 0.001 M cysteine 0.01 M EDTA 0.001 M M@+ L(-)-Tartaric acid D-(+)-Tartaric acid 0.01 M cu*+ 0.001 M Cup+ 0.01 M F0.001 M F0.001 M Mo'+

100 104 94

100 100 102 89 90 8.3 -

8.6 38 2 9

7.8 0.5

From Chersi et al. (103). Assays were performed uskg the following incubation mixture: 2 pmoles of substrate in 2 ml of 0.05 M acetate (pH 5.0); activator or inhibitor in 0.2 ml; enzyme, diluted with bovine serum albumin, in 0.2 ml. The concentration of activator or inhibitors refer to the solutions before adding the enzyme. 0

b

19.

495

ACID PHOSPHATASES

specific acid phosphatase activity. Trimmed bovine spleen was extracted with 0.1 M HCl, brought to pH 2.5, and the supernatant fractionated with ammonium sulfate. Appropriate fractions were then subjected to DEAE, hydroxylapatite, Sephadex G-100, and CM-Sephadex chromatography. The substrate specificity of the purified enzyme is shown in Table XXIII (103).The sedimentation constant was 5.6 S by a sucrose density gradient centrifugation. The final acid phosphatase portion was free of any acid deoxyribonuclease, acid ribonuclease, exonuclease, and phosphodiesterase activity. It can be seen that p-nitrophenyl phosphate is an excellent substrate. Table XXIV (103)shows the effect of a variety of inhibitors on the purified enzyme. pH-Activity curves show a broad maximum between pH 3 and 4.8. The Michaelis constant a t 37" and pH 5.0, p = 0.05, is 7.25 x M . Phosphate ion is a competitive inhibitor of the enzyme.

VI. Acid Phosphatase in Serum

Zucker and Borrelli (31,105) showed that platelet-rich and plateletpoor plasma were completely different when assayed against P-glycerophosphate for acid phosphatase activity. No activity was found in serum prepared from platelet-poor plasma, whereas values up to 0.49 Bodansky unit were found in serum from platelet-rich plasma. This suggested that the platelets were the major source of the normal serum acid glyceropliosphatase activity. When p-nitrophenyl phosphate was used, serum from native platelet-poor plasma had some activity, indicating that the platelets were not the only source of serum activity with the substrate. hctivity was always greater in serum from platelet-rich plasma. Forty to eighty percent of the activity could be attributed to the platelets. Platelet acid nitrophenyl phosphatase is inhibited to a considerable degree by formaldehyde but very little by D-(+)-tartrate or Mg2+. I n these respects, it behaves like the enzyme in normal serum. I n contrast, prostatic, liver, and spleen acid phosphatases are strongly inhibited by D- (+)-tartrate. Formaldehyde partially inhibits the enzyme in serum and many tissues but completely inhibits the acid phosphatase activity from the erythrocyte. The enzymic activity can be found in suspension of intact, washed platelets and greater activity is obtained by freezing and thawing the platelet suspensions. No role for platelet acid phosphatase in blood clotting or any other function has been elucidated. The clinical interpretation of elevations of serum acid phosphatase is compli105. M. B. Zucker and J. Borrelli, Ann.

N. Y . Acad. Sci. 75, 203 (1958).

496


cated because of the presence of prostatic, platelet, and erythrocyte phosphatases in a complex mixture (106, 107). The methodology, accuracy, and precision of manual and automated methods for serum acid phosphatase estimation for clinical purposes have been reviewed by Bodansky and Schwartz (108).

VII. Miscellaneous Sources

A. GAUCHER ACIDPHOSPHATASE Serum of individuals with Gaucher’s disease have electrophoretic peaks of enzymic activity distinct from that of the prostatic enzyme (109), and acid phosphatase activity is very often elevated (110, 111). Unlike the acid phosphatase of prostatic and erythrocyte origin, this serum phosphatase is not significantly inhibited by D-(+)-tartrate or Cu2+.It is moderately inhibited by formaldehyde. The source of the acid phosphatase is apparently the Gaucher cell, a large macrophage containing nonmetabolized glucocerebroside. This cell contains abundant acid phosphatase by histochemical examination. It splits phenyl phosphate much more rapidly than glycerophosphate, is an excellent nucleosidetriphosphatase, and has maximal activity a t pH 4.0-4.2. It does not seem to be identical with spleen acid phosphatase (112).The more general interest in Gaucher acid phosphatase lies in the concept that many variants of acid phosphatase activity may exist in mammalian organisms but are not detected because the particular cell population producing the activity is very small. I n some pathological situations where the cell population increases enormously, new variants may appear.

B. BONEACIDPHOSPHATASE Bone contains two distinct enzymes, acid and alkaline phosphatases, which are associated with osteoclasts and osteoblasts, respectively (113). Further study of acid phosphatase and bone should be rewarding. Many 106. H. Q . Woodard, A m . J . Med. 27, 902 (1959). 107. R. Bases, New Engl. J . M e d . 266, 538 (1962). 108. 0. Bodansky and M. K. Schwartz, Methods Med. Res. 9, 79 (1961). 109. A. F. Goldberg, K. Takakura, and R. L. Rosenthal, Nature 211, 41 (1966). 110. L. R. Tuchman, G. Goldstein, and M. Clyman, A m . J . Med. 27, 959 (1959). 111. B. Estborn and P.-0. Hillborg, Scand. J . Clin. & Lab. Invest. 12, 504 (1960). 112. A . C. Crocker and B. H. Landing, Metab., Clin. Ezptl. 9, 341 (1960). 113. M. S. Burstone, J . HCtochem. Cytochem. 7, 39 (1959).

19. ACID

PHOSPHATASES

497

physiological studies show that bone resorption is accompanied by increased levels of enzymic activity, but no systematic studies have been carried out on the chemical characterization of the activity.

C. PLANTACIDPHOSPHATASE Seedlings are a rich source for nonspecific acid phosphatase. Newmark and Wenger (114) have reported on a 1000-fold purification from lupine seedlings. The purified enzyme hydrolyzes phosphate monoesters and pyrophosphate with p-nitrophenyl phosphate as substrate. The optimal activity was a t pH 5.2-5.5, and K,,, was 3 X lO-*M. Fluoride inhibition was noncompetitive. Shaw (115) reported a 300-fold purification of enzyme from tobacco leaves. Activity of the enzyme was optimal a t pH 5.5-5.7, and divalent cations were not required for activity. The enzyme possessed high activity toward ribonucleoside 2'- and 5'-monophosphates and glucose l-phosphate. There was no activity toward RNA or phosphodiesters. Fluoride acts as a noncompetitive inhibitor for this enzyme. This behavior of fluoride is in contrast to the behavior with prostatic acid phosphatase where the inhibition is strictly competitive.

D. Neurospora crassa An acid phosphatase from the mycelium of the fungus Neurospora crassa has been purified 1400-fold with a 40% recovery. The pH maximum is 5.6 with P-glycerophosphate as substrate. Fluoride and D-(+) tartrate are competitive inhibitors so that this enzyme fits into a rather general pattern for others of its type which have been described (116).

-

E. Saccharomyces PHOSPHATASE A repressible acid phosphatase of Saccharomyces mellis develops when the organism is grown in a medium free of phosphate. Only traces of enzymic activity are found when media containing inorganic phosphate are used. The enzyme is inhibited by phosphate, arsenate, molybdate, and borate (117). 114. M. 8.Newmark and B. S. Wenger, ABB 89, 110 (1960). 115. J. G.Shaw, ABB 117, 1 (1966). 116. M.-H. Kuo and H. J. Blumenthal, BBA 52, 13 (1961). 117. R. Weimberg and W. L. Orton, J . Bacterial. 86, 805 (1963).

498


F. STAPHYLOCOCCAL ACIDPHOSPHATASE Acid phosphatase of S. aureus PS 55 is eluted from the cell surface by 1.0 M KC1 a t pH 8.5. Gel filtration of this material gave a 44-fold purification. The protein seems homogeneous by gel filtration, starch block electrophoresis, and analytical ultracentrifugation with the weight of approximately 58,000 ( l g a ).

G. AMEBICPHOSPHATASE The acid phosphatase activity of the ameba, Chaos chaos, is largely confined to particulate bound enzyme which exhibits latency. A noncompetitive heat-stable inhibitor is present in the particulate fraction. The role of this inhibitor in the mechanism of lysosomal activation is not clear (I%, I&). H. E . coli ACIDPHOSPHATASE At least four acid phosphatase fractions have been obtained by ammonium sulfate, DEAE hydroxylapatite, and electrophoretic separation. One type of activity, nucleoside 2’- or 3‘-phosphatase was purified 1500fold. Hexosephosphatase activity was also obtained in three separate fractions. All three fractions were different with respect to rate of splitting of different substrates and pH optimum (118).

I. Melanogaster MacIntyre and Dean (119) report that acid phosphatase from D. melanogaster has “slow and fast” electrophoretic variants specified by co-dominant alleles. Thus, acid phosphatases AA, BB, and AB were studied. Types AA and BB could be inactivated by exposure to acid. Reactivation of enzymic activity could be accomplished by dialysis against buffers a t pH 6.5. Mixtures of AA and BB produced some AB reconstituted enzyme. From this evidence it seems very probable that acid phosphatase, a t least in this species, consists of a t least two polypeptide chains.

118. D. Rogers and F.J. Reithel, ABB 89, 97 (1960). 119. R.J. MacIntyre and M.R.Dean, Nature 214, 274 (1967)

Inorganic Pyrophosphatase of Escherichia coli JOHN JOSSE

SIMON C. K . WONG

I . Introduction . . . . . . . . . . . . . I1. Molecular Properties . . . . . . . . . . . A . Purification . . . . . . . . . . . . B . Homogeneity . . . . . . . . . . . C . Size . . . . . . . . . . . . . . D. Physical Properties . . . . . . . . . . E . Electron Microscopy . . . . . . . . . F. Subunits in 5 M Guanidine Hydrochloride . . . . G . Reconstitution of Native Enzyme Particles from Subunit Polypeptide Chains . . . . . . . . . . H . Chemical Composition . . . . . . . . . I . Chemical Modification of Residues Essential for Enzymic Activity . . . . . . . . . . I11. Catalytic Properties . . . . . . . . . . . A . p H Effects . . . . . . . . . . . . B. Effects of Ions and Inhibitors . . . . . . . C . Reversal of Reaction . . . . . . . . . D. Substrate Specificity and Stoichiometry . . . . . E . Nature and Binding of Active Substrate and the Role of Magnesium . . . . . . . . . . . . F. Interactions with Inhibitors . . . . . . . . 1V . Conclusions . . . . . . . . . . . . . .

499 501 501 602 504 504 506 508

510 612

514 518 518 518 519 520 522 525 626

.

1 Introduction

Enzymes which catalyze hydrolysis of inorganic pyrophosphate ( PPI) are ubiquitous throughout nature. Insofar as the authors are aware. there 499

500

J. JOSSE AND S. C. K. WONG

is no instance in which this activity has been sought and not found in any organism-animal, vegetable, or microbial. Furthermore, when assay conditions have been optimal, a marked abundance of the activity has invariably been demonstrated, far more than would appear to be necessary for binding and catalytic hydrolysis of the estimated amounts of PP, produced in cell metabolism. For example, in Escherichiu coli a simple calculation, employing the specific activities of pure enzyme and of crude cell extracts, leads to the estimate that inorganic pyrophosphatase constitutes 0.2% of the total soluble protein of this bacterium. This corresponds to approximately 1000 molecules of enzyme (molecular weight: 120,000) per bacterial cell, an amount that will catalyze hydrolysis of about 1.5 X log molecules of PP, per minute a t 37” (enzyme turnover number: 1.5 x 108). It is unlikely that E . coli metabolism produces even 1/1000 that much PP,. We have deliberately tried to isolate mutant E . coli strains in which this enzyme might be missing or altered. To date and after processing over 5000 colonies of cells treated with mutagenic agents, we have found only three strains with diminished activity, the lowest level of which was 2% of normal. This mutant behaved quite normally, even after adaptation to growth on PP, as the sole source of phosphorus ( I ) , and was indistinguishable from wild type cells in growth kinetics in various media and under different conditions of temperature and atmosphere. When either mutant or wild type cells were grown on 3zP, (0.1 mCi/pmole) as the source of phosphorus, 32PPiwas not found in cell extracts (isolated in the presence of excess unlabeled carrier PPi). Even 2% of the usual amount of pyrophosphatase activity is apparently adequate to dispose of all of the PP, formed during the course of E . coli metabolism. I n those instances where PP, has been demonstrated in vivo in other organisms (2-4), its mode of formation and means of escape from intracellular pyrophosphatase action have not been clarified. Whatever the role of PP, in cell metabolism, nature seems to have assured that little of it has a chance to survive for long. Is this a “vital enzyme,” one without which cells cannot grow and survive? Again in E . coli, we have attempted to answer this question by isolation of temperature-sensitive mutants that will grow a t a low (15”1. Escherichia coli can be adapted to grow on PPI aa the sole murce of phosphorus; in this situation the cells are dependent upon intracellular inorganic pymphosphatase activity to make PI available. To avoid possible confusion from the inducible alkaline phosphatase of E . coli, which also has pyrophosphatase activity (see Chapter 17, by Reid and Wilson, this volume), all mutant isolation studies began with an E coli strain unable to synthesize this inducible protein. 2. T. Mann, BJ 38, 345 (1944). 3. P. E. Lindahl and K. Kiessling, Arkiv Kemi 3, 97 (1951). 4. H. Fleisch and S. Bisaz, Am. J . Physiol. 203, 671 (1962).

20.

INORGANIC PYROPHOSPHATASEI OF

E . coli

601

25") but not a t a higher (35"-40") temperature (6). Of all of the mutants thus far isolated (>250) , none was found to have abnormal levels of inorganic pyrophosphatase when cell extracts were assayed a t either high or low temperatures. Use of penicillin selection techniques (6) and media containing PPI as the sole source of phosphorus ( I ) gave the same result, which may be related to the marked temperature insensitivity of E . coli as well as most other inorganic pyrophosphatases (e.g., see Fig. 7 below) ( 7 ) . Therefore a clear-cut answer to this question cannot be given at the present time. A metabolic role for intracellular PPI hydrolysis has been suggested by Stetten (8) and by Kornberg (9). The argument is as follows: PPI is a by-product of numerous important enzymic syntheses, including the reactions of deoxyribonucleic acid (DNA) and ribonucleic acid (RNA) polymerization, coenzyme synthesis, and amino acid and fatty acid activations. Although some of these reactions are themselves exergonic, the free energy change often is not great, as, for example, in amino acid activation (10). However, by coupling such reactions with catalytic hydrolysis of concurrently produced PP, [AF' (standard free energy change a t neutrality) = -5 kcal], the overall equilibrium can be markedly shifted in favor of synthesis. In this way inorganic pyrophosphatases may be vital to assure ongoing progress of the anabolic phases of cell physiology. Because so much of modern biochemistry has utilized E.coli as a model cell for investigative purposes, it was of interest to study the inorganic pyrophosphatase of this organism in greater detail. The enzymes from yeast and other sources are described in Chapter 21 by Butler, this volume.


A. PURIFICATION

Escherichia coli pyrophosphatase was first isolated by Schito and Pesce (11) and later by Josse ( 1 2 ) . A large-scale purification method 5. R. H. Epstein, A. BollB, C. M. Steinberg, E. Kellenberger, E. Boy de la Tour, R. Chevalley, R. S. Edgar, M. Susman, G. H. Denhardt, and A. Lielausis, Cold Spring Harbor Symp. Quant. Bwl. 28, 375 (1963). 6. J. Lederberg and N. D. Zinder, JACS 70, 4287 (1948). 7. A. Schaffner and F. Krumey, 2.Physbl. Chenz. 255, 145 (1938). 8. D. Stetten, Jr., Am. J . M e d . 28, 867 (1900). 9. A. Kornberg, in "Horizons in Biochemistry" (M. Kasha and B. Pullman, eds.), p. 251. Academic Press, New York, 1982.

502


TABLE I PURIFICATION OF E . coli PYROPHOSPHATASE~ ~

Fraction I. 11. 111. IV. V. VI. VII.

Extract Streptomycin Heat (SO", 10 min) Ammonium sulfate Sephadex G-150 DEAESephadex Crystals

Total protein (mg) 225,000 126,000 12,300 8,000 1,000 390 320

Specific activity (units/mg protein) 4

7 70 100 750 1,800 2,000

Yield of activity

(%) 100 98 95 89 83 78 71

From Wong et al. (IS). A unit of enzymic activity is that amount which will catalyze hydrolysis of 10 pmoles of PPi in 15 min at 37" in the standard assay procedure (1.9).

with improved yields has recently been described (13). This procedure, in which 300-350 mg of crystalline, homogeneous protein are obtained from 2 kg of packed E . coli cells (wet weight) in about 3 weeks, is summarized in Table I. The enzyme crystals are cubic in shape (Fig. l a ) and can be stored in suspension a t 2" indefinitely ( > 2 years) without loss of activity. Very large crystals can be formed by recrystallization from 0.15 M potassium phosphate buffer, p H 7.5, 1.26M ammonium sulfate a t 38" ; after 5-15 days crystals with the habit and dimensions shown in Fig. l b are obtained. However, these proved exceedingly difficult to handle and shattered readily into multiple small cubes upon the slightest manipulation (e.g., upon attempted transfer to capillary tubes for X-ray studies or even upon removal of excess mother liquor from crystals grown in the capillaries themselves). When the crystals shown in Fig. 1 were examined in polarized light, no optical activity whatever was detected.

B . HOMOGENEITY Purity of the crystalline product was confirmed according to the following criteria: 10. P. Berg, F. H. Bergmann, E. J. Ofengand, and M. Dieckmann, JBC 236, 1726 (1961). 11. G . C. Schito and A. Peace, Gwrn. MiCrobioZ. 13, 145 (1965). 12. J. Josse, JBC 241, 1938 (1966). 13. S. C. K. Wong, D. C. Hall, and J. Josse, JBC 245, 4335 (1970).

20.

INORGANIC PYROPHOSPHATASE OF

E. Cali

503

(1) Usually a single, sharp band was observed when the protein was examined by polyacrylamide or starch gel electrophoresis (14, 15) a t either pH 7 or 9 ( 1 3 ) .Occasionally traces of a slower moving band have been noted, but there is reason to believe that this represents a specific aggregate of pyrophosphatase molecules (see Section II,G and Fig. 4 ) . (2) Only one symmetrical schlieren boundary was observed during sedimentation velocity and diffusion analyses (13). (3) The plot of log (concentration) vs. (radius)', obtained during both low- and high-speed sedimentation equilibrium analyses, was invariably linear across all portions of the solution column which could be examined (13, 16, 17).

FIG.1. Crystals of E . coli inorganic pyrophosphatase. The cubic forms in (a) were obtained in the final step of the routine purification procedure (IS). The large crystals shown in (b) were grown slowly at higher temperature (see the text). 14. B. J. Davis, Ann. N . Y . Acad. Sci. 121, 404 (1964). 15. 0. Smithies, BJ 71, 585 (1959). 16. E. G. Richards, D. C. Teller, and H. K. Schachman, Biochemistry 7, 1054

(Km. 17. D. A. Yphantis, BiochemCtly 3, 297 (1964).

504

J . JOSSE AND S. C. K. WONG

C. SIZE Molecular weight (MW) of the purified enzyme in 0.1 M NaCl, 0.01 M sodium phosphate buffer, pH 7 (NaC1-NaPO4 buffer), has been established by both low- and high-speed sedimentation equilibrium techniques (13, 16, 17). M W (mean) = 121,000 f 2,000 (S.D.); range: 117,000-125,000 (1) With use of the s,”,, and DZOo,wvalues cited below (Section II,D,l) an additional estimate of the molecular weight can be made by means of the Svedberg equation (18); the value obtained is 118,000. [Calculations employ as partial specific volume: V = 0.745 cms/g, obtained from the amino acid composition of the protein given in Table I11 (19).]

D. PHYSICAL PROPERTIES 1. Sedimentation and Diffusion Coefficients

Sedimentation coefficients were obtained in NaCl-NaPO, buffer by standard methods (20), and diffusion coefficients were determined in the same medium both by analyses of schlieren boundary spreading during sedimentation analysis (20) and by tracing the time-course of interference fringe movements during approach to low-speed sedimentation equilibrium (21).Values obtained over a range of protein concentrations (0.5-9 mg/ml) were corrected to standard conditions (water a t 20”) and plotted as reciprocal parameters vs. protein concentration. The straight lines so obtained had the following equations (13): [SZO,~]-’

=

0.1427

+ 2.48~; lo7+ 1 . 9 6 ~ ;

[D20,w]-1 = 0.1755 X

s;~,, = 7.01 S

Die,, = 5.70 X

cm2 sec-l

(2) (3)

where c is the concentration in grams per milliliter. These data indicate hydrodynamic behavior typical of the compact globular class of proteins and equivalent to that of an unsolvated prolate ellipsoid of axial ratio ( a / b ) = 3-4 (22).However, solvation of the protein molecules may give them a more nearly spherical shape in solution. 18. T. Svedberg and K. 0. Pedersen, “The Ultracentrifuge,” p. 39. Oxford Univ. Press, London and New York, 1940. 19. E. J. Cohn and J. T. Edsall, “Proteins, Amino Acids and Peptides,” p. 370. Reinhold, New York, 1913. 20. H. K. Schachman, “Methods in Enzymology,” Vol. 4, p. 32, 1957. 21. K. E. Van Holde and R. L. Baldwin, J. Phys. Chem. 62, 734 (1958). 22. C. Tanford, “Physical Chemistry of Macromolecules,” p. 317. Wiley, New York, 1961.

20.


E . coli

505

2. Viscosity The intrinsic viscosity of the enzyme in NaCl-NaPO, buffer, measured in a long capillary viscometer (ZO), was [.I] = 4.00 ml/g (13). When this value for [ v ] is inserted into the Scheraga-Mandelkern equation (23) together with M W = 120,000, V = 0.745 ml/g, and s& = 7.01 S, a p-factor of 2.31 x loe is obtained. These figures lead to estimates of hydrodynamic behavior equivalent to that of a prolate ellipsoid with axial ratio ( a / b ) = 6-7 (22, 2 3 ) . 3 . Optical Properties

Optical rotatory dispersion of the protein in NaC1-NaP0, buffer showed a Cotton trough a t 230-231 mp with a reduced mean residue rotation [m’] = -3100 deg cm2 decimole-l (13) ; the a-helix content corresponding to this extent of optical activity is 12% (24). Analyses of the visible portions of the dispersion (300-600 mp), according to the treatments of Moffitt and Yang (25) and of Carver, Schechter, and Blout ( 2 6 ) , gave respective &-helix content estimates of 1% and 12% (IS). The presence of a Cotton trough a t 230-231 m,p suggests the presence of P-pleated sheet structure in the enzyme (27, 28), but study of the dispersion a t lower wavelengths (not possible with instruments available to us) will be necessary to corroborate this possibility. Circular dichroism studies in the far ultraviolet region (190-260 mp), kindly provided by Dr. F. R. Brown, revealed a [ O ] value of -9000 deg cm2 decimole-l a t 222 mp. This and the general shape of the circular dichroism trace, in comparison with that observed previously for various mixtures of ahelix, &pleated sheet, and random coil configurations of poly-L-lysine (29), suggest that E . coli pyrophosphatase contains a mixture of all three types of conformations with an estimated a-helix content of about 20%. The presence of P-pleated sheet structure ( > 15%) was also suggested by infrared spectroscopy in D,O, which showed an amide I band a t 1533 cm-l and a shoulder a t 1655 cm-l. The native protein had a typical protein ultraviolet absorption spectrum with Amax = 278 m p (E;:” = 1180 cm2 g-l), A min = 250 mp, and a 23. H. A. Scheraga and L. Mandelkern, JACS 75, 179 (1963). 24. N. S. Simmons, C. Cohen, A. G. Szent-Gyorgyi, D. B. Wetlaufer, and E. R. Blout, JACS 83, 4766 (1961). 25. W. Moffitt and J. T. Yang, Proc. Natl. Acad. Sci. U.S . 42, 696 (1956). 26. J. P. Carver, E. Schechter, and E. R. Blout, JACS 88, 2562 (1966). 27. B. Davidson, N. Tooney, and G. D. Fasman, BBRC 23, 156 (1966). 28. P. K. Sarkar and P. Doty, Proc. Natl. Acad. Sci. U. S. 55, 981 (1968). 29. N. Greenfield and G. D. Fasman, Biochemistry 8, 4108 (1969).

506

J. JOSSE AND S. C. K . WONG

shoulder a t 283 mp (SO). The fluorescence spectrum was similarly unre= 348 mp ( I S ) . markable and showed a smooth emission curve with ,,A

E. ELECTRON MICROSCOPY Electron micrographs of negatively stained preparations of E . coli pyrophosphatase revealed a round object 65 A in diameter (Fig. 2). The same structure was observed whether the protein had been treated with an anionic or cationic stain (Figs. 2a and c) and whether or not i t was previously fixed with glutaraldehyde (Figs. 2a and b) ( I S ) . If this object represents a spherical particle of 6 5 A diam, its calculated volume is 1.44 x 10-19 cm3, and if particle density is equated to the reciprocal of protein partial specific volume ([GI-' = [0.745]-l = 1.342 g/cm3), the particle weight is 1.99 x lO-lS g. This corresponds to a molecular weight of 117,000, which is near that found for the native enzyme in solution (Section 11,C). However, this proposed spherical shape is not in agreement with the unsolvated particle asymmetry deduced from hydrodynamic studies cited in Sections II,D,l and 2. Although there may have been distortions of shape introduced during the staining and drying procedures preparatory to microscopy, such disagreements between structural dimensions seen in electron micrographs and those deduced from physical transport studies are usual, and the discrepancy is often much greater than that observed here ( 3 1 ) .At the present time the exact shape of the native enzyme molecule cannot be described with certainty. When mother liquor from an enzyme recrystallization was stained with sodium silicotungstate at pH 7, most of the forms were indistinguishable from the 65-A structures of Fig. 2. However, there were also a few round objects 130-140A in diameter, most of which appeared to have a central cavity (Fig. 3a). When the pH was lowered to 6, many more of the larger objects, both solid and with cavities, were observed (Fig. 3b), and a t pH 5 nearly all of the forms were of this larger type (Fig. 3c). Uranyl acetate stained preparations at pH 4 also showed the 130-140-A objects almost exclusively, both with and without the ringlike appearance (Fig. 3d). These large forms may represent a specific molecular intermediate with which single enzyme particles and the crystals are in equilibrium. As the pH is lowered, the equilibrium between the 65-A and the 130140-A structures is shifted in favor of the latter ( I S ) . 30. S. C. K. Wong, P. M. Burton, and J. Josse, JBC 245, 4353 (1970). 31. R. C. Valentine, Nature 184, 1838 (1959).

20.


E.

COli

507

FIG.2. Electron micrographs of negatively stained preparations of E . coli pyrophosphatase. (a and b) Particles stained with sodium silicotungstate (4 g/100 ml, pH 7) ; the particles in (b) were first fixed in glutaraldehyde (0.5 g/100 ml). (c) Nonfixed particles stained with uranyl acetate (2 g/100 ml, pH 4).

508

J . JOSSE AND S. C. K. WONG

FIQ. 3. Electron micrographs of negatively stained aggregates of E . coli pyrophosphatase in 1.4 M ammonium sulfate. The objects in (a), (b), and (c) were stained with sodium silicotungstate (4 g/100 ml) at pH 7, 6, and 5, respectively. The forms in (d) were stained with uranyl acetate (2 g/lW ml) at pH 4.

F. SUBUNITS IN 5 M GUANIDINE HYDROCHLORIDE Chemical analyses of the enzyme indicate that cysteine, but not cystine, residues are present (see Section 11,HJ). Therefore all physical studies in the denaturing environment of 5 M guanidine hydrochloride

20.


E . coli

609

(Sg) were either conducted in the presence of 0.01 M dithiothreitol to maintain reduction of these sulfhydryl groups and prevent formation of disulfide cross-links, or the cysteines were first alkylated with N-ethylmaleimide (13). 1. Size

Molecular weight of the denatured, randomly coiled, subunit polypeptide chains was measured during both low- and high-speed sedimentation equilibrium (IS, 16, 1 7 ) . Results indicated a size-homogeneous subunit population with a molecular weight of 18,300-20,600. [The lower figure was obtained from low-speed analyses, where nonideality effects were present resulting from the higher protein concentrations required in this method ; we have reason to believe that in 5 M guanidine hydrochloride this method yields falsely low molecular weight estimates. The higher figure was obtained from six high-speed determinations, in which nonideality was not a significant complication, and is believed h be a more accurate size estimation (IS).] Additional size estimates of the 5 M guanidine hydrochloride subunit can be made from the Svedberg equation (18),using the s:o,w and D & w values cited in Section II,F,2, and from the empirical equations of Tanford and associates (32,SS), which relate s and [.I] values in concentrated guanidine hydrochloride solvents to polypeptide chain length. These calculations yield molecular weights of 21,300, 20,700, and 19,800, respectively, for the E. coli pyrophosphatase subunit (IS). It appears that this enzyme (MW, 120,000) consists of six subunits, each with MW of 20,000. Size homogeneity of the subunits was further confirmed by the presence of a single band in polyacrylamide gel electrophoresis in 8 M urea containing 0.1 M thioglycolate (SO), 2. Sedimentation and Diffusion Coefficientsand Intrinsic Viscosity

These were determined as described in Sections II,D,l and 2 (IS). s & w= 1.30 S

D20,w= 5.63 X lo-' cm2 sec-l [v] = 21.8 cma/g

(4)

(5) (6) The values of these hydrodynamic properties are at the levels expected for randomly coiled, single polypeptide chains of molecular weight 20,000 and devoid of intrachain cross-link restraints (32,33). 32. C. Tanford, Advan. Protein Chem. 23, 121 (1988). 33. C. Tanford, K. Kawahara, and S. Lapanje, JACS 89, 729 (1967).

510


3. Optical Properties Optical rotatory dispersion of the polypeptide subunits in 5 1 1 1 guanidine hydrochloride was similar to that of native protein except that the 230-231 mp Cotton trough was abolished (IS).The ultraviolet absorption spectrum of the denatured chains had a Amax = 276 m p (Ei7m = 1110 cm2 g-l), Amin = 248 mp, and a shoulder a t 282 mp (SO).

G. RECONSTITUTION OF NATIVEENZYME PARTICLES FROM SUBUNIT POLYPEPTIDE CHAINS As shown in the preceding section, in the presence of 5 M guanidine hydrochloride the six subunits of the enzyme were separated from one another and opened up to form randomly coiled, single polypeptide chains. These were catalytically inactive ( 1 2 ) . When the guanidine hydrochloride was removed by dialysis, enzymic activity was restored, provided the protein was in a reducing environment during the dialysis (Table 11).Under optimal conditions (No. 5 of Table 11) there was 8090% recovery of activity. If there was opportunity for persistent disulfide TABLE I1 RECONSTITUTION OF ACTIVE ENZYME FROM SUBUNIT POLYPEPTIDE CHAINSO

No. 1 2 3 4 5 6

Denaturation treatmentb

Renaturation treatmentb

None GuHCl GuHCl GuHC1-DTT GuHC1-DTT GuHCl-NEM

None NaCl-NaP04 ME-NaCl-NaP04 NaCl-Nap04 MENaCl-Nap04 NaCl-Nap01 or ME-NaCl-NaP04

Specific activity (units/mg protein) 2000 that of phosphohydrolase. Supraoptimal levels inhibit, as do all tested concentrations of sodium lauryl sulfate and Tweens 20 and 80. (See also “Lysolecithin,” “Fatty acids,” and “Long-chain fatty acyl-CoA esters,” above) Oxalate, citrate, l,l0-phenanthroline, aside, diethyldithiocarbamate, cyanide, and 8-hydroxyquinoline inhibit, suggesting the metalloensyme nature of the catalyst Inhibits noncompetitively both phosphohydrolase and phosphotransferase activities; inhibition of former potentiated by cetrimide and abolished or significantly ameliorated by deoxycholate, cholate, Triton X-100, or digitonin treatment of microsomes. Cetrimide reduces inhibition by phlorizin of phosphohydrolase Inhibition even a t fiM levels. pH dependent (37) Inhibition noted with Mg2+,Caa+, ZnP+, Cu2+

PI Bicarbonate

Detergents

Metal-binding agents

Phlorizin

(118-120)

(7, is, 16,IY, 89,101,146)

581

22. GLUCOSE-6-PHOSPHATASE TABLE X (Continued) Inhibitors Bile acids and other detergents Fluoride ion Sulfhydryl inhibitors

References

Comments

(See Sections II,A and C for details and references on biphasic effects of these compounds) (18,38) (7, 119) Inhibition noted with p-chloromercuribenzoate, p-chloromercuriphenyl sulfonate, N-ethylmalemide, iodoacetate, and alloxan

11. Inhibition Demonstrated with Glucose-6-P Phosphohydrolase Activity Iodide ion Arsenate Borate

(266) (18) (146)

Silicic acid

(147-149)

Oral hypoglycemic agents

(150-163)

“Phosphataseinactivating system” “Cell sap inhibitor”

(26‘6)

(164)

Inhibition observed with rat liver, brain, and kidney preparations Inhibition, which is reversible, believed to involve monosilicic acid. Inhibition kinetically noncompetitive; mM concentrations effective Inhibition noted with 2-phenylsulfonamido-5-alkyl1,3,4-thiaeole; l-butyl-3-(p-tolylsulfonyl) urea; l-(p-tolylsulfonyl)-% butylurea (U-860) ; and Orinase In cytosol; effects reversed by epinephrine and EDTA Develops in rabbit cell sap fraction on storage; heat stable; high molecular weight nonprotein not further characterized

a Part I from R. C. Nordlie, Ann. N . Y . Acad. Sci. 166,699 (1969). Copyright (1969), The New York Academy of Sciences. Reprinted by permission.

certain detergents. Inhibition by molybdate, for example, appears to be pH dependent (37), for equimolar amounts of this ion inhibited much more effectively a t pH 5.2 (37) than at pH 6.5 (19). Micromolar levels of the ion were found to inhibit a t the lower pH (37), while no inhibition was observed by concentrations less than 6.3 X M a t pH 6.5 (19). As mentioned in Section II,C,3, inhibition of both phosphotransferase and phosphohydrolase activities by citrate (118), as well as other metalbinding agents (119, 120), also is highly pH dependent. Little or no inhibition by the former compound was observed above pH 6.5 (118). Variations in the degree of inhibition with assay pH have suggested to Dyson et al. (141) that HC0,-, and not CO,, is the active inhibitory 141. J. E . D. Dyson, W. B. Anderson, and R. C. Nordlie, JBC 244, 560 (1969). 142. R. J. Paur, Master’s Thesis, University of North Dakota, 1968.

582

ROBERT C. NORDLIE

species. Recent studies by Vianna and Nordlie (104) indicate that inhibition of phosphohydrolase and phosphotransferase activities by Pi is both pH and detergent dependent. As described in Section III,D,l, inhibition by various nucleosidetriphosphates and nucleosidediphosphates, as well as by PP,, of glucose-6-P phosphohydrolase activity is observed over a broad range of pH values, in contrast with phosphotransferase reactions involving these compounds as substrates. The mechanistic significance of these observations is considered in detail in Sections III,D,l and 111,D15.Certain natural and synthetic detergents markedly potentiate this inhibition of glucose-6-P phosphohydrolase activity by these compounds. Specific details relating to this phenomenon are given in Section II,C,3. Detergents also exert interesting, activity-discriminating modifying effects on the inhibition by phlorizin of phosphohydrolase and phosphotransferase activities of the enzyme, as described in detail in Section II,C,3. Clearly, the interactions of inhibitors with this multifunctional, particulate enzyme are highly complex and deserving of further detailed experimental consideration. The need for caution in reaching conclusions relating to the significance of inhibitor action without thorough exploration of the phenomenon from a variety of angles is also pointed up by the observations described above. 5 . Reaction Mechanism

a . Mode of Enzyme-Substrate Interactions. Detailed kinetic studies relating to the reaction mechanism of glucose-6-P phosphohydrolase have been carried out by Segal (31) and by Hass and Byrne (69,SO), who 143. R. C. Nordlie, T. L. Hanson, and J. D. Lueck, unpublished observations (1970). 144. T. L. Hanson, Doctoral Dissertation, University of North Dakota, 1968. 145. T. Fossa, Boll. SOC.Ital. Biol. Sper. 38, 903 (1962). 146. C. Zerr and W. B. Novoa, BBRC 32, 129 (1968). 147. N. C. Ganguli, B. S. Arora, and V. R. Bhalerao, Indian J . Exptl. Biol. 1, 228 (1963). 148. K. Krisch, 2.Physiol. Chem. 314, 211 (1959). 149. W. Kersten, K. Krisch, and H. Staudinger, Wks. Forschungsber., N a t u m . Reihe 66, 117 (1958). 150. R. Jasmin and W. Johnson, J . Am. Pharm. Assoc. 48, 113 (1959). 151. G. Mohnike and W. Knitsch, Naturwissenschaften 43, 449 (1958). 152. G. Mohnike, K. W. Knitsch, H. Boser, G. Werner, and S. Werner, Deut. Med. Wochschr. 82, 1514 (1957). 153. G. Weber and A. Cantero, Metab. Clin. Ezptl. 7, 333 (1958). 154. T. R. Ricketts, Exptl. Cell Res. 34, 557 (1964).

683

22. GLUCOSE-6-PHOSPHATASE

also included a consideration of catalysis of 14C-glucose:glucose-6-P exchange and glucose-6-P: fructose phosphotransferase activity of the enzyme. Arion and Nordlie (4,41) have extended these initial studies to include PPi-glucose phosphotransferase, mannose-6-P :glucose phosphotransferase, and nucleotide-glucose phosphotransferase activities of the enzyme. On the basis of the observed phosphotransferase reaction, of the apparent noncompetitive inhibition of phosphohydrolase reaction by glucose (which actually is a manifestation of phosphotransferase activity), competitive inhibition between glucose-6-P and various phosphoanhydride substrates, and of the variations of activity with substrate concentrations in the phosphotransferase reactions as described in Fig. 6 and Section 1II1D,2, the reaction mechanism described in Fig. 4 has been proposed (40, 4 1 ) . This mechanism incorporates both phosphohydrolase and phosphotransferase activities of the enzyme. The left half 5, Fig. 4) describes glucose-6-P of the diagram (reactions 4 + 3 phosphohydrolase activity of the enzyme, as proposed both by Segal (31) and Hass and Byrne (SO), while reactions 1 2 3 in Fig. 4 describe the hydrolysis of PPi, nucleosidetriphosphate or nucleosidediphospkate. In both instances the first step in the process involves formation of binary enzyme-phosphoryl substrate complexes, which then dissociate to yield a common phosphoryl-enzyme intermediate. The existence of such an intermediate has also been suggested by Hass et al. (155) on the basis of their observation that the enzyme, when allowed to hydrolyze glucose-6-P in H2180in the presence of excess glucose, did not catalyze an incorporation of water l80into glucose-6-P concomitant with the glucose incorporation. Transfer of the phosphoryl group from the phosphoryl-enzyme intermediate to water (reaction 5 in Fig. 4) completes the hydrolysis process. Alternatively, the transfer of the phosphoryl group to glucose, for example, leads to the production of a binary enzyme-glucose-6-P complex which ultimately dissociates (reaction 4, Fig. 4) to yield glucose-6-P and free enzyme. Thus, reactions 1 2 3 4, Fig. 4, constitute phosphotransferase activity of the enzyme. Reactions 4 3 plus reversal of reactions 3 4 depict the exchange reaction observed by Segal (31) and by Byrne and Hass (SO), and also explain the apparent competitive inhibition by glucose of the hydrolysis of glucose-6-P observed by these workers and by Arion and Nordlie (4). Applicable rate equations, with accompanying definitions of kinetic expressions, for the various activities of the partially purified rat liver microsomal enzyme a t pH 6.0, derived on the basis of the mechanism in Fig. 4, are as follows (40) (to simplify notation, k , x [ H 2 0 ] is set =

+

+ +

+

155.

+ +

+

+

L. F. Ham, P. D. Boyer, and A. M. Reynard, JBC 238, 2284

(1961).

584

ROBERT C. NORDLIE

k,' since H 2 0 concentration may be presumed relatively very high and constant (40) ) (i) For phosphotransferase activity with R P (PP,, mannose-6-P, nucleosidetriphosphates or nucleosidediphosphate, etc.) as phosphoryl group donor:

.

Vmax(Trf) --

- 1 + - KRP +-

K G I ~ KRP-GI~ (Glc) (RP)(Glc)

(RP)

V

(5)

where

KRP= =

KGIC

KRP-OI~ = and Vmax(Trf)

=

Here, E, stands for total enzyme concentration and the subscript "Trf" indicates the RP-glucose phosphotransferase reaction. (ii) For glucose-6-P hydrolysis in the absence of glucose: Vmax(Glc-6-Paee)

-

KGIUC-6-P

V

-

(Glc-6-P)

where

and .

.

.

(iii) For hydrolysis of R P ( PPI, mannose-6-P, nucleosidetriphosphate or nucleosidediphosphate, etc.) in the absence of glucose :

where

and

22. GLUCOSE-6-PHOSPHATASE

585

(iv) For glucose-6-P hydrolysis in the presence of glucose: Vmax(Glc-6-Paae) 2,

-

KGIc-6-P

(Glc-6-P)

( g) I i!I +

Here, I stands for glucose, Kolc,-p is as defined in Eq. (6a), Vmax(Glc+ Pase) is as defined in Eq. (6b), and

(v) For RP hydrolysis in the presence of glucose:

where K R Pis as defined in Eq. (7a), Vmax(RPase) is as defined in Eq. (7b), and

Reactions 2 and 3, Fig. 4, involving the formation of phosphoryl enzyme through dissociation of binary enzyme-phosphate substrate complexes, are the rate limiting steps in the various reactions catalyzed (40). Inability of such complexes to dissociate when composed of enzyme and PE-, ADP3-, or ATP4- has been considered in Section 111,DJ and will be further considered in Section III,D,5,c. Assuming that the rate-limiting steps in the overall reaction processes involve the dissociation of binary enzyme-phosphoryl substrate complexes to yield the phosphoryl-enzyme intermediate (31, 40) (reactions 2 and 3 in Fig. 4), it follows that k , k,, k , k,, ks k,, k , Q k,, and k , k,. By applying these inequalities to Eqs. (5a), (5b), (6a), (7a), (8a), and (9), above, it may be shown, consistent with experimental observations (37, 40, 41, 43, 45, 46, 155a) that a t any pH where activity is manifest K , for R P in the phosphotransferase reaction [Eq. (5a)l = K , for R P in the phosphohydrolase reaction [Eq. (7a)l = K I for R P as a competitive inhibitor of glucose-6-P phosphohydrolase = k , / k , ; that K , for glucose-6-P [Eq. (6a) J = K , for glucose-6-P acting as a competitive inhibitor of R P hydrolysis = k , / k , ; and that K , for glucose in the phosphotransferase reaction [Eq. (5b)I = K , for glucose functioning as an apparent noncompetitive inhibitor of phosphohydrolase activity with glucose-6-P [Eq. (8a)l or RP [Eq. ( 9 ) ] = k , / k , ( 1 5 5 ~ ) . From the mechanism depicted in Fig. 4 it is apparent that glucose-6-P and various phosphoryl donors should compete for a common site on

k

40

-I

20 v) v)

W

sa o 0.5

0

-

I

I

1

I

I

1

I

1

1

I

I

I

I

I

1

I

I

I

I

I

I

I

I

I

200

w W

0

*

100

0 -100

I

FIG.5 (a)-(d).

24.

661

BOVINE PANCREATIC RIBONUCLEASE

I

I

I

I

I

I

I

I

I

I

I

I

I

.I

r

I

1

I

1

I

I

I

I

I

I

1

I

I

I

I

i

I

I

1

1

I

I

I

I

I

175

150

2 3 125 6 w E 0 100 E E

75

50 W

- 150

4'

125

0

a a

-

100 75

FIG.5. Various parameters of accessibility, twist, and bend plotted vs. sequence number. Part 1: (a) Solvent-accessible area of side chains. (b) Fractional accessibility (referred to full sphere) of backbone carbonyl oxygen and peptide nitrogen. The separate plot for values less than 1% is meant to show that no accessibility was detected for many atoms. The actual nonzero values are not to be taken too literally. Part 2: (c) Backbone anglcs as normally defined. (d) Angles between sequentially adjacent carbonyl vectors in the backbone plotted between the sequence numbers of the two residues involved. Part 3: (e) Distance in A between the tips, T, of adjacent residues as defined in the text. ( f ) Distances in A between peptide center, M, and the third sequcntial pcptide center (open circles), and between carbon a and the sixth sequential a-carbon (crosses) plotted opposite the central carbon atom in each case. (g) Angles between lines joining the centers of successive peptide bonds plotted between the residues defining the central bond. (11) Angles between lines joining successive a carbons plotted opposite the central carbon. (Note that the accessibilities were calculated with coordinate set 4 and the other parameters with set 6 ; see text.)

*

TABLE I11 STATIC ACCESSIBILITIES OF ATOMSOF RIBONUCLEASE-S" ,* OlY 68 88 112

CA MN MC MO M A I N NPS PS

CYS

CA MN MC MO CB SG M A I N NPS PS

25 0 33 13 27 7 AVG 28 6

3 44 6 32 5 5 5 27

2 6 0 0 0 0 40 0 3 0 1 6 5 8 0 0 + + 6 5 0 0 1 3 7 2 0 0 0 0 8 4 0 0 0 0 9 5 0 3 + 2 1 1 0 0 0 0 + AVG 0 1 0 3

18 21 11

0 2 + 0 0 0 0 0 0

0 1 * 3 0 0 7 0 1

0 5 0 1 0 0 1 0

0 1 0 1 0 0 4 0

CA MN I,1C MO CB M A I N NPS PS

ALA

2 0

7 0 034 1 1 232 6 1 1 + 133 5 2 1 2 + 1 5 1 5 6 7 + 0 1 3 1 64 0 4 + 19 49 9 6 + 0 0 0 3 0 102 1 0 + + 35 109 0 0 0 0 10 122 5 1 0 0 21 AVG 2 2 0 2 33 4

5

PRO

42 93 114 1 1 AVG

2 34 1 3 2 0 33 1 5 1 2 3 1 6 49 0 3 0 0 35 0 10 2 21

CA MN MC td0 CB CG CD M A I N NPS PS 1 26 + 0 0 2 29 37 12

1 0 2 0 25 31 3 0 0 5 28 31 7 0 0 0 2 0 5 1 0 0 2 20 26

21 34 0 17

1 2 0

25 31 2

CA PIN MC MO CB G 1 G2 N A I N NPS PS 43 0 0 + 5 1 2 4 2 8 1 1 5 0 0 4 7 0 0 0 0 0 - + 5 4 0 0 0 * 0 - + 0 0 5 7 0 0 0 0 0 + 0 0 0 63 5 0 + 0 5 2 1 6 3 1 1 1 0 0 1 0 8 O O O O O + * 116 0 0 0 0 0 1 4 6 0 7 1 1 8 0 0 0 6 + 2 5 2 2 AVG 1 0 0 1 3 5 5

VAL

LEU

CA MN E1C MO CB CG D1 D 2 M A I N NPS PS 3 5 + 0 0 0 4 0 0 2 0 2 51 + 5 0 0 2 6 2 5 3 9 1 1 8 AVG 0 2 0 0 15 1 2 21 CA MN MC MO CB G 1 G2 CD M A I N NPS PS 8 1 0 0 0 + + 0 1 4 0 1 0 0 1 0 6 0 0 0 0 0 + * * 107 0 0 0 0 0 1 + 2 6 0 7 AVG 0 0 0 0 0 0 0 1 0

ILE

CA MN t.IC MO CB CG SD CE M A I N NPS PS 1 3 0 0 3 2 5 3 + 0 1 2 2 9 0 + 0 0 0 4 2 1 0 0 G 3 0 0 0 0 0 0 0 0 0 0 0 7 9 0 0 1 0 0 6 1 2 0 2 AVG 0 0 1 1 1 3 5 1

MET

TABLE I11 (Continued) CA MN M C MO CB CG D1 El CZ E2 02 MAIN NPS PS 0 0 0 0 0 0 0 0 + 2 2 0 1 0 0 4 6 0 0 0 0 0 0 0 0 0 0 0 1 2 0 0 0 + 1 5 1 0 6 1 1 * 0 4 1 AVG 0 0 0 5 0 0 2 0 0 1 1

PHE

8

SER CA MN MC MO CB OH MAIN NPS 32 6 t 1 19 37 10 6 37 50 3 0 0 0 2 5 5 1 2 5 59 7 + 0 27 35 38 8 35 0 0 7 5 0 0 0 0 0 2 77 3 t 0 1 29 38 1 29 80 0 0 0 0 10 21 0 10 89 9 6 0 3 14 47 4 14 3 0 9 0 0 0 0 1 3 t O 123 0 0 t 12 9 31 3 9 AVG 3 1 0 8 17 21

PS 10 5 38 2 38 21 47 0 31

THR

CA MN M C MO CB CG2OH MAIN NPS PS 4 0 4 0 16 38 18 2 27 18 0 3 0 3 6 0 0 0 1 1 4 0 4 S 0 + 0 0 0 1 3 2 0 6 2 5 7 0 0 1 + 1 6 1 5 0 5 4 2 6 3 7 34 78 0 t + 13 0 13 34 82 0 0 0 0 0 * 1 2 0 0 12 8 7 3 0 5 1 4 8 1 6 9 5 1 2 9 9 9 0 1 0 2 5 0 3 2 2 7 1 2 2 1 13 32 100 5 0 1 0 12 13 32 AVG 1 0 1 8 4 17 15 3

CA MN M C M O CB CG D1 El C Z E2 02 OH MAIN NPS PS 25 2 2 0 0 2 0 1 2 + 1 2 1 2 2 + 1 1 0 0 7 3 0 0 0 0 0 0 0 0 0 7 0 2 3 0 1 2 3 6 11 48 76 2 0 t 23 15 2 11 10 3 20 14 48 6 8 41 92 0 2 0 2 2 1 6 2 1 7 1 7 1 2 + 4 1 0 0 9 7 0 0 0 8 0 0 0 0 0 + 0 0 2 115 0 2 t 11 16 1 7 15 3 9 14 18 3 9 18 AVG 1 1 0 11 11 1 6 7 1 10 8 22

TYR

ASP CA M N MC MO CB 14 0 0 0 1 2 0 38 t + 0 27 30 5 3 t 0 0 2 1 2 8 3 0 0 0 0 2 1 2 1 1 0 2 2 0 AVG 0 0 0 9 7

CG 01 02 MAIN NPS PS 0 410 3 0 7 1 1 49 7 15 25 1 9 4 7 1 2 3 3 0 1 1 8 0 3 7 + 0 2 2 3 1 0 1 3 1 13 26

ASN CA MN M C MO CB CG X1 X2 MAIN NPS PS 24 10 1 0 + 10 2 21 10 3 6 16 2 7 0 0 0 + 6 1 1 4 + 0 3 7 3 4 0 0 0 6 3 5 2 6 3 2 1 4 2 9 0 0 0 4 4 2 0 0 0 0 0 1 0 62 0 1 + 22 21 3 33 5 6 12 19 5 32 6 7 4 0 t 38 8 2 44 20 10 0 4 1 0 7 1 + 0 0 0 8 0 1 8 1 8 13 28 9 4 14 5 3 11 25 1 56 0 1 0 3 0 * 0 9 7 3 4 9 2 4 2 5 3 7 113 0 6 + 33 25 2 41 64 10 14 52 AVG 3 1 0 12 11 2 30 16 GLU

2 9 49 86 111 AVG

CA M N M C MO CB C G CO 0 1 0 2 MAIN NPS 0 + 2321621 1 7 1 8 12 1 4 0 0 3 + 2 8 14126 0 0 0 0 0 0 0 0 02025 6 2 0 0 t25 4 0 1 9 2 7 3 6 1 0 3 9 18 0 1 37 25 0 0 2 13 8 6 1 23 21

PS

4 33 23 18 31

664

F. M. RICHARDS AND H . W. WYCKOFF

TABLE I11 (Continued) GLN CA MN MC MO CB CG CD X 1 X2 M A I N N P S PS 0 0 3 1 1 0 0 0 0 0 0 1 5 2 28 1 0 0 1 4 1 4 3 8 5 7 0 7 32 1 4 27 55 0 0 1 3 9 1 1 2 2 3 2 60 3 0 + 4 0 3 1 1 9 1 5 2 2 17 69 5 0 + 0 8 0 * 3 4 3 5 1 3 34 0 0 1 4 7 4 0 0 0 0 + 0 0 2 9 0 3 1 4 33 1 0 1 0 1 t 1 2 5 27 11 23 4 4 AVG 1 0 0 3 4 7 2 2 0 2 6 CA MN MC MO CB CG N1 C 2 N 3 1 2 0 0 0 0 0 0 0 4 0 4 8 0 0 0 4 t l t 7 5 105 0 0 0 0 0 0 1 1 3 5 8 1 1 9 0 0 0 t 1 7 2 5 5 25 AVG 0 0 0 1 4 1 4 1 3 9

HIS

LYS

7 31 37 41 61 66 91 98 104 AVG

CA MN MC MO 0 0 0 0 0 0 + 0 8 2 1 7 0 0 0 0 7 + 0 0 6 1 1 27 0 0 0 4 4 0 0 0 0 0 0 0 3 0 0 4

CB CG CD 0 + 2 3 t 1 6 29 20 1 2 0 0 3 0 1 1 5 0 11 1 8 1 8 1 4 29 1 0 8 2 29 1 1 2 22 6 12 14

ARC CA MH MC MO CB CG CD 10 5 t 0 t 1 7 7 9 33 1 0 0 7 0 0 1 1 39 4 0 0 0 1 2 0 7 85 0 0 0 0 4 + 2 0 AVG 2 0 0 2 8 2 1 2 *** END OF DATA S E T ***

CE 1 2 15 20 3 0 1 9 17 39 19 3 19

C4 M A I N NPS PS 0 0 1 0 3 t 1 2 0 0 5 18 16 0 12 10 4

NZ M A I N 0 5 0 0 35 4 36 1 4 0 3 9 2 9 67 1 54 1 49 31 0 42

NPS 9 15 13 8 9 16 23 14 9

PS 50 35 36 14 39 67 54 49 31

NE CZ N 1 N2 M A I N NPS PS 1 0 0 1 2 2 4 8 12 2 3 17 1 8 2 1 0 2 2 2 1 1 5 7 2 2 1 5 33 7 1 5 5 5 3 0 6 38 12 1 33 30

==>

a The accessibility of each atom was calculated as discussed in the text. The numbers given are areas of the hydrosphere surface defined as the locus of possible centers of solvent molecules in contact with the given atom or atom group (CHZ and CHa). The calculated areas in k have been rounded off to the nearest integer. Values between 0 and 0.5 have been indicated by a and atoms contacting the internal void have been indicated with an “*”. Atom designations are standard except that three or four character designations have been shortened, e.g., CG1 is indicated as G I . Single character main chain designations have been prefixed with M. The sums of the areas for the main chain atoms (MAIN), nonpolar side chain atoms (NPS), and polar side chain atoms (PSI are given, and the average area for each atom in a given amino acid is tabulated. Residues 1, 17 through 23, and 124 are omitted from the tabulation although they were included in the calculation. * The following comments refer to the fit of the model to the electron density map. The uncertainties discussed should be considered in any use of the accessibilities listed in Table 111. The backbone peptide chain is clearly defined in regions of high electron density except as indicated below. The carbonyl oxygen atoms are frequently, but not always, visible as “bumps.” In some areas it is not possible to get good fits for the oxygen positions without changing some of the main chain bond angles. The present “best fit” model has not yet involved an intentional change of any of these angles.

“+”

24.


665

Footnotes to Table I11 (Continued) The chain and most of the associated side chains are not well defined in the following regions: residues 2,65-72, and 119-123. The chain is very poorly defined or not visible a t all in regions: residues 1, 18-20,21-23, and 124. Except for the small peptide loop, these regions are all associated with the chain ends. Cursory inspection of the sums of the electron densities in the backbone and 8-carbon positions shown residues 3 through 9 to be systematically low, and preliminary calculations show that a simple shift of these residues by 0.6 A in the crystallographic z direction would bring these sums into line with those for the remainder of the structure. Alanine. Residues 4, 5, 56, 96, 102, and 109: CB clear in positive, but perhaps weak, electron density. Residues 6,52,64, 122: CB outside of lowest electron density contour or poorly defined. Residue 5: carbonyl direction apparently pointing somewhat away from helix axis with no obvious explanation. Residues 19 and 20: uncertain region of very poorly defined chain. Arginine. Residue 10: fair fit only; plane of guanidino group not clear; possible confusion of one lobe with solvent; alternate position possible. Residues 33, 39, and 85: clear and well defined. Asparagine. Residues 24, 27, 44, and 62: clear and well defined. Residues 34, 103, and 113: plane of terminal group not well defined. Residues 67, 71, and 94: not clear beyond CB; mostly positive density but poorly defined. Aspartic Acid. Residues 14 and 121: groups in positive electron density, but latter formless and somewhat spread out. Residues 38 and 53: clearly defined except for plane of terminal group of 53. Residue 83: not defined beyond CB. Cysteine (8-8). Residues 26-84 and 40-95: well defined and clear, chirality of S-S bonds clearly left-handed; S atom in 84 needs slight shift. Residues 58-110: electron density reasonably clear, chirality probably left-handed. Residues 65-72: electron density much broader than for the other three S-S bonds; chirality not completely certain, but probably righehanded. Glutamine. Residues 11,55, 60, and 74: clear and well defined. Residue 28: not defined beyond CB. Residue 69: poorly defined beyond CG; could be shifted to area of higher density. Residue 10: plane of terminal group not defined. Glutamic Acid. Residues 9 and 49: clear and well defined; CB of 9 slightly above positive density. Residue 86: plane of terminal group not well defined; no forking. Residues 2 and 111: weak electron density, poorly defined beyond CB. Glycine. Residues 88 and 112: clearly in positive electron density. Residue 68: weak electron density a t CA position (area of uncertain chain, see above). Histidine. Residue 12: electron density “ball” not flat enough to clearly define the plane of the imidazole ring. Residues 48 and 105: clearly defined but some ambiguity in plane of 105 ring. Residue 119: possible fit shown, but the ‘ ball” of electron density is almost spherical and is somewhat larger and denser than the other histidine residues or than would be expected for a 5-membered ring; alternate positions are possible with some reorientation of the chain on both sides of 119. Zsoleucim. Residues 81, 106, and 107: clear and well defined. Leucine. Residue 35: clear and well defined. Residue 51: electron density weak but terminal fork fairly clear. Lysine. Residues 7, 61, and 66: clear and well defined. Residue 41: weak electron density but probable position reasonably clear. Residues 37,98, and 104: electron density negative, weak or poorly connected beyond CB. Residues 31 and 91: not defined beyond CA. Residue 1 : uncertain; region of poorly defined chain. Methimine. Residues 29 and 79: clear and well defined, although S peak of 79 has low density even with slight shift and S C D direction of 29 is not defined. Residue 13: alter-

666

F. M. RICHARDS AND H. W. WYCKOFE’

Footnotes to Table I11 (Continued) nate position of CB and CD possible. Residue 30: CB-CG connection weak; S-CD direction reasonably clear. Phenylalanine. Residues 8 and 46: clear and well defined; CB of 8 just above region of positive density. Residue 120: C.4 and CB clear but ring poorly defined, density weak and spread out. Proline. Residues 42 and 117: rings clearly defined, both in trans conformation. Residues 93 and 114: rings clear, both shown in cis conformation; reorganization of the tight loop does permit a fit with 114 trans, but the fit is not quite as good as in the cis conformation and 113 would be in radically different position; alternate fit for 93 also possible but not as satisfactory. Serine. Residues 16, 59, 75, 80, and 90: clear and well defined. Residues 32 and 77: clear but two possible positions for OH group. Residues 15, 50, 89, and 123: not well defined OH in weak or negative electron density. Residues 18, 21, 22, and 23: uncertain, regions of poorly defined chain. Threonine. Residues 3, 36, 70, 78, 82, and 100: dear and well defined. Residues 17, 45, 87, and 99: positive electron density but forking not clear. Tyrosine. Residues 73, 92, and 115: clear and well defined. Residues 25 and 76: ring electron density irregular in shape, but OH density clear. Residue 97: OH slightly outside of electron density bump; movement difficult to keep CB, ring and OH all in positive density. Valine. Residues 43, 47, 54, 57, 63, 108, 116, and 118: clear and well defined. Residue 124: uncertain, region of poorly defined chain.

chain atoms with a radius of 1.4A for the solvent while a hydrogen bond distance should be 2.8-3.0 A and a 2-A radius for the methyl and methylene groups might be a better average value. Any pattern in the side chain accessibility as a function of sequence is difficult to see. As a direct consequence of the structure of helices the backbone N and 0 are relatively inaccessible to the solvent except a t the ends. Extended structures would tend to have alternate bonds exposed if any. The static accessibilities calculated as noted above are plotted in Fig. 5b as a percentage of a full sphere. Since very low but finite accessibility can be significant, values less than 1% are plotted on an expanded scale on an auxilliary ordinate. The calculation is not necessarily as precise as implied by this scale. It is easily noted that the backbone nitrogens are exposed much less than the pendant carbonyl oxygens. Only 17 nitrogens are exposed more than 1% as opposed to 52 oxygens, while there are 11 additional nitrogens above the 0.1% level and 6 oxygens in this category. The published hydrogen bonding scheme (62) has been slightly modified in the latest adjustment of the model to the 2-A map, M30, based on 6000 reflections. Many side chain bonds to the backbone have been assigned, and a number of solvent molecules have been tentatively located. These are indicated schematically in Fig. 6a. The number of (Y

24.


667

backbone-backbone bonds assigned in the a-helical scheme is 16 while 3 or 4 bonds are in a short 3,, section and approximately 26-35 bonds are in the p scheme. A number of bonds a t ends of helices are made to side chains. There is one reverse turn a t 34. Some of the groups with unsatisfied intramolecular bonds are actually bonded to adjacent molecules in the crystal, some additional bonds may be assigned in the future, and some are clearly left unsatisfied according to the electron density map. It should be noted that the accessibility calculations were based on coordinate set 4 (62) while the hydrogen bonding scheme of Fig. 6a includes some revisions. For example, the bonding in the 50-58 helical region has been changed to include the insertion of a solvent molecule a t 53 and a bond to the Ser 50 side chain. The 55-58 bond was accidentally shifted, and the 56-59 bond is questionable. In the active center the motion of His 119 exposed the backbone nitrogen of 120 and it bonds to the sulfate ion in the pl site (see Section VI). Also note that residues 1, 16-23, and 124 were included in the model during the accessibility calculations but are omitted from this tabulation since their positions are uncertain and the calculated parameters would be meaningless. The bending and twisting of the backbone, calculated from the most recent coordinate list (unpublished), is indicated in Fig. 5 in several ways. The twist is indicated by the angle between successive carbonyl bond vectors in Fig. 5d. These bonds are nearly parallel in a helices and antiparallel in J structures. The Ik angle of the Ramachandran plot is the one parameter most indicative of the a and p regions (omitting left-hand helices), and this is also plotted vs. sequence number in Fig. 5c. Four parameters are graphed indicating the bending and looping of the backbone. The angle between lines joining sequential a-carbon atoms is plotted opposite the residue number of the central atom in Fig. 5h. Haas (72) has used this angle for model building and lists the values for several proteins. The angle for fully extended chains is only 155". If one chooses the centers of peptide bonds, M , instead of the a carbon the angle can be 180". The M M M angles are plotted in Fig. 5g between the residue numbers bracketing the central peptide. Progressive curvature producing loops and gross bends and reverse curvature producing nearly extended regions in spite of local curvature are best indicated by distances between sequentially more remote reference points. The distance between terminal M's of M a M a M a M is plotted above the residue number of the central a carbon and the distances between terminal a carbons in a M a M a M a M a M a M a are plotted similarly in Fig. 5f. 72. D. J. Haas, "Three Dimensional Models of Protein Molecules," manual. Electronics and Alloys Inc., Englewood, New Jersey, 1969.

668

F. M. RICHARDS AND H . W. WPCKOFF

I

-120

-

I

I

0

1

60

___---.--.

(-t\

1

0

$

(b)

FIG.6. (a) Backbone hydrogen bonding scheme as currently delimited. Disulfide bridges are indicated in heavy dashed lines and poor or less certain bonds in light dashed lines. W indicates solvent molecule. Side chain to side chain bonds are not generally indicated. The scheme of bonding currently assigned for 3'-CMP is also

24.

669


Sequentially adjacent side chains may or may not be near each other as seen in the plot of the distances between successive terminal reference points, T, in Fig. 5e. For carboxyl, amide, valine, leucine, and arginine side chains, T was defined as the center between the two terminal atoms. I n proline C, was chosen, in methionine the sulfur, in threonine the oxygen, and in histidine the point between C, and N,. A Ramachandran plot of the dihedral angles and is shown in Fig. 6b. The outline of the allowed area is taken from Fig. 38a in Ramachandran and Sasisekharan (7'2cz).The only residue well outside of this contour is 60. If the three-dimensional structure of rat RNase is assumed to be identical to that of the cow, it is seen that all of the 41 changes in sequence are sterically permitted. There is a curious tendency for the changes to occur in pairs so that the charge distribution on the surface is maintained. The active site is invariant, however. Wyckoff (7'2b) has discussed in detail the sequence changes in relation to the structure of bovine RNase-S.

+

IV. Modiflcation of Covalent Structure

A. ENZYMIC CLEAVAGE OF THE MAINCHAIN A summary of the products is given in Table IV (25, 28, SO, S4,7S-77'). 1. Subtilisin (RNase-S)

The proteinases of various strains of B . subtilis will cleave the peptide chain of native RNase-A. Although the eventual extent of proteolysis 72a. G. N. Ramachandran and V. Sasisekharan, Advan. Protein Chem. 23, 283 (1968). 72b. H. W. Wyckoff, Brookhaven Sump. Biol. 21, 252 (1968). 73. F. M. Richards and P. J. Vithayathil, JBC 234, 1459 (1959). 74. W. A. N e e , JBC 240, 2900 (1965). 75. C. B. Anfinsen, JBC 221, 405 (1956). indicated partially in the upper right with several residues repeated and several solvent molecules indicated. A portion of this scheme is also included a t the lower left. The scheme for (Y helix, 310 hrlix, and /3 structures are self-explanatory. (b) Ramachandran plot of backbone torsion angles 9 and \k with energy level contours (7%) superimposed. The axes are extended beyond 360" and several residues are repeated, once with a solid circle and once with an open circle. The glgcine residues are labeled G. Residues 1, 16-24, and 124 are omitted since they are very poorly defined in the electron density map. The calculations are based on coordinate set 6.

670


TABLE IV NATURALLY OCCURRING FORMS OF PANCREATIC RIBONUCLEASE AND WELLDEFINEDMAIN CHAINCLEAVAGE PRODUCTS Activity ~~

Preparation and/or abbreviation

Primary structure description (residues)

(8)

~

RNA

C > P (U > p)

Ref.

Isolated from Tissue Extract RNase-A RNase-B des-Lysyl RNase-A des-Lysylpyroglutamyl RNase-A

1-124 as a single chain 1-124 oligosaccharide 2-124 pyroglu-3-124

+

100 100 100 60

100 100 99 38

100 0 0

100 0 0

100

100

0

0

25 30

14 25

12

-

Products of Proteolysis Subtilisin RNase-S S-Protein S-Peptide Elastase RNase-E Pepsin PIR~ Trypsin Component 3 Component 4 Component 5

+

1-20(21) 21(22)-124 21(22)-124 1-20(21) 1-19

+ 21-124

1-120

+ 34-124 + 32-124 and 1-33 + 34-124 1-31 + 38-124 1-31 1-31

Here PIR stands for pepsin-inactivated ribonuclease.

is extensive (78), approximately 30 of the 123 bonds, one or two bonds are cleaved very much more rapidly than others resulting in the accumulation of an intermediate designated RNase-S. A description of the preparation and initial characterization of this material is given by Richards and Vithayathil (73). These authors concluded that no amino acids were lost in the conversion of RNase-A to RNase-S and that a single bond between residues 20 and 21 had been cleaved. A later study by Ottesen and Szekely (79) found no evidence of any other catalytically active component during later stages of digestion, and it must thus be concluded that the next bond split causes inactivation. More detailed 76. T. Oi, J. A. Rupley, and H. A. Scheraga, Biochemistry 2, 432 (1963). 77. T. Oi and H. A. Scheraga, Biochemistry 3, 641 (1964). 78. F. M. Richards, C o m p t . Rend. Trav. Lab Carlsberg, Ser. Chim. 29, 322 (1955). 79. M. Ottesen and M. Szekely, Compt. Rend. l'rav. Lab Carlsberg 32, 319 (1962).

24.


671

studies by Doscher and Hirs (80) and by Gross and Witkop (81) indicated that either the 20-21 bond or the 21-22 bond were broken and that the product ratio depended on the particular proteinase preparation used for the digestion. The term RNase-S will be used to refer to the collection of products without regard to the particular distribution of bonds cleaved. As will be seen later, no confusion is ordinarily introduced because of this uncertainty. Ribonuclease-S can be separated into S-peptide [residues 1-20 (21) ] and S-protein [residues 21 (22)-1241 by precipitation with trichloroacetic acid (73) or better, Sephadex chromatography in 5% formic acid (82). The best preparations of these components show no detectable hydrolytic enzymic activity and little if any transphosphorylation activity (see Section VI) . Isolated S-peptide appears to have no regular secondary structure (83, 84) or 10-2076 helicity (86, 86). (These slightly different interpretations are based on almost identical C D data.) When equimolar amounts of S-protein and S-peptide are mixed a t neutral pH and room temperature or below, essentially full catalytic activity is recovered (73, 87). A schematic diagram is shown in Fig. 7. For a detailed summary of the preparative procedures see Doscher (88). Potts et al. (89) have shown that the 5 C-terminal residues of S-pepticre can be removed with carboxypeptidase. The resulting derivative (residues 1-15) forms a strong complex with S-protein having full catalytic activity. It is clear from the X-ray structure that these 5 residues interact little, if at all, with any part of S-protein, and they are remote from the active site. The various changes produced in this component by synthesis and by chemical modifications are discussed later. Carboxypeptidase action a t 25” on S-protein removes Val 124 very rapidly with no effect on the RNA activity regenerated with added Speptide (90). Further digestion removed Ser 123 with an activity drop to 4576, but the peptide-protein binding constant changed very little. More 80. M. S. Doscher and C . H. W. Hirs, Biochemistm 6,304 (1967). 81. E. Gross and B. Witkop, Biochemhy 6, 745 (1967). 82. E. Gross and B. Witkop, BBRC 23, 720 (1966). 83. A. Scatturin, A. M. Tamburro, R. Rocchi, and E. Scoffone, Chem. Commun. No. 24, p. 1273 (1967). 84. A. M. Tamburro, A. Scatturin, R. Raniero, F. Marchiori, G. Borin, and E. Scoffone, FEBS Letters 1, 298 (1968). 86. W. A. Klee, Biochemistry 7, 2731 (1968). 86. J. E. Brown and W. A. N e e , Biochemistry 8, 2876 (1969). 87. F. M. Richards, Proc. Natl. Acad. Sci. U.S. 44, 162 (1958). 88. M. S. Doscher, “Methods in Enzymology,” Vol. 11, p. 640, 1967. 89. J. T. Potts, D. M. Young, and C . B. Anfinsen, JBC 238, 2593 (1963). 90. J. T. Potts, M. Young, C. B. Anfinsen, and A. Sandoval, JBC 239,3781 (1964).

672


I I S-Peptide I

Y-RNase-A S-Protein dimer

a

FIQ.7. Schematic diagram of RNase-S system. The single bond is cleaved converting RNase-A to RNase-S. RibonucleaseS dissociates reversibly to S-peptide f S-protein. The latter can recombine with denatured forms of RNase-A where the “tail” is loosened from the rest of the molecule. Reproduced from Richards ( 9 1 ~ ) .

extensive proteolysis at 37” removed residues through Phe 120 and resulted in total loss of potential activity. This derivative bound Speptide a t least a factor of 10 more weakly than S-protein itself. Although the enzymic activity of RNase-S is very similar to RNase-A, it is not identical. An extensive comparative study has been reported by Takahashi et al. (91). These authors varied the substrate, the temperature, and the pH. The pH optima are the same with RNA but different with C > p or CpA as substrates. The pH profiles vary with temperature. The effect of the lower thermal stability of RNase-S is evident above 30”. 2. Elastase (RNase-E)

Klee (74) has shown that porcine pancreatic elastase has an effect on RNase-A a t pH 8 similar to subtilisin. I n this case Ala 20 is excised by 91. T. Takahashi, M. Irie, and T. Ukita, J . Biochem. ( T o k y o ) 65, 55 (1969). 91a. F. M. Richards, in “Structure and Activity of Enzymes” (T. W. Goodwin, J. I. Harris, and B. S. Hartley, eds.), p. 5. Academic Press, New York, 1964.

24.


673

the cleavage of two bonds. The product, RNase-El is then a complex of residues 1-19 and 21-124. This complex shows properties similar to RNase-S. 3. Pepsin [ P I R or des- ( i H - l 6 4 ) - R N a s e ] Anfinsen (76)has shown that digestion of RNase by pepsin a t pH 1.8 and 37" produces initially a free tetrapeptide from the C-terminal end of the chain and a macromolecular component corresponding to the rest of the molecule, residues 1-120. This material shows no catalytic activity and is designated PIR for pepsin-inactivated ribonuclease. The work was confirmed by Fujioka and Scheraga (9.2) in digests prepared a t pH 2.0 and 25". The latter authors tentatively identified some of the bonds cleaved during later stages of the digestion. All of the digestion products isolated were missing the C-terminal tetrapeptide. Neither of these two studies found any evidence for pepsin digestion products which had any catalytic activity. The cause of the discrepancy between these reports and that of Ginsburg and Schachman (93)indicating catalytically active intermediates is not clear, but this latter study did not involve the actual isolation of components. Activity measurements on total digests can produce some very strange results [see, e.g., Allende and Richards (94)]. I n a recent study, Lin et al. (94a) has found that although the activity of PIR is low it is not zero. In carefully purified material assayed with C > p a t p H 6 the K,,, was about twice that of RNase-A while the turnover number was 0.5% of the native value. This material bound the inhibitors 2'-CMP about 12 times less strongly than RNase-A. When Phe 120 is removed with carboxypeptidase from PIR to give des- ( 1 2 ~ 2 4- ) RNase, all activity and binding properties are lost, 4. Trypsin Trypsin will attack native RNase-A a t neutral pH and room temperature very slowly or not at all. At higher temperatures where the thermal transition begins, cleavage by trypsin does occur. Oi et al. (76) have identified three macromolecular components in a digest a t 60" and pH 6.5, corresponding to the cleavage of bonds 31-32,33-34, or both of these. I n the latter case the dipeptide Ser-Arg is excised from the native molecule. All of these components show between 15 and 30% of the activity of RNase-A toward RNA and C>p. Limit digests a t very long times showed 92. H. Fujioka and H. A. Scheraga, Biochemistry 4, 2197 (1965). 93. A. Ginsburg and H. K. Schachman, JBC 235, 115 (1900). 94. J. E. Allende and F. M. Richards, Biochemistry 1, 295 (1962). 94a. M. C. Lin, B. Gutte, S. Moore, and R. B. Merrifield, JBC 245, 5169 (1970).

674

F.

M. RICHARDS AND H . W. WYCKOFF

the cleavage of 11-12 bonds near the maximum permitted by the specificity of trypsin. Ribonuclease-S differs from RNase-A in its great sensitivity to trypsin inactivation a t room temperature (94).The cleavage of S-peptide a t bond 10-11 causes loss of all ability to interact with S-protein and to regenerate catalytic activity. S-Protein plus trypsin leads to several intermediates with partial regeneratable activity and a limit digest with 7 bonds cleaved. This number is less than the maximum of 10 expected. With RNase-S, some protection of the S-peptide component was noted although cleavage eventually occurred. However, the maximum number of bonds cleaved was 6-7, less than the total of S-peptide and S-protein separately and much less than the expected maximum of 12. This would appear to indicate that the course of proteolysis is different in the complex and in the isolated components, and clearly different from the high temperature reaction on RNase-A. The absolute rates also indicate that the tryptic attack occurs on the complex, RNase-S, and does not require prior dissociation of the components. Some evidence was obtained for the loss of a dipeptide Ser-Arg in agreement with the RNase-A data. Tryptic attack a t room temperature on the components of RNase-A that are missing bonds 31-32 or 33-34 caused further cleavage, removing in the limit all residues between 31 and 38 to give a derivative with about 12% activity (77). Five other potentially susceptible bonds were not cleaved a t this low temperature, reminiscent of the RNase-S observations. 5. Chymotrypsin Chymotrypsin does not attack RNase-A a t neutral pH and room temperature. At elevated temperatures it will (95).The list of sensitive bonds proposed by Rupley and Scheraga (95)in the order of decreasing ease of cleavage was 25-26, 79-80, 97-98, 35-36, 76-77, and 4 6 4 7 . Demonstrably pure components with clearly defined primary structure were not obtained.

B. CHEMICAL MODIFICATION OF FUNCTIONAL GROUPS I n the following section attention is directed principally to those derivatives whose stoichiometry and properties have been well characterized. Discussion of the properties of the products is left largely to later sections. 95. I. A . Rupley and H. A. Scheraga, Biochemistry 2, 421 (1963).

24.

675


1. Carboxyl Groups

The modification of carboxyl groups has been carried out (1) by esterification with dry methanol and HCI, (2) by esterification with aliphatic diazo compounds, (3) by the formation of adducts with carbodiimides, or (4) by the formation of amides through activation with carbodiimides. Both complete and, apparently specific, partial modification of the 11 free carboxyl groups have been obtained. I n general, the first method suffers from the denaturing medium, the second from incomplete reaction, and the third from the uncertain nature of the products. The fourth procedure is perhaps subject to the least question. There are a total of 11 free carboxyl groups in native RNase-A; la(Val), !$(Asp), 5y (Glu) . A summary of the derivatives is given in Table V.

a. Methanol. Sela et al. [ (96) ; see also Vithayathil and Richards (97)] TABLE V SUMMARY OF CARBOXYL GROUPDERIVATIVES" ~~

Activity (%) of RNase-A No.

Derivative or abbreviation

1 (0Me)ll 2 (OMeh

(0-acetog1ycinamide)t 4 (WSC)I 5 (WSC)s 6 (WS'C)a 3

7 (WSC)e 8 9 10 11

(Gly-phth ester)ll (Gly)n (Ala-Gly-phth ester)?,* (Ala-Gly),,,

Modified carboxyl groups All All except 14, 38, 83 53 53 53, 49, 111 53, 49,

111, 9, 86 53, 49, 111, 9, 86, 38

All All NR NR

RNA

C>P > p)

(U

No. of buried TYr residues

0 1

0 NR

98 97 16

NR NR NR NR

3 3

2

NR

2

0 93 0 0

0 0 0 0

58

Ref.

2-3at 1" 0 at 25' 3

0 NR

The derivatives are indicated as methyl esters (OMe),, water-soluble carbodiimide adducts (WSC),, etc., where the subscript indicates the established stoichiometry, moles of modifying function per mole of protein. Phth stands for phthalimidomethyl and NR stands for not reported. 96. M. Sela, C. B. Anfinsen, and W. F. Harrington, BBA 26, 502 (1957). 97. J. J. Vithayathil and F. M. Richards, JBC 236, 1380 (1961).

676

F. M. RICHARDS AND H. W. WYCKOFF

obtained a fully esterified preparation by treatment of the protein in dry methanol, 0.1 M in HCl a t 25" for 24 hr. Broomfield et al. (98) obtained partial esterification in MeOH HC1 a t 2" for 1 week. Eight of the 11 groups were esterified. It was subsequently shown that the free carboxyl groups were Asp 14, Asp 38, and Asp 83 (99). The acid conformational transitions (see Section IV,B) all occurred a t lower temperatures a t any given pH value indicating a loosening of the structure. At 1" alkaline titration indicated 2-3 tyrosyl residues still buried, although all appeared to be available on titration a t room temperature. Ribonucleic acid assays (Kunitz) a t room temperature showed less than 1% residual activity. Assays a t 1" were not reported. Over 80% of the activity was recovered on saponification a t pH 10.44 and 25" for 26 hr. Fully esterified RNase-A when mixed with S-protein regenerates activity indicating that the "tail" in the (OMe)l, RNase-A derivative is no longer tightly bound to the rest of the molecule. This may result from either a general disruption of the structure or specifically from Asp 14 whose esterification no longer permits interactions with Tyr 25, His 48, and Arg 33. I n RNase-S this interaction is important but not crucial because of the flexibility produced by the missing peptide bond. I n RNase-A the loss of this interaction might force dissociation of the Nterminal residues although the explanation based on a general loosening of the structure seems more likely. It is also possible that carboxyl group 121 is involved in an important interaction with His 119, or that methylation blocks the active position of His 119. The tetramethyl ester of S-peptide (97) was prepared by treatment with dry MeOH and HC1. This peptide derivative is bound an order of magnitude more weakly than the unmodified peptide, but the interaction is still strong. The complex shows about 28% of the normal activity toward RNA and about 25% toward C>p. Full activity and binding constant are recovered on saponification.

+

b. Diazoacetoglycinamide. The reaction of diazoacetoglycinamide a t pH 4.5 and 10" never produced more than 25% esterification, but a derivative with a single ester group on Asp 53 was obtained (100). This carboxyl group appeared to be particularly reactive toward this reagent. The derivative showed full enzymic activity in the Kunitz RNA assay. c. Water-Soluble Carbodiimide Adducts. The water-soluble carbodiimide, 1-cyclohexyl-3- (2-morpholinoethyl) carbodiimide metho-p-toluenesulfonate is abbreviated WSC. Treatment of RNase-A with WSC a t 98. C. A. Broomfield, J. P. Riehm, and H. A. Scheraga, Biochemistry 4, 751 (1965). 99. J. P. Riehm, C. A. Broomfield, and H. A. Scheraga, Biochemistry 4, 760 (1965). 100. J. P. Riehm and H. A. Scheraga, Biochemktty 4, 772 (1965).

24.


677

pH 4.5 and room temperature leads to a number of chromatographically separable derivatives (101) where the detailed nature of the modification is unknown. With increasing extent of substitution the derivatives show decreasing catalytic activity and decreasing thermal stability. The normalization of one of the 3 buried tyrosyl residues, B, was associated with the modification of Asp 38. No evidence for intra- or intermolecular amide formation was found.

d. Phthalimidomethyl Esters. Glycine or alanylglycine as the phthalimidomethyl ester could be added by peptide bond formation to the carboxyl groups through activation by the water-soluble l-ethyl-3- (3dimethylaminopropyl) carbodiimide hydrochloride (10.2). The phthalimidomethyl protecting groups could be removed in 0.5 M piperidine in 30 hr a t 4" to yield free carboxyl groups on the newly added glycines. I n these latter derivatives the net charge on the protein was not changed, but the position of the COOH groups in the structure was altered. The retention of activity toward RNA and loss of C > p activity by the GlyIl derivative is particularly notable. 2. Amino Groups Ribonuclease-A contains 11 amino groups, la and 1 0 groups ~ on the lysine residues. Ribonuclease-S has one additional a-amino group a t position 21(22). A large variety of reagents will react with these functional groups. A summary of some of the reported derivatives is given in Table VI (103-119). See Section IV,B,9 for cross-linking to amino groups. 101. J. P. Riehm and H. A. Scheraga, Biochemistry 5,99 (1966). 102. M. A . Wilchek, A. Frensdorff, and M. Sela, Biochemistry 6, 247 (1967). 103. W. A. Klee and F. M. Richards, JBC 229, 489 (1957). 104. J. H. Reynolds, Biochemistry 7, 3131 (1968). 105. R. Goldberger and C. B. Anfinsen, Biochemistry 1, 401 (1962). 106. J. P. Riehm and H. A. Scheraga, Biochemistry 5,93 (1966). 107. C. H. W. Hirs, Brookhaven Symp. Biol. 15, 154 (1962). 108. C. H. W. Him, M. Halmann, and J. H. Kycia, A B B 111, 209 (1965). 109. R. P. Carty and C. H. W. Hirs, JBC 243, 5254 (1968). 110. R. P. Carty and C. H. W. Hirs, JBC 243, 5244 (1968). 111. C. B. Anfinsen, M. Sela, and J. P. Cooke, JBC 237, 1825 (1962). 112. D. Wellner, H. I. Silman, and M. Scla, JBC 238, 1324 (1963). 113. A. FrensdorE and M. Sela, European J. Biochem. 1, 267 (1967). 114. A. Frensdorff, M. Wilchek, and M. Sela, European J. Biochem. 1, 281 (1967). 115. R. R. Becker, Polyamino Acids, Polypeptides, Proteins, Proc. Intern. Symp., Madison, Wisc. 1961 p. 301. Univ. of Wisconsin Press, Madison, Wisconsin, 1962. 116. C. J. Epstein and R. F. Goldberger, JBC 239, 1087 (1964). 117. A . H. Nishikawa, R. Y. Morita, and R. R. Becker, Biochemistry 7, 1506 (1968). 118. R. L. Heinrikson, JBC 241, 1393 (1966). 119. J. Goldstein, J M B 25, 123 (1967).

678


TABLE VI SUMMARY OF AMINOGROUPDERIVATIVESO ~~

Activity

(%) of RNase-A No.

Derivative or abbreviation

1 2 3 4 5 6 7 8 9 10 11

(Guanidino)lo (Guanidino) 9.5 (Acetimido)11 (TFA)n (Dicyanoethy1)ll dl-DNP t, t'-7, 41-diDNP or-1-DNP a-l-SNP &I-SNP (polyMa)43

12 13 14 15 16 17 18 19 20 21 22 23 23a 24 25 26 27 28

Modified amino groups All E 9.5 E All All A11 Lys 41 Lys7, 41 Lys 1 Lys 1 Lys 41 Allexcept LYS 7 , 37, 41

7 7 7.5 NR 6.5 7 7.5 2 NR 8 7 7.6

7 Lys 41 Lys 41 Lys 41 Lys 1

RNA 0 33 0 0 0 b NR NR NR NR 100

50 150 18 4 8 0 28 60 100 P > p)

(U

NR NR 0 NR NR p as compared to the native enzyme, indicating that the synthetic material was not pure. Denkewalter and Hirschmann and their colleagues (191, 223-226), synthesized 75 pg of blocked Sprotein by solution methods. After deblocking of the SH groups, the reduced synthetic S-protein was combined with natural S-peptide and allowed to reoxidize. About 2 pg of RNase-S' activity was generated. Since synthetic S-peptide had been prepared in 1966 (227) the complete synthesis of RNase-S was achieved. Gutte and Merrifield (228) have recently reported the synthesis of S-protein and des-(21-25) S-protein by the solid phase method. When 223. R. G. Denkewalter, D. F. Veber, F. W. Holly, and R. Hirschmann, JACS 91, 502 (1969). 224. R. G. Strachen, W. J. Paleveda, R. F. Nutt, D. A. Vitali, D. F. Veber, M. J. Dickinson, V. Garsky, J. E. Deak, E. Walton, S. R. Jenkins, F. W. Holly, and R. Hirschmann, JACS 91, 503 (1969). 225. S. R. Jenkins, R. F. Nutt, R. S. Dewey, D. F. Veber, F. W. Holly, W. J. Paleveda, T. Lanza, R. G . Strachan, E. F. Schoenewaldt, H. Barkemeyer, M. J. Dickinson, J. Sondey, R. Hirschmann, and E. Walton, JACS 91, 505 (1969). 226. D. F. Veber, S. L. Varga, J. D. Milkowski, H. Joshua, J. B. Corn, R. Hirschmann, and R. G. Denkewalter, JACS 91, 506 (1969). 227. K. Hofman, M. J. Smithers, and F. M. Finn, JACS 88, 4107 (1966). 228. B. Gutte and R. B. Merrifleld, Federation Proc. 29, 727 (1970).

24.


699

mixed with a sample of S-peptide these synthetic protein components produced complexes with 5 and 9% respectively of the RNA activity (Kunitz) of RNase-A. The lack of effect of the removal of the 3 Ser residues 21, 22, and 23 is not surprising in view of the X-ray structure. However, both Asn 24 and Tyr 25 are clearly involved in interactions with other residues. These interactions must not be important for the formation or maintenance of an S-protein structure closely resembling that of the native enzyme. Apart from these complete syntheses, the major efforts in the past few years have been those of Hofmann and Scoffone and their respective colleagues. They have concentrated on synthesizing S-peptide and a series of fragments and derivatives related to S-peptide. I n the RNase-S system one can test the activity of an S-protein: S-peptide-derivative complex and separately obtain an estimate of the S-protein :S-peptidederivative binding constant either directly or from inhibition studies. Strongly bound derivatives will show maximum activity a t about 1 :1 molar ratio to S-protein under assay conditions. The more weakly bound the derivative, the higher the molar ratio required to approach full complex formation. For peptide derivatives whose complexes are inactive, the binding may be studied by inhibition of S-peptide or any of the active derivatives. Here the molar ratio of derivative peptide to S-peptide required to obtain 50% inhibition is a measure of the relative binding affinities. A summary of this synthetic work is given in Table XI (89, 94, 139, 153, 2 2 9 - 9 4 ) . 229. F. M. Finn and K. Hofmann, JACS 87, 645 (1965). 230. K. H. Hofmann, F. M. Finn, M. Limetti, J. Montibeller, and G . Zanetti, JACS 88, 3633 (1966). 231. K. Hofmann and H. Bohn, JACS 88, 5914 (1966). 232. F. M. Finn and K. Hofmann, JACS 89, 5298 (1967). 233. K. Hofmann, J. P. Visser, and F. M. Finn, JACS 92, 2900 (1970). 234. F. M. Richards and P. J. Vithayathil, Brookhaven Symp. Biol. 13, 115 (1960). 235. R. Rocchi, F. Marchiori, L. Moroder, A. Fontana, and E. Scoffone, Gazz. Chim. Ital. 96, 1537 (1966). 236. E. Scoffone, R. Rocchi, F. Marchiori, A. Marzotto, A. Scatturin, A. Tamburro, and G. Vidali, JCS,C p. 606 (1967). 237. L. Moroder, F. Marchiori, R. Rocchi, A. Fontana, and E. Scoffone, JACS 91, 3921 (1961). 238. R. Rocchi, L. Moroder, F. Marchiori, E. Ferrarese, and E. Scoffone, JACS 90, 5885 (1968). 239. F. Marchiori, R. Rocchi, L. Moroder, A. Fontana, and E. Scoffone, JACS 90, 5889 (1968). 240. R. Rocchi, F. Marchiori, A. Scatturin, L. Moroder, and E. Scoffone, Gazz. Chim. Ital. 98, 1270 (1968). 241. E. Scoffone, R. Rocchi, F. Marchiori, L. Moroder, A. Marzotto, and A. M. Tamburro, JACS 89, 5450 (1987).

TABLE XIA S Y N T I i E T l C P E P T I D E R E L A 4 T E D TO %PEPTIDE"

RNase-S' activity (%) a t peptide: Sprotein molar ratio

No. Ref. 1 2 3 4

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 H.LYS.GLU.THR.ALA-ALA.ALA.LYS.PHE.GLU.ARG.GLN.HIS.~ET.ASP.SER.SER.THR.SER.ALA.ALA.OH 100 200 300 700 ZOO0 5ooo 8OOO 8 4 3

10 7

7

10

12

15 13 11 18

6 7 8 9 10 11

15 24 65 100 80 65 100 10 16 21 26

39 48

61 71

70 74

12 13

52 100 0

0

BUT NH2 1

5

0

5

0

Activity (Yo) Mo- of RNase-A lar ratio RNA C > p 14 15 16 17 18 19 20 21 22 23

PY 1. NHZPY3.NH2-PY3PY3.0-C 02-

2 100 1 100 1 100 5500 0 1500 0

1

0 0 70

32

50

loo0

n

Derivative: Speptide ratio a t 50% inhibition

100 100 100

> 1OO0 1 600 8

h3

F

24 (233) 25 (233) 26 (233) 27

3CM3CM*+ 1CM1, 3 X dichf-

(233)

ORNORN-3CM-

28 29

1400

0.8

0

2400 100 25 50050 0

P Y 3-

3

No inhibition 20 6

0 0

RNase-S' activity (%) a t peptide: Sprotein molar ratio No. Ref.

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 H.LYS.GLU.THR.ALA.ALA.ALA.LYS.PHE.GLU.ARG.GLN.HIS.~ET.ASP.SER.SER.THR.SER.ALA.ALA.OH

1

2

5

10 30

100 600

3

z

M

9

3

0

E m Z C

0 30 31 32 33 34

(235) (236) (237) (237) (237)

35 36 37 38 39

(238) (238) (239) N P O --(239) R (240)

40 (2441) 41 (241) 42 (249) 43 (236) 44 (24.9) 45 (244) 46 (244)

V.1_.

-

ORN ORX

I-^-

SER-

ORN ORN

-- SER-

GLY-ORN AL A-ORN IL L O R N T Y R 4 R N ORN-GLU ORNLYS.GLU*THR.ALA.ALA-ALA ORN LYS.GLU.THR.ALA.ALA*ALA.LYkORN-

63 63 57 55 68 20 30

73 88 80 70 30

40 50 40 15 27

50

0

NLE

0 0 82 37 30 40

0 0

85 88 80 70

0

F

M

6

60 50

30

30

30

0

0

20 0 82

28

56

56

16

82

48 50 0 0

15 15

15 15

a The assays referred to in the right-hand columns employed RNA as a substrate unless otherwise indicated. These data cannot be used to provide accurate Sprotein: Speptide derivative binding constante (see reference 234).but they do give a qualitative picture of the effect on binding and activity of the indicated changea in covalent structure. The peptides are indicated by the horizontal lines with reference to the sequence at the top. Only altered residues are specifically indicated.

4

s

702

F. M. RICHARDS AND H.

W.

WYCKOFF

The data given on No. 14 in Table XIA clearly show that residues 15-20 of S-peptide play no significant role in either the binding of the peptide to S-protein or in the activity of the resulting complex. Removal of residues 1, 2, and 3 from the N-terminus is not serious but does show a slight fall in activity will all three removed (Nos. 32, 33, and 34). Methionine 13 and Asp 14 obviously play important parts in the binding. Addition of oxygen atoms to the sulfur, or conversion to a sulfonium salt by alkylation, invariably results in a dramatic decrease in binding constant but relatively little effect on the activity of the complex (Table XIB, Nos. 10, 11, and 12). The histidine residue in position 12 is crucial for activity. Its destruction by photooxidation or modification by iodination (No. 13, Table XIB) or carboxymethylation in the 3 position on the ring (No. 24, Table XIA) all destroy potential activity. The first two also substantially lower the association constant while the latter has no effect or may even increase it slightly. The CM group in the 3 position is, of course, easily accommodated and the interaction of the free carboxyl group with other positive charges nearby, e.g., His 119, may explain the increase in association constant. Conversion to a pyrazolyl residue destroys activity but not binding (No. 19). A most striking observation is the apparent activity of the 1-CMHis-12 derivative (No. 26). This is most unexpected. The N1 position of His 12 appears to be hydrogen bonded to the main chain in the X-ray structure. The residue is quite tightly packed in with only the N3 really exposed. It is not clear what movements could take place to accommodate a carboxymethyl group and presumably a counterion without seriously disrupting the geometry of the active site. The synthesis of this pcptide was actually carried out by chemical modification of a histidine-containing precursor peptide. Thus the synthetic route has all the difficulties normally associated with modification and separation of products. A contamination of the synthetic material with about 0.2% of a peptide containing His instead of l-CM-His would have been enough to explain the activity results. It is unfortunate that the authors did not also test the activity with varying S-protein and constant S-peptide concentrations. A comparison of the two curves could then be used to

242. F. Marchiori, R. Rocchi, L. Moroder, and E. Scoffone, Gazz. Chim. Ital. 96, 1549 (1966). 243. R. Rocchi, A. Scatturin, L. Moroder, F. Marchiori, A. M. Tamburro, and E. Scoffone, JACS 91, 492 (1969). 244. R. Rocchi, F. Marchiori, L. Moroder, G. Borin, and E. Scoffone, JACS 91, 3927 (1969).

W

0

4 z

m TABLE XIB DERIVATIVES OF NATURAL SPEPTIDEOJ RNsse-S’ activity (%) at peptide: Sprotein molar ratio 1

No. Ref. 1 2 3

(89) (f39)

4

5

(94) (234)

6

7

(2.994)

8

(234) (284)

9 1w

2

3

4

5

6

7

8

9

10

11

12

13

14

15

16

17

18

19

20

H.LYS.GLU.THR.ALA.ALB.ALA.LYS.PHE.GLU.ARGGLNHISMETASPSER.SER.THR.SER.ALA..4LbOH 1

5

10 30

100

600

-HSL

60

0 0

~

-G

G

100

(2S4) G

G

c,

A A

G A

P

100 100 40

__ ME

A -

M

. 02

M

ME 70

2 E

Ei

100

(94)

(2S4)

2

9

5? B 40

40

40 90

2r

40 100

40

CM 50 70 11 (234) CAM 50 70 12 (234) 13d (163) I 0 a Abbreviations: BUT. a-aminobutyric acid; NH2. amide; PY 1. &(pyraeolyl-l )Galanine; 0 2 . methionine sulfone; 0, methionine sulfoxide; CM. carboxymethyl; CAM, carboxamidomethyl; HSL, homoserine lactone; G , guanidino group; A, acetyl group; ME, methyl ester; I, idohistidine; and NLE. norleucine. b See footnote a to Table XIA. * Inhibition studies indicate a decreaee from the Speptide: Sprotein binding constant of a factor of 700. d Inhibition studies indicate a decrease from the Speptide: S-protein binding constant of a factor of 3200.

704

F. M . RICHARDS AND H . W. WYCKOFF

test for just this possibility as had been done previously by Vithayathil and Richards (138) in a similar situation involving Met 13. The glutamine in position 11 can be converted to glutamic acid without a major effect on the activity (Nos. 11 and 43). An earlier statement to the contrary (97)was probably wrong. I n the case of No. 11 one must consider possible impurities such as residual blocking agent at a 0.1% level. In the case of No. 43 it must be noted that residue 10 is ornithine and this flexible charge could be in a salt link to Gln 11. Introduction of a negative charge adjacent to the pl site would be expected to have a significant effect (see Section VI). Arginine 10 contributes to the binding since replacement with ornithine lowers the association constant by a factor of 20. Phenylalanine 8 like Met 13 is important to the binding. A drastic reduction in the association constant is shown by all derivatives tested with substitution in position 8 (Nos. 39, 40, 41, and 46) except tyrosine (No. 42). In the latter case neither binding nor activity is significantly affected. Apparently tyrosine occupies the position of Phe 8. I n the XTABLE XI1 SUMMARY Property Partial specific volume Refractive index increment Radius of gyration Diffusion coefficient translation rotation Sedimentation coefficient Molecular weight Frictional ratio Intrinsic viscosity Isoionic point Hydration Axial ratio Molar absorbancy a t 278 nm E:.fz See text for references.

OF PHYSIC AL PROPERTIES"

Conditions Neutral pH, 25" or less pH 7.7, 25", X = 546nm

Infinite dilution Neutral pH, 25" 0 SZ0,W

Anhydrous

f/fo

pH 3-9, -25" or less, Z = 0.1

As prolate ellipsoid Neutral pH, 25" or less Neutral pH, 25" or less

Average or "best" value 0.703 ml/g 0.185 ml/g 18.3 A 1.26 X 10-6 cm*/sec 1.07 X 10-6 cmz/sec 230 X 106 sec-I 1.78 S 13,680 1.26 3 . 3 ml/g 9.60 0 . 3 - 0 . 4 g/g -2 9700 cmz/mole 0.71

24.


705

ray structure there is room to accommodate the extra oxygen atom with very slight shifts in the hydrophobic core, However, there is not enough space for water molecules as well, and there is nothing in the core area for the phenolic OH to hydrogen bond to. One would expect that a marked decrease in association constant would accompany this substitution, but that does not appear to happen. Spectral data show that this tyrosine is clearly buried and titrates abnormally. Alteration of the net charge on the peptide has little effect on the binding constant and changes the activity only within a factor of two or so (Table XIB, Nos. 8 and 9 ) . The nonpolar and H bonding interactions obviously make the most important contributions in the Speptide :S-protein interaction. Kato and Anfinsen (245) have reported on the use of agarose-bound S-protein in the form of a column for the purification of synthetic Speptide derivatives. This procedure has shown the presence of closely related products in material from the Merrifield solid phase techniques. Some of these products produced enaymically inactive complexes with S-protein.

V. Molecular Properties

A. PHYSICAL PARAMETERS A summary of some physical properties is given in Table XII. 1. Partial Specific Volume Many measurements of this parameter have been made over a long period of time and by a variety of different techniques. The early pycnometric measurements of Rothen (246) gave a value of 0.709 cm3/g. Later measurements in a gradient column by Harrington and Schellman (247) gave 0.694 cm3/g. The very precise magnetic float procedure developed by Beams and his colleagues gave a series of different values shown in Table XI11 (248). I n general all careful measurements reported by others for p H values near neutrality and temperatures at or below 30" 245. I. Kato and C. B. Anfinsen, JBC 244, 5849 (1969). 246. A. Rothen, J . Gen. Physiol. 24, 203 (1940). 247. W. F. Harrington and J. A. Schellman, Compt. Rend. Trav. Lab. Carlsberg, Ser. Chim. 30, 21 (1956). 248. D. V. Ulrich, D. W. Kupke, and J. W. Beams, Proc. Natl. Acad. Sci. U. S. 52, 349 (1964).

F.

706

M. RICHARDS AND H.

W.

WYCKOFF

TABLE XI11

THEPARTIAL SPECIFICVOLUMEOF RIBONUCLEASE-A SAMPLES~ ~

~

Sampleb

t(ml/g)

0.15 M KC1-0.007 M Phosphate (K), pH 7.6, 20"

Sigma, Type 111, lot R-22B-70 ( N

=

Sigma, Type 111-A, lot 43B-772-3 ( N

17.0 f 0.2%)

=

16.9 f 0.2%)

Worthington (P04free),lot RAF 6060 ( N

=

16.4 f 0.2yQ)

Worthington (PO4 free), lot RAF 6065 ( N Worthington (PO4 free), lot RAF 6067 ( N

=

1 6 . 3 k 0.2%) 16.5 f 0.2%)

=

0.7060 0.7046 0.7040 0.7036 0.6979 0.6955 0.6988 0.7004 0.7030 0.7005 0.7134

f 0.0006 f 0.0016 f 0.0019 f 0.0008

f 0.0011 f 0.0012 f 0.0006

k 0.0010 f 0.0010 f 0.0013

k 0.0006

0.15 M KC1-0.010 M Glycine (K), p H 9.6

Sigma, Type 111-A, lot 43B-772-3

0.6919 f 0.0017 0.6923 f 0.0020 0.15 M KC1, p H 7.8

Sigma, Type 111-A, lot 43B-772-3

0.6949 f 0.0015

Table taken directly from Ulrich et al. (248). The range of protein concentration over which densities were determined for computing each value of t was 1.52 f 0.18% to 0.07 f 0.02%. a

b

also fall in this range. Particularly in the work of Beams, the observed variation between the different samples is well outside the experimental error of the technique. The principal error lies in the measurement of concentration where the definition of dry weight is elusive as always. The importance of the Donnan correction a t p H values removed from the isoelectric point has been pointed out by Krausz and Kauzmann (249). Even here the variation cannot explain the range of observed differences. The cause of the variation between samples remains to be found. For many purposes the mean value of about 0.703 cm3/g can be used. This agrees nicely with the value calculated from the amino acid composition (250). For careful work, however, even with purified RNase-A, the value must be determined for each lot under the precise conditions of interest: By dilatometry, Holcomb and Van Holde (251) measured the values 249. L. M. Krausz and W. Kauzmann, Proc. Natl. Acad. Sci. U.S. 53, 1234 (1965). 250. T. L. McMeekin and K Marshall, Science 116, 142 (1952). 251. D. N. Holcomb and K. E. Van Holde, J . Phys. Chem. 66, 1999 (1962).

24.

707


of b from -15" to 70" through the range of the thermal transition as seen in Fig. 8. The solvent was 9 mM phosphate buffer, 0.15 M KCl, pH 7.7. Note that although b increases with temperature, the volume change for the transition (see Section V,B,6) native to denatured is negative. The value a t the midpoint of the transition is about 240 ml/mole (see, however, Section V,B,6). The temperature dependence of B for ml/g/deg (963) and ml/g/deg or 8 X native RNase is 4.5 X ml/g/deg. There is no change in fi for the denatured form 10 X with pressure between 0 and 400 atm (959). 2. Refractive Index Increment

The best value appears to be that of Beams et al. (253), dn/dc = 0.185 ml/g a t 25.0O0C,pH 7.7, and X = 546 nm.

3. Radius of Gyration Analysis of low angle X-ray scattering data gives a value for the radius of gyration at infinite dilution of 18.3 A (954). The scattering curve can best be explained on the basis of a cylinder or prolate ellipsoid I

I

I

0.72

0.71

-3 E

IA

0.70

0.69 T

(OC)

FIG.8. The dependence of partial specific volume on temperature. Reproduced from Holcomb and Van Holde (261). 252. P. F. Fahey, D. W. Kupke, and J. W. Beams, Proc. Natl. Acad. Sci. U.8. 63, 548 (1969). 253. A. M. Clarke, D. W. Kupke, and J. W. Beams, J . Phys. Chem. 67, 929 (1963). 254. I. D. L. Filmer and P. Kaesberg, Brookhaven Sump. Biol. 15, 210 (1962).

708


with axial ratios between 2 and 3. The molecular volume of the hydrodynamic ellipsoid is of the order of 30,000 AS,much larger than the actual molecular volume of about 15,800A3. The difference is attributed to hydration of the order of 0.6 gram of water per gram of anhydrous protein.

4. Diffusion Coefficients A careful investigation by Creeth (255) using the Gouy interference of system gave a value for the translational diffusion coefficient, D,,,,, cm2/sec a t 25.0"C where the solvent cm2/sec or 12.22X 10.68 X conditions for measurement were KC1 0.1 M , potassium phosphate buffer 0.035 M , pH 7.74,25.0",with an average protein concentration of 0.38 g/100 ml. Van Holde and Baldwin (266) reported the values in the accompanying tabulation. Protein conc. ( d l 0 0 ml)

(cmz/sec)

1.08 0.54 0.27

1.17 X 1.22 x 10-6 1.24 x lo-'

Dabvw

A recent study has employed the new technique of quasi-elastic light scattering for estimation of diffusion constants (257). The result for the native enzyme a t 24" is in close agreement with the data given in the tabulation. The value of D as a function of temperature was measured through the thermal transition. The method was also used to follow the kinetics of the urea-induced transition. The rotational diffusion constant in water at 25" and neutral pH as measured by electric birefringence (958) is 230 X lo5 sec-l or 0.73x 1 P sec as a relaxation time. For a hydrodynamic ellipsoid of dimensions 66 X 22 A and a molecular weight of 14,000,the calculated relaxation sec. However, the apparent asymmetry of the molet i d e is 0.72x cule from the X-ray structure corresponds to an axial ratio of no more than 2:l rather than 3:l. 255. J. M. Creeth, J. Phys. Chem. 62, 66 (1958). 256. K. E. Van Holde and R. L. Baldwin, J. Phys. Chem. 62, 734 (1958). 257. L. Rimai, J. T. Hickmott, Jr., T. Cole, and E. B. Carew, Biophys. J. 10, 20 (1970). 258. S. Krause and C. T. O'Konski, Biopolymers 1, 503 (1963).

24.


709

5. Sedimentation Behavior Ribonuclease-A carefully purified to remove aggregates sediments as a single symmetrical peak. I n most preparations of the enzyme there will be a fast moving shoulder on the main peak or at least a slightly unsymmetrical peak. The sedimentation coefficient of RNase-A at neutral pH, s&,, is given as 1.78 2 0.01 S by Yphantis (259). Many reported values for the unfractionated enzyme or for samples of uncertain purity lie in the range of 1.8-2.0 S. The fractionated material shows almost ideal behavior in equilibrium ultracentrifugation while the cruder samples give higher average molecular weights at the cell bottom than a t the meniscus. 6. Molecular Weight

The anhydrous molecular weight calculated from the amino acid composition is 13,680. Osmotic pressure measurements by Kupke (260) in 6 M guanidine hydrochloride gave a best estimate of 14,000. All estimates of M from sedimentation require a knowledge of V . Measurements have not always been made of both parameters on the same sample. The accuracy of literature estimates cannot always be evaluated. Most estimates lie in the range of 13,000-14,000. Very careful measurements in a Beams magnetically suspended centrifuge (261) gave an equilibrium value of 13,650 which is embarrassingly close to the value calculated from amino acid analysis. Yphantis (259) and Van Holde and Baldwin (256) found essentially the same value from the average of a series of short column equilibrium runs. Erlander and Foster (262) obtained about 13,800 from Archibald approach-to-equilibrium determinations. There can be little doubt at t,his point t'hat deviations from the theoretical value either result from errors in the procedure used or from a small amount of aggregated material in the sample used. 7. Hydration and Axial Ratio

This property remains difficult to define and to estimate. The value of 0.6 g/g quoted above from low angle X-ray scattering appears high. In a careful study of sedimentation behavior as a function of salt concentration, Cox and Shumaker (263) concluded that the preferential 259. D.A. Yphantis, J. Phys. Chem. 63, 1742 (1959). 260. D.W.Kupke, Compt. Rend. Trav. Lab. Carlsberg 32, 107 (1961). 261. R.D.Boyle and P. E. Hexner, Science 134, 339 (1961). 262. S. R.Erlander and J. F. Foster, J. Polymer Sci. 37, 103 (1959). 263. D.J. Cox and V. N. Shumaker, JACS 83,2433 (1961).

710

F. M. RICHARDS AND €I W. . WYCKOFF

hydration for ribonuclease was 0.34 g/g. This value is close to the average estimate by a variety of techniques for a number of proteins. To the extent that the X-ray structure can be represented by an ellipsoid of revolution, it appears to be an oblate ellipsoid with an axial ratio no greater than 1:2. The approximate dimensions are 25 x 45 x 45 A. From molecular weight and free diffusion data the frictional ratio f/fo is 1.26. This corresponds to an unhydrated prolate ellipsoid with an axial ratio of 5.2 or an oblate ellipsoid with a ratio 0.18. If the hydration is assumed to be 0.34 g H,O/g protein, the axial ratio values would be 3.0 and 0.33, respectively. The maximum hydration for a sphere would be 0.7 or 0.55 g/g for a prolate ellipsoid of axial ratio 2. In their study of the polarization of fluorescence of labeled RNase, Young and Potts (264) employed the Scheraga-Mandelkern treatment of the hydrodynamic data and obtained a value of the coefficient @ = 2.12 x lo6, which is compatible with an axial ratio of 1 : 2 for an oblate ellipsoid. 8. Viscosity The intrinsic viscosity of native ribonuclease is very low. Harrington and Schellman (247) reported 3.3 ml/g a t neutral p H in 0.1 M KC1. Buzzell and Tanford (265) found values of 3.3-3.5 ml/g over the entire pH range from 1 to 11 and ionic strengths from 0.05 to 0.25 M . This value increases dramatically on denaturation even without oxidation or reduction of the disulfide bonds to 8.5 ml/g ( 2 6 6 ) . I n the presence of reducing agents and 6 M guanidine hydrochloride the value is 16.0 ml/g ( 2 6 7 ) . 9. Electrophoretic Mobility The isoionic point of RNase-A determined as the pH of a concentrated salt-free solution is 9.60 (268). Estimates of the isoelectric point in buffers not containing phosphate all give values above 9, Anderson and Alberty (269) reporting 9.45. I n phosphate buffers specific anion binding dramatically reduces the measured isoelectric point to an extent dependent on the total phosphate concentration. Values below 6 have been measured (270). 264. 265. 266. 267. 268. 269. 270.

D. M. Young and J. T. Potts, Jr., JBC 238, 1995 (1963). J. G. Buzzell and C. Tanford, J . Phys. Chem. 60, 1201 (1956). M. Sela, C. B. Anfinsen, and W. F. Harrington, BBA 26, 502 (1957). C. Tanford, K. Kawahara, and S. Lapanje, JBC 241, 1921 (1966). C. Tanford and J. D. Hauenstein, JACS 78, 5287 (1956). E. A. Anderson and R. A. Alberty, J . Phys. Colloid Chem. 52, 1345 (1948). A. M. Crestfield and F. W. Allen, JBC 211, 363 (1954).

24.


711

Electrophoretic heterogeneity was indicated in early reversible boundary spreading experiments. Although careful examination of free boundary electrophoretic runs can show several components in normal preparations (271), the heterogeneity is most easily seen in studies on supporting media. Both paper (870) and starch (272,273) give multiple zones. The minor zones can be correlated a t least in part with the fastmoving chromatographic peaks. Purified RNase-A moves as a single zone. 10. Hydrogen Ion Equilibria The titration data of Tanford and Hauenstein (268) indicate that all ionizable groups are accessible in the native enzyme except for 3 tyrosine residues (274) (see discussion below). The guanidyl groups had a pK, value greater than 12, the three accessible phenolic groups a value of 9.95, the p) in the presence of 2 mM SDS (308). The midpoint of the transition at pH 5.8 in 0.02 M phosphate buffer, 27”, as detected by fluorescence occurs at about 1 mM SDS. There is no additional acid transition detectable by fluorescence in spite of the fact that the SDS transition is supposed to “normalize” one residue while the acid transition affects two. Jirgensons (365) has made the intriguing suggestion based on ORD studies of a number of proteins that detergents induce the formation of helical structure from either unordered or /3 structures. Bigelow (334) [with the correction supplied by Sarfare and Bigelow (SSS)] has summarized a great deal of the work by himself and others. The experimental data considered were obtained by difference spectra, ORD, and viscosity. They may be summarized as follows: Native RNase 365. B. Jirgensons, JBC 241, 4855 (1966). 366. P. S. Sarfare and C. C. Bigelow, Can. J. Biochem. 45, 651 (1967).

24.

737


(N) undergoes denaturation by a path which has two intermediate states (I and 11), before reaching the completely random coil (111).All of the N e I e I1

~

I11

steps are reversible. This is the minimum number of states required to explain presently available data. The properties of these different states are shown in Table X I . The assumption is an all-or-none “normalization” of each of the 3 residues. It is interesting how well this appears to explain the data although there is no a priori reason why that should be so. The residues initially labeled A, B, and C have been tentatively identified as T y r 25, 92, and 97, respectively (300). The normalization of Tyr 92 (B) causes a decrease in molar absorbance a t 287 nm of 700 with little change in either viscosity or rotation. Of the three this is the most accessible residue in tho X-ray structure. The normalization of the other two causes changes of 1000 each in absorbance and is accompanied by both viscosity and rotation changes. The second to normalize is Tyr 25 (A) presumably by dissociation of the N-terminal portion of the chain from the body of the molecule, which would expose this residue as in S-protein. The last is Tyr 97 (C) whose exposure requires disruption of the entire structure. This residue is also the most buried of all the tyrosine residues according to the X-ray structure. However, the accessibility of Tyr 97 to chemical modification in S-protein and Met 30 t o alkylation in RNase-A indicate that the region around Tyr 97 may be easily deformable. If this is so, why should Tyr 97 be the last to normalize. TABLE X X I

DENATURED CONFORMATIONS

OF

RIRONUCLEASE AND THEIR PROPERTIES“

h1 State Native I I1

I11

Denaturant

pH 1 at 15°C pH 6 at 60°C pH 1 at 40°C 5 M LiBr pH 2, p = 0.1, 2 M dioxane 8 M urea 6 M guanidinium chloride

As281 nmb

0 -700 (B) - 1700 (B and A) -1700 (B and A) -1700 (B and A) -1700 (B and A)

-2700 (B and A) -2700 (B, A, and C)

(dl/g)

0

0.033 0.035 0.065 0.072 0.073 0.070

21-23 24 23 23

0.093-0.095 0.094

46 51

7

Taken from Bigelow (334). and C identify the tyrosyl residues which contribute to the observed changes in the various parameters. B, Tyr 92; A, Tyr 25; and C, Tyr 97. a

* Here B, A,

738


W.

WYCKOFF

5. Transitions in Derivatives of RNase-A

a. RNase-S. In general, RNase-S is less stable t o perturbing conditions than RNase-A, and S-protein is less stable than RNase-S. Sherwood and Potts (367) measured the transition temperatures of these three proteins as a function of urea concentration using the rotation a t 366 nm as an index. The results are shown in Fig. 17. For the thermal transition in 0.9% NaCl, 0.01M phosphate buffer, pH 6.8, the molar absorbance change was the same for both RNase-A and RNase-S and was equal to 1750, equivalent to the normalization of 2 tyrosine residues (304). The change for S-protein was about 850. At the higher temperatures all of these samples still have 1 tyrosine buried. Richards and Logue (368) examined the acid transition of RXase-S in 0.1 M NaC1-0.01 M acetate-tris at 22". The midpoint for RNase-S was about pH 3.3 and for S-protein about 4.4.The complex with the tetramethyl ester of S-peptide was intermediate between these values, all being at higher pH than RNase-A. The difference spectra obtained on association of S-peptide and S-protein clearly showed the perturbation of phenylalanine as well as tyrosine residues. Phenylalanine 8 is certainly involved in the spectral shift and perhaps also Phe 120. A peculiar facet of the association was the time dependence of the spectral changes. There was a very rapid change probably reflecting the

0

1

I

I

I

2

3

4

,

5

I

6

J

I

7

8

Urea molarity FIG.17. Comparison of the transition temperatures for RNase-A (circles), RNase-S (triangles), and S-protein (squares) as determined by optical rotation (open symbols) and ultraviolet absorption difference spectroscopy (filled symbols). Reproduced from Sherwood and Potts (367). 367. L. M. Sherwood and J. T. Potts, Jr., JBC 240, 3799 (1965). 368. F. M. Richards and A. D. Logue, JBC 237,3693 (1962).

24.


739

bimolecular association. This was followed by a relatively slow first-order change, in the same direction, with a half-time of 60-90 sec. These measurements were made in 0.1 M acetate buffer pH 4.5. On neutralization from pH 1.7 to 4.6, S-protein showed a similar first-order change, this time with no initial fast portion. The origin of the slow isomerization is not clear. Using circular dichroism, Simons and Blout (369, 369a) found that the acid titration of the 240-nm peak showed the same pH dependence as observed by AcZs7.The C D spectrum of S-protein changes continuously from 10" to 40". Lowering the temperature from 40" does not result in complete reversal. The product, however, is fully activatable on addition of S-peptide. b. C M Derivatives. Yang and Hummel (370) found l-CM-His-119RNase and 3-CM-His-12-RNase (Table VIII, Nos. 5 and 6) to be denatured less rapidly in 8 M urea pH 5 than RNase-A. I n acid solution both derivatives were slightly more stable than RNase-A. I n basic solution the His 12 derivative was markedly less stable than the other two. Although the spectral titration curve was still biphasic, the time-dependent changes in absorption began about 1 pH unit lower than for RNase-A or the His 119 derivative. The free carboxyl group and its interaction with the strongly positive center might explain the increased stability a t pH 5, and, if the primary interaction is with His 119, this would explain why such an effect disappears a t pH 8 and above for the His 12 derivative. The pyrophosphate ion a t 0.5 mM, which binds between His 12 and 119, strongly protects RNase-A against denaturation, but it has no effect on the two CM derivatives nor is there any evidence for the binding of the nucleotide 2'-CMP. From the X-ray structure the CM group would be expected to block sterically the phosphate binding site. c. DNP-Lys-41-RNase. Ettinger and Hirs (371) have shown by solvent perturbation methods that the D N P group is partly buried. This agrees with the X-ray data on this derivative (120) where the D N P group tucks up near Arg 39 and Asp 38. The thermal transition for the protein part of the derivative estimated by AcZs7 in 0.009M tris pH 7.2 had a T, of 61.9" (compared to 62.1" for RNase-A). However, the transition a t 358 nm specifically reflecting the D N P group occurred a t 55.5". This is further evidence of the tendency of the structure around Asp 38 to loosen more easily than the bulk of the molecule. 369. E. R. Simons and E. R. Blout, JBC 243, 218 (1968). 369a. E. R. Simons, E. G. Schneider, and E. R. Blout, JBC 244, 4023 (1969). 370. S.-T. Yang and J. P. Hummel, JBC 239, 3775 (1964). 371. M. J. Ettinger and C. H. W. Hirs, Biochemistry 7, 3374 (1968).

740

F. M. RICHARDS A N D N. W. WYCKOFF

d. (PoZyVaZ) RNase. The observations on this material by Becker and his colleagues (117,372) were of a different sort. The process followed was aggregation, presumably directly involving the valine side chains added to the native enzyme. The lowering of the rate of aggregation on increasing the pressure corresponded to an activation AV of between 200 and 250 ml/mole. The aggregation rate increases a t higher ionic strength and is decreased by urea and ethylene glycol. These effects are in the same direction as those seen for detergent micelle formation and to those expected for hydrophobic bond formation.

6. Thermodynamics of the Conformational Transitions The most extensive and detailed studies of the equilibrium processes are those of Brandts and his colleagues (336, 339, 373). A careful calorimetric study has been reported by Tsong et al. (374).This work and that of others is discussed by Tanford in a recent review (333).Although the primary data are of very high quality, considerable uncertainty is involved in their interpretation. The problems are perfectly general in protein chemistry. ( 1 ) I n order to convert measured variables to fractional conversion in a transition, careful attention must be given to estimation of the values of these variables for the limiting conformations in the transition region. Absorbance, rotation, or other properties of a given conformer may be marked functions of temperature, pH, or urea concentration. Usually this dependence can be adequately represented by a linear relationship. On the other hand, it is impossible to state with certainty that the variation of some property outside of an obvious transition region is not in fact resulting from a variable distribution of the protein among several or many conformers. (2) The transition must be shown to be reversible. If the protein concentration is low (less than 0.5 mg/ml), this can be shown to be true for RNase over a wide range of temperature but only a t p H values below about 3. The transitions are frequently so sharp that accurate estimates of equilibrium constants can be made only over a narrow range of the variable. Thus comparisons of parameters usually involve alteration of more than one variable a t a time (i.e., pH and temperature) in order to keep the measurements in an accessible range. Paying as much attention 372. M. S. Kettman, A. H. Nishikawa, R. Y . Morita, and R. R. Becker, BBRC 22, 262 (1966).

373. J . F. Brandts, R. J. Oliveira, and C. Westort, Biochemistry 9, 1038 (1970). 374. T. Y. Tsong, R. H. Hearn, D. P. Wrathall, and J . M. Sturtevant, Biochemistry 9, 2666 (1970).

24.


741

to these problems as is presently possible, the available data on RNase still cannot be fully reconciled. The basic question is how many stable states are involved in the transition. When only two are present, detailed thermodynamic calculations are possible. With more than two, calculations become increasingly uncertain. Evidence for a two-state process can be indicative but never compelling. Evidence against a two-state process may be compelling in some cases, but it is not necessarily always so. The significance of a multistate process will depend on the concentration of the intermediates and the values of the particular physical variable of interest applicable to each state. The summaries in the previous sections have indicated some of the difficulties. At present, meaningful thermodynamic parameters can be obtained only for a process which is at least approximately two state in character. Brandts (375)made the case for this, and the two-state process is assumed in the data quoted in Table XXII (336, 337, 339, 348, 355, 362, 374, 376, 377) and Fig. 18. The marked increase in heat capacity between the native and the denatured forms has been observed in the denaturation of several proteins (333). This appears to be related to the solvent structure around newly exposed side chains in the unfolded conformation. The heat capacity difference is responsible for the temperature dependence of AH and for the curvature almost invariably seen in van't Hoff plots of equilibrium data. The measured free energy changes must always be small in the range of actual measurement since 0.1 < K < 10 for accurate estimates. It can be seen (Table XXII, No. 1) that the free energy is the small difference between large enthalpy and entropy terms. Small changes in either of the latter will markedly affect the former. The numbers a t 0" and 60" were derived from the extrapolated values of the power series curve fitted to the ,AGOvalues in the measurable range. The dramatic change in AH" is undoubtedly real. The AC, values are obtained by double differentiation; their reliability is uncertain, The variety of values obtained in earlier studies reflect some of the difficulty. The value a t pH 2.1 and 30" from Ginsburg and Carroll (Table XXII, No. 5) does not agree well with that given above, but the effect of phosphate and sulfate in stabilizing the structure, even a t acid pH, is clearly shown. Direct calorimetric results (Nos. 6 and 7) do not agree with each other and bracket the data of Brandts and Hunt. Aggregation may be a problem in studies with more concentrated solutions. The more 375. J. F. Brandts, "Biological Macromolecules," (S. N. Timasheff and G. Fosman, eds.), Vol. 2, p. 213. Marcel Dekker, New York, 1969. 376. G. C. Kresheck and H. A. Scheraga, JACS 88, 4588 (1966). 377. A. K. Beck, S. J. Gill, and M. Downing, JACS 87, 901 (1965).

TABLE XXII

THERMODYNAMIC PARAMETERS OF THE ACID TRANSITION (TWO-STATE PROCESS ASSUMED) (Units for AGO, AH", TAS" = kcal/mole, for AC, = kcal/mole/degree) No.

pH

AGO

AH"

TAS"

ACp

AHo

30"~

AC,

AH"

0"b

AC,

60"

~~

~

1

1.13 2.10 2.50

2.77 3.15

Water, no added electrolyte, pH adjusted with HCl 60.3 -1.1 61.4 2.1 11 1.2 137

-0.5 62.3 0.9 57.2 2.0 56.6 3.1 53.0

62.8 56.3 54.6 49.9

2.0 2.0 2.0 2.0

15 10 8 6

1.2 1.2 1.2 1.2

135 131 132 126

3.0 2.9 2.9 2.9 2.9

Calorimetric and van't Hoff values No.

pH

T or T,

Solvent

AHcal

RNase-A 2 4.8 O.1MKC1 62 3 6.5-6.8 0.01M phosphate-0.12 M 61.5

AH&

AC,

76.5 121-135

-

Ref.

KCl

4

1-7 1-7

5

2.1

0.01M buffer O.16MKC1 0.01M buffer 0.16M KC1 Ionic strength 0.019M

6 7

2.2 2.8

8

0.36 1.05 2.02 2.80 3.28 4.04 5.00 6.23 7.00 7.80

chloride Ionic strength 0.019M phosphate Ionic strength 0.019M sulfate 0.1M KCl 0.15M KC1, 0.01M phthalate HCl HCl Glycine 0.2M Glycine 0.2M Glycine 0.2M Acetate 0.2M Acetate 0.2M Acetate 0.2M NaCl 0.2M NaC10.2 M

7.0

NaC10.2 M

T, 30

51 110 48

-

32

57

-

44

72

-

45

-

-

-

109 f 5 70 f 1

31.5 29.9 31.2 40.6 45.8 52.3 57.8 60.8 61.3 61.2

61 59 66 88 105 126 151 155 168 178

62 59 67 68 67 64 60 66 97 72

1.9 2.0 2.0 2.1 2.0 2.1 2.2 2.0 2.1 2.0

107

62

2.1

111

63

2.1

55

30

1.4

0.66

RNase-S

47.7 RNase-S'

7.0

NaCl 0.2M

47.1 SProtein

7.0 0

NaC10.2 M

Brandts and Hunt (336).

37.6 Brandts (339).

.

. I

24.

743


-10

10

0

20

30

40

50

Temperature

- 400

(b) 100

-

-

-

AH-

-

AC,/lO

As 0

- 20

0 0

20

40

60

Temperature

FIQ. 18. (a) The temperature dependence of the free energy change for the RNase transition at different pH values. The points represent AF" values calculated from the data. The solid curves are the best fit to a quadratic equation by least squares analysis. The dashed lines indicate the range of relatively high experimental accuracy. (b) The values of AH", AS", and ACp for the RNase transition from data at pH 2.50. Reproduced from Brandts and Hunt (886).

744


recent data of Tsong et al. (374) indicate that analysis by a two-state process is not valid above a pH of about 2. In the presence of other solvent components such as urea and ethanol the parameters for the thermal transition change in a complicated way. At concentrations of 10-15% and a t pH 3.2, ethanol is a stabilizer a t low temperature and a denaturant a t temperatures of 30" and above. Urea a t pH 3.3 is a denaturant a t all concentrations with the effect on being roughly linear in urea molarity. An excellent discussion of these problems is given by Brandts (336). Effect of Pressure. The volume change occurring during the transition can be estimated directly by dilatometry or indirectly by studying the pressure dependence of the equilibrium constant. On the basis of the temperature dependence of partial specific volume (pH 2.8, 0.15 M KCl), Holcomb and Van Holde (661) estimated the change to be -240 =k 100 ml/mole. Gill and Glogovsky (378) estimated the volume change in the same solvent from d In K / d P and arrived a t a value of -30 =k 10 ml/ mole. Brandts et al. (S'Z?), in a very extensive study, found numbers similar to that of Gill and Glogovsky. The investigation covered the temperature range 0"-65" and the pressure range 0-50,000 psi. All changes in were demonstrably reversible a t the very low protein concentration used (< 0.5%). The volume changes were pressure dependent becoming more negative as the pressure increased. At pH 2 and 25" the volume change was about -45 ml/mole, but this dropped t o -5 ml/ mole a t pH 4 and 50". The compressibility of the denatured protein appeared to be larger than the native by about 1.5 x atm-'. These figures are hard to reconcile with the volume change indicated by the pycnometric values. The values of AV are all small and are close to the error of many of the experiments. Most of the expected effects from current theories of protein structures would predict much larger negative values of AV. As pointed out by Brandts et al. (373) most of the large volume changes expected on denaturation are negative. On the basis of studies on model compounds, increased electrostriction and insertion on nonpolar residues into an aqueous environment would be expected to produce volume contractions of the order of several hundred milliliters per mole for RNase. Apparently, this does not happen. A G O

C. AGGREGATION Crestfield et al. (379) have shown that most RNase samples contain dimers and higher aggregates. The amount of this aggregated material 378. S. J. Gill and R. L. Glogovsky, J . Phys. Chem. 69, 1515 (1965). 379. A. M. Crestfield, W. H. Stein, and S. Moore, ABB Suppl. 1, 217 (1962).

24.

745

BOVINE PANCREATIC RIBONCCLEASE

can be markedly increased by lyophilization of the enzyme from 50% aqueous acetic acid. The aggregates appear to be fully active enzymically. Fruchter and Crestfield (580) found that two dimers could be separated on sulfoethyl Sephadex chromatography. The structural difference between these two is not clear; they are both fully active. The probable general structure of the dimers was established in elegant experiments by Fruchter and Crestfield (3881)involving alkylation with iodoacetate. The two isomeric dimers referred to above behave identically in these reactions. The two active sites in the dimers behave just like that of the monomer. Histidines 12 and 119 both react, but the reactions are mutually exclusive. The proposed structure is outlined in Fig. 19. The “tail” of one monomer combines with the “body” of the other and vice versa. The His 12 and 119 pairs are now on separate molecules. When the dimers, fully inactivated by reaction with iodoacetate, are dissociated by heating a t neutral pH, the following monomers would be expected: native RNase (active), CM-His-12-RNase (inactive) CM-His-119(inactive). These RNase (inactive), and di-CM-His-12-His-119-RNase were, in fact, found. About 25% activity reappeared from the inactive dimer. Equally important the di-CM compound was found. This material Homologous dimers I

ii

iii

..;.........;..

; .ill9 ..........

..i

. ......... .

Hybrid dimers iv

;...119

f

.

(A/119)

V

vi

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

119 .i .:........

(A/12)

vii

.. (12/119)

...

ill9 i 12

......

(A/12-119)

FIG.19. Diagrammatic representation of the homologous and hybrid dimers of RNase-A and its carboxymethylhistidine derivatives. In each dimer one polypeptide chain is represented by a dashed line and the other by a continuous line. Carboxymethylhistidine residues are indicated by the solid ovals. Sites containing one or more residues of carboxymethylhistidine are catalytically inactive. Unalkylated, and therefore active, sites are indicated by the stippled ovals. Reproduced from Crestfield and Fruchter (382). 380. R.G.Fruchter and A. M. Crestfield, JBC 240, 3868 (1965). 381. R. G.Fruchter and A. M. Crestfield, JBC 244 3875 (1965).

746


does not appear on alkylation of monomer RNase under the normal conditions. Hybrid dimers have been prepared from alkylated monomer fractions (38.2) and the native enzyme in various permutations. The products gave the predicted activities. I n particular a dimer prepared from Chl-His-12RNase (inactive) and CM-His-119-RNase (inactive) had 50% of the specific activity of RNase-A. This activity was lost on dissociation of the dimer. All of these experiments together strongly confirm the proposed model. This also fits nicely into the observed behavior of the RNase-S system as outlined in Fig. 7.

VI. Catalytic Properties

A. NATUREOF

THE

REACTION CATALYZED

Both the base-catalyzed and the RNase-catalyzed hydrolysis of polyribonucleotides take place in two steps: (1) a chain cleavage or depolymerization step resulting in a 2’:3’-cyclic phosphate terminus and a free 5’-OH group on the other side of the bond cleaved, and (2) hydrolysis of the cyclic phosphate to yield the free 3’-phosphate monoester group. These reactions are shown in Fig. 20. The base B, will normally be a pyrimidine, uracil, or cytosine. The base B, can be either a pyrimidine or a purine. More detailed discussion of the specificity is given below. It was shown by Hilmoe et al. (383) that P-0 bond cleavage rather than G O bond cleavage occurs in RNase action. Cyclic phosphates are mandatory intermediates in the RNase-catalyzed reaction (384) and can be isolated in good yield. The two steps in the reaction are quite separate and ordinarily differ by orders of magnitude in rate. There is no detectable activity toward DNA although deoxynucleotides do bind to the enzyme. Thermodynamically, step 1 is easily reversible while step 2 is not. The reverse reaction of step 1 consists of the formation of a phosphodiester from a pyrimidine cyclic phosphate and a primary alcohol, either a simple aliphatic alcohol or a nucleoside or nucleotide (386-387). The 382. A. M. Crestfield and R. G. Fruchter, JBC 242, 3279 (1967). 383. R. J. Hilmoe, L. A. Heppel, S. S. Springhorn, and D. E. Koshland, Jr., BBA 53, 214 (1961). 384. K. Brocklehurst, E. M. Crook, and C. W. Wharton, Chem. Commun. No. 2, p. 63 (1967). 385. L. A. Heppel and P. R. Whitfield, BJ 56, ii (1954). 386. L. A. Heppel, P. R. Whitfield, and R. Markham, BJ 58, iii (1954). 387. L. A. Heppel, P. R. Whitfield, and R. Markham, BJ 60, 8 (1955).

24.


747

synthesis occurs only with primary alcohols (388). Incubation of a phosphodiester such as cytidine-3’-benzyl phosphate with methanol in the presence of RNase will lead to formation of the methyl ester by exchange (389).These reactions are not surprising in view of the energetics. I n a series of papers, Bernfield has reported such synthetic reactions for the preparation of specific oligonucleotides (390-592). The remarkable aspect of this work is not that it was possible with the native enzyme but that it was possible with three enzyme derivatives normally considered devoid of step 1 and step 2 activity, S-protein, l-CM-His-119RNase, and DNP-Lys-41-RNase. Each of these derivatives showed reduced “synthetic” activity but much more markedly reduced step 2 activity. Thus the yield of synthesized oligonucleotides was much higher than with RNase-A or RNase-S. Interestingly the inactive derivative PIR showed no synthetic capacity. Although the enzyme preparations used were not completely pure, especially the His 119 and Lys 41 derivatives, the observed activities could not be attributed to contaminating native enzyme. It is axiomatic that an enzyme in trace amounts cannot shift a thermodynamic equilibrium. If catalysis occurs in one direction, it must also in the other for a single step process. Each of the enzyme derivatives used is deficient in 1 of the 3 residues thought to be essential to the mechanism of action of the native molecule. Unfortunately, the nature of the synthesis and hydrolysis of a single phosphodiester bond in properly protected compounds was not tested. The mystery behind these experiments remains to be clarified. None of the detailed mechanisms to be discussed considers the macromolecular association that may be involved in the action of RNase on high molecular weight polyribonucleotides. Preiss reported from light scattering studies that very large RNA-enzyme aggregates may be formed (393).Their significance for the catalytic mechanism is unknown.

B. ASSAYSFOR ENZYMIC ACTIVITY A wide variety of procedures have been used for assaying ribonuclease activity. Some measure only step 1 or step 2, some measure both, some use high molecular weight substrates, some low, some substrates are 388. (1957). 389. 390. 391. 392. 393.

G. R. Barker, M. D. Montague, R. J. Moss, and M. A. Parsons, JCS p. 3786

L. A. Heppel and P. R. Whitfield, BJ 60, 1 (1955). M. R. Bernfield, JBC 240, 4753 (1965). M. R. Bernfield, JBC 241, 2014 (1966). M. R. Bernfield and F. M. Rottman, JBC 242, 4134 (1967). J. W. Preiss, Biophys. J. 8, 199 (1968).

748


His 119

Lys 41

1

CpA

+

H,O

It C>p

-lAI1 + A H,O

+

t 3’Cp

+

A

FIO.20. The diagram at the top is a schematic view of the active center as deduced from the X-ray data from the protein and several substrate related complexes. BI, RI, pl, R,, and BZ indicate the relative positions of the bases, riboses and phosphate of the dinucleotide analog UpcA. Position pl is occupied by SO-: in the protein crystal. CMP, UMP, and analogs of these occupy BI, R1, and pl p occupy predominantly. %-AMP occupy B2, R, and pl while 3’-AMP and 3’5’-A Bz and R, predominantly, and possibly t o a lesser extent, B1 and RI. Bz’ is the probable position of the second pyrimidine in dinucleotides such as CpU. The phosphate position in C p cannot be observed owing to digestion but would be a t pl if the base occupies the same position as in CMP. Four His 119 positions are indicated. I coincided with pl but is a possible position in the absence of SO?or nucleotides. I1 is behind I11 and may be occupied by solvent. I11 is slightly stabilized by 3’-CMP. IV is the position occupied when Bz and R, are occupied by adenosine phosphates. His 12 is behind pl and R I . There is a solvent molecule, presumably water, behind pl as indicated by H20. Lys 41 enters from the upper right and is not in contact with P I but might contact PI’. Asp 121 enters from

>

>

24.


749

clearly defined, others are mixtures; different buffers and different values of pH and ionic strength are used. Work on this enzyme spans the period of the establishment of the structure of RNA and many procedures no longer used were historically important in that context. The most commonly used assay procedures today are based on commercial yeast RNA or on cytidine-2' :3'-cyclic phosphate, C>p, as substrates. In the Kunitz procedure (394) the decrease in absorbance a t 300 nm of an RNA solution is measured as a function of time. The decrease primarily results from the hypochromicity of the cyclic phosphate relative to either oligonucleotides or free nucleotides. Measurements are normally done a t room temperature in 0.1 M acetate buffer at pH 5. This pH is well removed from the point of maximum activity, 7.0-7.5. Why pH 5 is routinely used is not clear, the rate a t pH 7 being much higher. The decrease in absorbance is linear only for very short times and thus calibration of each substrate solution is usually required. The procedure is fast and simple. Since step 1 is much faster than step 2, the Kunitz assay largely reflects the step 1 process. Hydrolysis of the cyclic phosphate end groups is, of course, occurring concurrently to some extent and thus affecting the absorbance change. I n the Anfinsen procedure (43) the acid-soluble nucleotides produced after a fixed time of RNA digestion are measured. The undegraded RNA is precipitated with perchloric acid or a mixture of uranyl acetate and trichloroacetic acid. The absorbance of the supernatant solution after filtration or centrifugation is used as a measure of activity. Here also the assay is a measure largely of step 1 activity. With cytidine cyclic phosphate as a substrate the step 2 process alone is measured. The procedure of Crook et al. (395) relies on the increase in absorbance a t 286 nm which occurs on hydrolysis of the cyclic phosphate ring. Cleavage of the ring also results in proton release which is the basis of the titrimetric procedure described by Stark and Stein ( 1 3 2 ) . The assays employing C > p are usually carried out in 0.35 M NaCl. The ionic strength for maximum activity is much higher for this substrate than for RNA. A brief summary of the above procedures is given by Klee (396). 394. M. Kunitz, JBC 164, 563 (1946). 395. E. M. Crook, A. P. Mathias, and B. R. Rabin, BJ 74, 234 (1960) 396. W. A. Klee, Procedures Nucleic Acid Res. p. 20 (1966).

bdow and may contact His 119 in position IV. The reaction diagrams are meant to parallel the active site diagram. The arrows indicate physical motions of atoms rather than electron shifts. The presence of A is indicated during the hydrolysis step since i t is a known stimulant ns well as a potential acceptor in competition with water.

750


More critical examination of the step 1 process can be made employing well-defined dinucleoside phosphate substrates or a variety of other diesters. Extensive studies have been carried out by Witzel and are referred to below. Many other assay procedures have been described for a variety of different purposes. A list of some of these is given in Table XXIII (43, 103, 380,394, 395, 397-&S).

C. SPECIFICITY IN

THE

ENZYME-CATALYZED REACTION

The studies during the 1940’s and 1950’s on the specificity of RNase were closely connected with the elucidation of the structure of RNA itself. 397. J. A. Bain and H. P. Rusch, JBC 153, 659 (1944). 397a. C. A. Zittle and E. H. Reading, JBC 160, 519 (1945). 398. H. Edelhoch and J. Coleman, JBC 219, 351 (1956). 399. A. M. Crestfield, Anal. Chem. 28, 117 (1956). 400. L. Vandendreische, Acta Chem. Scand. 7 , 699 (1953). 401. J. S. Roth and S. W. Milstein, JBC 196, 489 (1952). 402. S. R. Dickman and K. Trupin, ABB 82, 355 (1859). 403. W. Fiers and K. M. Moller, Compt. Rend. Trav. Lab. Carbberg 31, 507 (1960). 404. W. Fiers, Anal. Biochem. 2, 126 (1961). 405. L. Lepoutre, J. Stockx, and L. Vanderdriesche, Anal. Biochem. 5, 149 (1963). 406. D. Shugar, Bull. Acad. Polon. Sci., Classe (II) 1, 39 (1953). 407. R. Shapira, Anal. Biochem. 3, 308 (1962). 408. V. G. Konarev and S. N. Amirkhanova, Biol. Nukleinovogo Obmena u Rust.,

Akad. Nauk SSSR, Otd. Biol. Nauk, Bashkirsk. Filial, Inst. Biol., Dokl. Ob’edin. Nauchn. Sessii, 1968 p. 137 (1959). 409. J. C. Houck, ABB 73, 384 (1958). 410. E. J. Altescu, Anal. Biochem. 8, 373 (1964). 411. J. Martin-Esteve, P. Puig-Muset, and F. Calvet, Rev. Espan. Fisiol. 12, 243 (1956). 412. H. von Heineeke, Z. Naturforsch. 12b, 527 (1957). 413. J. Polatnick and H. L. Bachrach, Proc. SOC.Exptl. Biol. M e d . 105, 486 (1960). 414. K. W. Mundry, Technicon Sump., 2nd N . Y., p. 612 (1966). 415. H. Barrera, K. S. Chio, and A. L. Tappel, Anal. Biochem. 29, 515 (1969). 416. F. M. Richards, Compt. Rend. Trav. Lab. Carlsberg, Ser. Chim. 29, 315 (1955). 417. J. Stockx, Arch. Intern. Physiol. Biochim. 68, 417 (1960). 418. R. Sperling and I. Z. Steinberg, BBA 159, 408 (1968). 419. J. T. Nodes, BBA 32, 551 (1959). 420. L. Josefsson and S. Lagerstedt, BBA 76, 471 (1963). 421. S. B. Zimmerman and G. Sandeen, Anal. Biochem. 10, 444 (1965). 422. F. Molemans, M. Van Montagu, and W. Fiers, European J . Biochem. 4, 524 (1968). 423. P. L. Ipata and R. A. Felicioli, FEBS Letters 1, 29 (1968).

24.

75 1


TABLE X X I I I PROCEDURES FOR RIBONUCLEASE ASSAY Substrate RNA

Property and/or variable Proton release Absorbance Volume change Acid-soluble nucleotides

Method or condition

Ref.

Manometric Ti trimetric Change a t 300 nm Dilatometry Acid-alcohol precipitant Perchloric acid Uranyl-acetate-trichloracetic acid Uranyl-acetate p H 4 BaClO4 Cellosolve Coprecipitation problems Methylene blue-AAaso nm Pyronine-fluorescence A400-7w nm (sensitivity 1 ng) With neomycin sulfate Precipitate with trichloroacetic acid (Sensitivity 100 ppg)

+

RNA-dye binding Turbidity BSA complex Turbidity Clearing in agar

Virus infectivity Automatic discontinuous or flow procedures U > P Absorbance AAz~o nm C > P Absorbance A A 2 8 0 - 3 ~nm Optical rotation h650-300 Paper chromatography Product separation of products Thin layer chromatography Column chromatography Perchloric acid Acid-soluble products Poly c (sensitivity 10 ppg) 14C extracted into toluene Beneyl esters Release of benzyl of U or C alcohol Coupled assay with adenine CpA, UpA Adenine release deaminase AAHK

(499)

The work, summarized by Brown and Todd (424), clearly indicated the 3',5' linkage in RNA, the occurrence of 2': 3'-cyclic phosphates as intermediates in the cleavage of RNA, and the high proportion of pyrimidines on the 3' side of the bond cleaved. Subsequent investigations have probed more deeply into the problem of specificity, investigating different parts of the nucleotide structure. 424. D. M. Brown and A.

R. Todd, J . Chem. SOC.p. 52 (1952).

752


1. The Sugar

The carbohydrate moiety of RNA is D-ribose with the /3-D-ribofuranoside ring. The 2'- and 3'-OH groups are cis to each other and easily form the cyclic phosphate intermediate. Although the 2'-deoxynucleotides bind to the enzyme, they only serve as inhibitors. The 2'-OH group is mandatory for the catalytic activity of pancreatic RNase. The sugar configuration about the l', 2', 3', and 4' positions can be changed by synthesis. A variety of pyrimidine nucleoside cyclic phosphates have been made. Ukita et al. (425) prepared J3-D-lyxo-uridine 2':3'-cyclic phosphate (Fig. 21a). The configuration about the 2' and 3' positions is inverted and the two OH groups are now cis to the base rather than trans as in the D-ribose series. No hydrolysis a t all of this compound was observed in the presence of RNase. However, both the cyclic phosphate and the free 2' (3')-nucleotides inhibit the enzyme. The SUGAR CONFIGURATIONS

T

425. T. Ukita, H. Hayatsu, and K. Waku, J . Biochem. (Tokyo) 50, 550 (1961).

24.

753


BASES Substrates

R

R

S

0

I

HNYNH 0

R

0

R I

Br

0

0

I

R I

cy0 HN- C-CH,

8

(XVI)

Nonsubstrate s

(XVII)

(XVIII)

754


PHOSPHATE DERIVATIVES

0

OH

0-

HZC,p/O o4 \o-

0

I

O=P-

OH 0-

FIG.21. Nucleotide components in substances tested as substrates or inhibitors: (a) sugar components, (b) pyrimidine base components, and (c) phosphate components.

a-D-lyxo compound is also not a substrate but the a-L-lyxo thymidine cyclic phosphate is a substrate (426). I n this latter nucleotide only the configuration about the 4’-carbon differs from the p-D-ribose series. These results are nicely correlated with the X-ray data on nucleotide binding. The pyrimidine biding site requires the @-&rib0 configuration for the 1’-, 2‘-, and 3’-carbon atoms, but puts no restrictions on the configuration a t the 4‘ position. 2. The Base

The following brief summary is considered in more detail in the discussion of mechanism. Nitrogen 3 will normally be protonated as in uracil or unprotonated as in cytosine, serving as a hydrogen donor or acceptor, respectively. Substitution of any other function on this nitrogen atom invariably converts the normal base to a nonsubstrate component. This has been shown by Gilham (427) in the formation of a carbodiimide 426. A. Holy and F. Sgirm, BBA 161, 264 (1968). 427. P. T. Gilham, JACS 84, 687 (1962).

24.


755

adduct, by Cramer et al. (4’38)for the 3-N oxide, and by Massoulie et al. for a 3-methyl group (429). I n most of the substrates there is a keto function in position 2. The 4 position is normally a keto or amino group. Hydrogen bonds involving these functions do appear to contribute to binding the normal nucleotides but they are not crucial. A 4-thio function still provides a substrate. The 5 position can be substituted by a methyl group (thymine) or by any of the halogens (429, 430). While changes are seen in rates of splitting of the homopolymers containing these modified bases, all are substrates for the enzyme. The order of decreasing rates is poly T > poly U > poly iodo U > poly BrU, and poly C1U a t pH 7. I n the binding site derived from the X-ray data, the 5 position of the pyrimidine base is the most exposed and the one most able to accommodate a substituent. The derivative 5-iodouridylic acid was in fact used to locate the active site and to define the ring orientation. 5,6-Dihydrouracil in a nucleic acid serves as a substrate component. The cyclic nucleotide containing this unusual base is hydrolyzed. Pseudouridine also provides substrates in model systems. These observations, along with confirmation of C and U as the principal cleavage sites for RNase, comes from RNA sequence studies, particularly those on tRNA [for recent examples, see Dube et al. (431-433) ; see also Rushizky et al. ( 4 3 3 ~] ); the oligonucleotides had as 3’ end groups C,U,T,H, and +. A summary of some of the base components of known substrates and nonsubstrates is given in Fig. 21b. The question arises as to whether cleavage ever occurs on the 3’ side of purine residues. The hydrolysis of polyadenylic and polyinosinic acids was originally described by Beers (434). The cleavage was very much slower than for pyrimidine-containing nucleic acids but it did occur. The cleavage appeared to be faster with the longer oligonucleotides. The same results were obtained with several different commercial enzyme preparations, but a highly purified enzyme preparation was not tried. 428. F. Cramer, F. Fittler, H. Kuentzel, and E.-A. Schaefer, Z . Naturforsch. 18b, 668 (1963). 429. J. Massoulie, A. M. Michelson, and F. Pochon, BBA 144, 16 (1966). 430. G. R. Barker, M. E. Hall, and R. J. Moss, BBA 46, 203 (1961). 431. S. K . Dube, K. A. Marcker, B. F. C. Clark, and S. Cory, European J . BWchem. 8, 244 (1969). 432. S. K . Dube and K. A. Marcker, European J . Bzbchem. 8, 256 (1969). 433. C. G. Alvino, L. Remington, and V. M. Ingram, Biochemistry 8, 282 (1969). 433a. G. W. Rushiaky, C. A. Knight, and H. A. Sober, JBC 236, 2732 (1961). 434. R. F. Beers, Jr., JBC 235, 2393 (1960).

756


This reaction was investigated in more detail by Imura et al. (495) with carefully purified RNase-A. The patterns of appearance of small oligonucleotides from poly A and poly U were compared (see Fig. 22). Although the rates differed by a large factor, the distributions of products as a function of time were remarkably similar. The cyclic phosphate end group of the A oligomers or of the mononucleotide was hydrolyzed extremely slowly. The ratio V, (poly U)/V, (poly A) is 1300; the ratio V,(U > p)/Vm(A > p) is about 500,000. The Michaelis constants for poly A and poly U differ by only a factor of 3 and are of the order of 1 0 - ~M . The effects of ionic strength on the rates of poly A and RNA hydrolysis are very similar showing a maximum a t 0.1 M , and both activities are reduced in parallel on carboxymethylation of His 119. There appears to be no doubt that A oligomers can be hydrolyzed but very inefficiently. In an earlier paper, Takemura e t al. (4%) investigated the hydrolysis of ribo-apyrimidinic acids by commercial RNase and by isolated RNaseA and RNase-B. They observed no detectable hydrolysis with the purified enzymes and concluded that the observed action with the commercial sample resulted from a minor contaminant. This conclusion also would

FIG.22. Comparison of the depolymerization of poly A and poly U by RNase-A. The incubations were performed in parallel a t 37", pH 72, potassium phosphate buffer 4 mM: (a) the poly A sample was digestrd for 5 hr and (b) the poly U sample was digested for 1 hr with the indicated amounts of enzyme. The various products were separated by paper chromatography. Reproduced from Imura et al. (4361, Figs. 1 and 2. 435. N. Imura, N. I r k , and C . Ukita, J. Biochem. (Tokyo) 58, 264 (1965). 436. S. Takemura, M. Takazi, M. Miyasaki, and F. Egami, J. Bwchem. (Tokyo) 46, 1149 (1959).

24.


757

apply to the data of Zamenhof et al. (437, 438) who reported earlier the hydrolysis of polyribose phosphate. It is possible that the assays were misleading because of the use of uranyl salts as precipitants. Dickman and Trupin (402) showed that only uridine compounds are soluble under certain conditions, all amino-containing nucleotides even monoiiucleotides being precipitated. Unfortunately, Imura et al. (435) did not discuss, or refer to, the paper by Takemura e t al. (436‘) in reference to the poly A observations. Simple substances containing a cyclic phosphate or ribose cyclic phosphate with a P-methyl substituent on the 1’carbon are not substrates (499, 499a). These rates could be just too slow to have been observed and, with the enhancement usually seen on going to polymers, action of polyribose phosphate might occur. In any event, normally all substrates other than those containing pyrimidines are split very slowly if a t all. Very interesting additional data come from studies on poly 3-isoadenylic acid (nonsubstrate) (440) and the unusual purines related to formycin (substrates), F, (441, 442). The activity of the enzyme on t,hese compounds is much higher than that toward the normal purines and is comparable to that toward the pyrimidines. Ikehara et al. (441) have reported the action of RNase on poly (F-C), poly (F-U), and poly (F-G) ; FpUp, FpCp, and GpFp were obtained. With more enzyme the first two yielded F > p and eventually 3’-FMP. Under the digestion conditions used poly (A-G) was not attacked a t all. Reich and his colleagues (442) have found that poly F is an excellent substrate for RNase and also that the nucleosides in the polymer have the unusual syn conformation. In contrast to Ikehara et al. (441), these authors found no evidence for the hydrolysis of F > p where the rate of degradation of poly F to give F > p was about equal to that on poly C to give C > p. The demonstration of the syn conformation was critical for an understanding of these data. The nucleoside can “fit” the pyrimidine binding site when in the syn conformation as discussed in more detail below. 437. S. Zamenhof, G. Leidy, P. L. Fitzgerald, H. E. Alexander, and E. Chargaff, JBC 203, 695 (1953). 438. E. Rosenbcrg and S. Zamenhof, JBC 236, 2845 (1961). 439. T. Ukita and M. Ire, Chem. & Pharm. Bull. (Tokyo) 9, 211 and 217 (1961). 439a. F. Egami, M. Takayi, I. Hayashi, and S. Takemura, Seikagaku 31, 120 (1959). 440. A. M. Michelson, C. Monny, R. A. Laursen, and N. J. Leonard, BBA 119, 258 (1966). 441. M. Ikehara, K. Murao, and S. Nishimura, BBA 182, 276 (1969). 442. D. C. Ward, W. Fuller, and E. Reich, Proc. Natl. Acad. Sci. U. S. 62, 581 (1969).

758


The influence on the kinetics of the ester group in pyrimidine nucleoside-3’-phosphate esters is discussed in the section on steady state kinetics. 3. The Phosphate Group Eckstein (443) has prepared the isomeric derivatives of uridine-2’: 3’-O1O-cyclophosphorothioatewhere one or the other of the two free oxygen atoms is converted to a sulfur atom. The maximum velocity for the hydrolysis of these compounds is one-fifth that of U > p under comparable conditions while the K , value for one is identical to U > p and the other is larger by a factor of 8. Phosphonate derivatives have also been synthesized. These are summarized in Fig. 21c and discussed in the section on mechanism.

D. STABLECOMPLEXES-INHIBITION-ACTIVATION 1. Macromolecular Inhibitors

Most polyanions, natural and synthetic, show some inhibitory effect on RNase (see, e.g., references 444-451). The interaction in many cases is largely electrostatic and is markedly reduced a t high ionic strength. The distribution of negative charges affects the inhibitory activity, and the inhibition can be relieved by adding polycations such as protamine or polyornithine. Inhibition is usually greatest near pH 5 , well on the acid side of the pH of maximum activity. The cyclic phosphate activity is frequently inhibited much more than that toward RNA. This may be merely reflection of the tighter binding of the macromolecular substrate. When the inhibition by heparin of step 1 and step 2 was tested with ethyl or benzyl esters of cytidine 3’-phosphate and cytidine 2‘:3‘cyclic phosphate, the inhibition constants were found to be the same (446, 452). There is no evidence for more than one catalytic center. 443. F. Eckstein, FEBS Letters 2, 85 (1968). 444. L.Vandendreissche, ABB 65, 347 (1956). 445. H. Heymann, Z. R. Gulick, E. J. DeBoer, G. De Stevens, and R. L. Mayer, ABB 73, 360 (1958). 446. J. P. Hummel, JBC 233, 717 (1958). 447. P. T. Mora, B. G. Young, and M. J. Shear, Makromol. Chem. 38, 212 (1960). 448. J. Coleman and H. Edelhoch, ABB 63, 382 (1956). 449. H. Sekine, E. Nakano, and K. Sakaquchi, BBA 174, 202 (1969). 450. P. T. Mora, JBC 237, 3210 (1962). 451. M. Sela, JBC 237, 418 (1962). 452. C. Ukita, T. Terao, and M. Irie, J. Biochem. (Tokyo) 52, 455 (1962).

24. BOVINE

PANCREATIC RIBONUCLEASE

759

The inhibitory effectiveness of a polyanion increases markedly with molecular weight between 5,000 and 12,000 as shown for polyethene sulfonic acid (453). This might mean that a weight of RNA comparable to that of the enzyme was involved in the interaction. There is no direct evidence from chemical or X-ray binding studies for any such large number of sites as this would imply. One of the most interesting synthetic inhibitors is the copolymer of glutamic acid and tyrosine described by Sela (451).As a 1 :1 copolymer this material was more effective than the 9 : l copolymer or than polyaspartic acid, the latter a very good inhibitor. Clearly some noncoulombic interactions are involved here. The effect of the tyrosine residues is perhaps related to the effectiveness of phenolic compounds in preventing recovery of activity on reoxidation of the reduced enzyme (see Section IV,B,7,c). The glutamic-tyrosine copolymer is an excellent inhibitor a t pH 5 but is almost ineffective a t pH 7 in agreement with the behavior of other anionic polymers. There are about 4 less protons bound to RNase a t pH 7 than a t pH 5 (26‘8). This difference largely results from the 4 histidine residues which are almost fully charged a t p H 5 and almost fully deprotonated a t pH 7. Only 2 of these are a t the active site. The total net charge on the protein is +4 a t pH 7 and +3! a t pH 5. The anionic charge on most of the polymers tested does not change in this pH interval. The importance for inhibition of the specific histidine charges as opposed to the overall net charge could be tested on a partly amino-acetylated enzyme. This does not yet appear to have been done. It would also be of interest to know whether the polymers bind to the enzyme a t pH 7 but without causing inhibition. 2. Small Molecule Effectors

a . Binding of Nucleotides. I n general the interaction of nucleosides is stronger than that of either ribose or the free bases, and nucleotides are more strongly bound than nucleosides. All three elements of the nucleotide are involved in the association. Where association results in inhibition of the enzyme-catalyzed reactions, the effect is usually classified as “competitive.” As will be seen in detail in the discussion below this does not imply identical binding sites for the whole range of compounds such as those listed in Table XXIV (464). Nucleotide and phosphate binding are strongest a t pH 5.5 (Table XXV (455).For phosphate453. M. K. Bach, BBA 91, 619 (1964). 454. T. Ukita, K. Waku, M. hie, and 0. Hoshino, J . Biochem. (Tokyo) 50, 405

(1961). 455. D. G. Anderson, G. C. Hammes, and F. G. Walz, Jr., Biochemistry 7, 1637 (1968).

760

F. M. RICHARDS AND H. W. WYCXOFF

TABLE XXIV APPROXIMATE INHIBITION CONSTANTS FOR SUBSTRATE-RELATED COMPOUNDSO Compound

2-OH-pyrimidine Uracil Uracil 5-methyl Uracil 5-bromo Uracil 5-amino Uracil 5-nitro Uracil 5-cyano Uracil 5-aminoethyl

Ki (d)

Compound

Free Bases 65 Uracil, 5-dimethyl19 (18) aminoethyl 21 Uracil 5, 6-dihydro 24 Uracil 1-methyl > 1000 Uracil 2-thio 6 Uracil 1, 3-dimethyl 14 Uracil 3-methyl 6 Cytosine Cytosine 1-methyl

Ki (mM)

11 36 34 49 > 1000 144 > 1000 >200

Nucleosides Uridine Uridine 5-bromo Uridine 5-methyl Uridine 4-deoxy Cytidine Cytidine 2’-deoxy

17 (13) 23 (14) 11 58 31 (25) 31

Adenosine Inosine Thymidine Thymine-xylof uranoside Thymine-glucopyranoside

>200 >200 16 (50) 1

175

Nucleotides* Adenosine 2’(3’)p Adenosine 5’p Adenosine 2’3’-cyclic p Adenosine 5’ tri p Guanosine 2’(3’)p Guanosine 5’p Guanosine 2‘, 3’-cyclic p Inosine 2’(3’)p Inosine 5’p

Orthophosphate ion Phenylphosphate Ethyleneglycol monophosphate Hydrobenzoin phosphate ~

~~

8 (35) 9 (33) >200 4 4 14 >200 6 6

23 (3) 66 (110) 49

Uridine 2’(3’)p Uridine 5’p

2 (2) 6 (4)

Uridine 2’,3’-isopropylidene-5’p 97 1 Uridine 2’(3’)p, 5-bromo Uridine 5’p, 5-bromo 5 Uridine 5’p, &methyl 9 Cytidine 2’(3’)p 2 3 Cytidine 5’p Lyxo-uridine-2’,3’-p 1ooc Lyxo-uridine-2’ (3’)-p 25” Lyxo-uridine-5’-p 25’ Others D-Ribose

> 1000

29

~

The numbers listed were derived from the “inhibition index” values given by Ukita et al. (454).The substrate was cytidine 2’: S’-phosphate, 8.8 mM. The Michaelis constant was reported to be 24 m M in the bicarbonate buffer 0.03 MI pH 7.6, 37”, that was used in all of the measurements. The inhibition constants were derived on the assumption of competitive inhibition in all cases. This type of inhibition was specifically shown for those compounds with a second number in parentheses. This latter number was obtained from multipoint double reciprocal plots in the usual manner. The agreement of the two

24.


761

containing compounds above pH 5.5 protons are taken up during the association with RNase while below pH 5.5 they are released (456‘). This pH of zero uptake is reported to be independent of the ionization constant of the phosphate group, and no uptake occurs with nucleosides. The phosphate group is apparently intimately involved with other ionizable groups in the association reaction. The pK values of these groups on the enzyme, if considered to be only two, are 5.1 and 6.1 as estimated from the pH dependence of the changes in proton uptake or release. The concentration of the enzyme species directly involved with the association is a t a maximum a t pH 5.5. These observed values would he expected to vary with ionic strength as do the pK values for the histidine residues. There is one binding site on RNase with a high association constant for pyrimidine nucleotides. This was demonstrated by difference spectra for 2’-CMP by Nelson and Hummel (457) and confirmed by Barnard and Ramel (458) by sedimentation. No evidence for a second binding site was seen. With l-CM-His-119-RNase no interaction with 2’-CMP was seen spectrally, and the sedimentation studies indicated that if any M interaction occurred the dissociation constant was greater than compared to 10-BM for the native enzyme. A close relationship between the phosphate group and a histidine residue is thus implied as it was in the proton uptake studies referred to above. Cathou et al. (459) found that the Cotton effect near 270 nm in the ORD spectrum of RNase disappeared on interaction with either 2’-CMP or 3’-CMP. The X-ray studies (120) (see Fig. 23) clearly show that no tyrosine residues are in close contact with the substrate. Thus the change in rotatory behavior must reflect either (1) a shift in protein structure on association of the nucleotide or (2) the induction of a Cotton effect of the opposite sign in the bound nucleotide. In the independent spectral and chemical studies of Irie and S5wada (460), the reduced nucleotide 5,6-dihydrouridine-2’(3’) -phosphate, known to interact with the enzyme, showed no difference spectrum. With nucleotides containing 456. J. P. Hummel and H. Witael, JBC 241, 1023 (1966). 457. C. A. Nelson and J. P. Hummel, JBC 236, 3173 (1961). 458. E. A. Barnard and A. Ramel, Nature 195, 243 (1962). 459. R. E. Cathou, G. G. Hammes, and P. R. Schimmel, Biochemistry 4, 2687 (1965). 460. M. Irie and F. Siiwada, J . Biochem. ( T o k y o ) 62, 282 (1967).

numbers may be taken as a qualitative measure of the reliability of the inhibition index values which come from data a t a single concentration. Where n o inhibition was reported, the values have been listed as >200 or >1000, the estimated detection limits under the conditions employed. * Here p stands for phosphate. From Ukita et al. (486).

TABLE XXV BINDINGOF NUCLEOTIDES TO RIBONUCLEASE-A~ Dissociation constant

Nucleotideb

2'-CMP 3'-CMP

4.0

4.5

5.0

indicated pH

5.5

5.5c

6.0

3.4 37

7.0 103

36 333

9 133

2'-UMP 3'-UMP

256

111

5.6 83 7.1 70

Pyrophosphate Orthophosphated

233 11.8

154 8.6

115 4.25

260

(pM) at

82 172 4.6

6.5

Ionization constants

7.0

20 192

147 625

164

435

385 6.5

1333 14.5

7.5

Phosphate

Ring N3

1100

6.02 5.90

4.32 4.32

5.74 5000 42

5.97 6.67

-

Solvent: 0.05 M tris, 0.05 M sodium acetate, 0.1 M KNOa, p H adjusted with acetic acid, 25". All nucleotide-RNase complexes shown to be 1:1 by method of continuous variations employing difference spectra measured a t the wavelength of maximum difference. c Same as footnote a except KNOa replaced with KCl (465). d Units, for orthophosphate only, mM. a

b


24.


763

4-thiouracil the spectral changes are seen but the wavelength is increased by 70 nm to the absorption region characteristic of the sulfurcontaining nucleotide ( 4 6 1 ) . The observed difference spectra thus appear to result from changes in the nucleotide chromophores and not from changes in the protein. Irie (462)compared the difference spectra of RNase complexes with various nucleosides and nucleotides with those obtained with the free ligands in various aqueous organic solvent mixtures. The observed red shift and hypochromism suggest a protein environment for the bound base with a higher refractive index than water and in addition the presence of an aromatic residue. The latter could be Phe 120 which the X-ray data (462a) show to be close to the pyrimidine base of the bound nucleotides. From studies on the effects of pH on nucleotide spectra, Deavin et al. (46’6s) concluded that a basic group on the enzyme, perhaps COO-, interacts with position 3 or 4 of uridine derivatives. Cytidine effects were not interpretable on this basis, and the X-ray data provide evidence for aliphatic OH groups near the 3 and 4 ring positions in the principal pyrimidine binding site and the COOH of Asp 83 distal to the OH of Thr 45 ( 4 6 ‘ 2 ~ )The . purine difference spectra were seen to be comparable to protonation of N7 of the purine ring. Although they attribute this to the presence of a conjugate acid, an alternate explanation would be an increase in pK of N7 caused by an adjacent COO-. Aspartic acid 111 is a prime candidate for this function and Asp 121 is also available. The effect of pyrimidine nucleotide binding on the apparent pK values of the histidine residues is shown in Table XXVI as estimated from the C2 proton resonances by NMR (464, 465). Marked upfield shifts are 461. F. Siiwada and F. Ishii, J . Biochem. (Tokyo) 64, 161 (1968). 462. M. hie, J . Biochem. (Tokyo) 64, 347 (1968). 46%. F. M. Richards, H. W. Wyckoff, and N. Allewell, in “The Neurosciences: Second Study Program” (F. 0. Schmitt, ed.), p. 901. Rockefeller Press, New York, 1970. 463. A. Deavin, R. C. Fisher, C. M. Kemp, A. P. Mathias, and B. R. Rabin, Biochem. 7, 21 (1968). European .I. 464. D. H. Meadows and 0. Jardetsky, Proc. Natl. Acad. Sci. 61, 406 (1968). 465. D. H. Meadows, G . C. K. Roberts, and 0. Jardetsky, J M B 45, 491 (1969).

FIQ.23. Stereodrawings of the dinucleotide phosphonate UpcA bound to RNaseS. Histidine 119 is in position IV where i t is forced to be by the adenosine. Lysine 41 is in the position found in DNP-Lys 41 derivative. I n the native protein it is lower and closer to the phosphate position, but it is not long enough to contact t,he phosphate. The UpcA is an interpretation of the 2-A resolution electron density difference map and the CH, which is bound to the phosphorus is crowding His 119. Some further adjustments may be necessary.

TABLE XXVI SUMMARY OF CHEMICAL SHIFT AND pK CHANGES ON BINDING OF INHIBITORS TO RNasen (Solvent:DzO, 0.2 M NaCl, 32”) RNase absorptions Inhibitor absorptions at p H 5.5 Inhibitor None 3’-CMP 2’-CMP 5’-CMP Cytidine 5-Methyl-2’deoxycy tidine Phosphate Sulfate

His 12 Aijb pK 6.2 8.0 8.0 8.0 6.2 6.2 6.9 6.6

His 48 A 8b

His 105 Asb pK

-

-

+10 +10 0 0

-7 - 10 - 10 - 10 5

6.7 6.7 6.7 6.7 6.7 6.7

0 0 0 0 0

0 0

-6 0

6.7 6.7

0

+8

0

His 119 pK A@ 5.8 7.4 >8.0 8.5. The observed relaxation time, T ~ ,for the process E’ + (EH + E ) will be pH dependent. It is assumed in what follows that the ligands either do not interact with (E’ E’H), or, more likely, interact with both forms but not necessarily equally (468). Neglecting the variety of possible ionization processes for each component, the minimal mechanism for the interaction of cytidine nucleotides is shown in Fig. 24 and some related rates in Table XXVII. For each nucleotide there is a bimolecular step followed by an

+

E

’

C

E

E’-E

+

+

CPC

c> P

E’-E

+ 3’-CMP

FIG. 24. Kinetic niechanism for the interaction of substrates and products with RNase. The various bimolecular association steps and isomerization processes are shown. Proton binding and p H dependence are not indicated. [Adapted from Hammes ( 4 6 6 ) , Fig. 2 ; note that second isomerization originally inserted between EP2 and ES, was probably the result of a second binding site a t high 3’-CMP concentration; see Hammes and Walz (4681.1 467. T. C.French and G. G. Hammes, JACS 87, 4669 (1965). 468. G.G.Hammes and F. G. Wala, Jr., JACS 91, 7179 (1969).

767

24. BOVINE PANCREATIC RIBONUCLEASE

TABLE XXVII RATECONSTANTS AND RELAXATION TIMESFOR OF

Nucleotide

pH

3'-UMP

4.5 5.0 5.5 6.0 6.5 7.0 5.0 6.0 6.6

3'-CMP

U>P

6.0 6.5

3'-CMPa 2'-CMP CpCb c > Pc

6 6 6 6

(I

THE INTERACTION NUCLEOTIDES WITH RIBONUCLEASE-A

10-7 kl (M-I sec)

lo-' k--5 (sec-1)

10-8 ki (sec-1)

0.2 M KC1 25" [Ref. (46'8)l 4.0 1.0 5.7 1.0 1.0 7.8 1.1 1.0 6.1 1.1 1.0 4.0 1.3 0.7 0.6 0.9 1.6 4.2 0.42 4.6 0.42 1.2 0.62 0.2 M NaCl 15' [Ref. (470)l 2.8 1.1 2.1 0.9 0.1M KNOI 15" [Ref. (46'8)l 6 0.4 -1 -3 1.4 7 2-5 10-20

-

10-8 k_t (sec-1)

1.8 1.2 0.8 0.6 0.2

10-8((1/~) (set-1)

1.8

1.8 2.6

-

1.1 8.6 12

Original reference 471. Independent of pH, range 6-7, errors ~ 2 5 %original ~ reference 472. Independent of pH, range 5.5-7, errors -25%,, original reference 473.

isomerization. Cytidine is released from CpC a t the rate limiting step. More detailed data are available for the steps involving C > p and 3'CMP, in particular the p H dependence (466). A possible mechanism for the interaction of the enzyme with 3'-UMP is shown in Fig. 25a (468). The observed pH dependence on the rate k, can be reasonably well accounted for if pKA1 = 5.4 and pKB, = 6.5 and (kzl kZ1Ix)= 2.5 x lo8 M-l sec-l. At least two ionizable groups on the free enzyme are implicated. The dissociation rate constant does not reflect any groups on the enzyme-nucleotide complex that ionize in the pH range 4.5-7. The pH dependence of the isomerization of the complex (Fig. 25b) requires a third ionizing group and can be adequately fit if k,' = 1000 sec-', kil = 420 sec-l, = 4200 sec-1 k-: = 2100 sec-l, k:' = 280 sec-l, k,"' = 2800 sec-', pKBz = 7.5, pKcz = 8.0, pKBs = 8.0, pKD3= 5.8, and pKAz, pKA3< 3.5. Other sets of constants would also fit the data, but

+

768

F. M. RICHARDS AND

H.

W. WYCKOFF

this set is internally consistent and agrees quite well with the equilibrium data on the binding of 3'-UMP. It is likely that the same general scheme is applicable to the binding of other nucleotides. Hammes (468) has drawn these general conclusions: (1) At least three ionizable groups on the enzyme are necessary for full catalytic activity. (2) Parallel reaction paths must be postulated (i.e., more than one ionized form of the enzyme must react).

HEH3P

E

HE&P

EHP

KB2

3

t

k!

KA2

1 ",%

HE'H3P

11

k-3

I

lKA3

HE'H,P

k."

KD3

3

C H E ~ H ~E'H,PP + ~H' C

E

EP (a)

i=o

i=o

(b)

FIG.25. Kinetic mechanisms for nucleotide binding including proton ionization steps. (a) Proposed mechanism for the initial interaction of RNase with 3'-UMP. PH is the monoanionic nucleotide, and P is the dianionic nucleotide. The horizontal arrows represent the kinetically significant steps in the pH range 4.5-7.0 (free protons are not indicated on the diagram). (b) Proposed mechanism for the isomerization of enzyme :3'-UMP complexes. Reproduced from del Rosario and Hammes (470).

(3) Initial binding of all substrates is followed by an isomerization or conformational change with a characteristic time constant of the order of magnitude of lo3 sec-I. (4) The second-order rate constants approach, but do not reach, the values expected for a diffusion-controlled reaction. Of the three ionizable groups on the free enzyme, two appear to interact directly with the nucleotides (pK values 5.4 and 6.5 a t 25", 0.2 M KCl) while one is affected by the association but is not in contact

24.


769

with the ligand (pK 5.8). Hammes and Wale (468) suggested that these may be His 119, 12, and 48, respectively. These numbers and assignments may be compared with those from the NMR titration data (see page 712 Section V,A,10) His 119 = 5.8, His 12 = 6.2, and His 48 = 6.4. The agreement is not particularly good, but the kinetically determined values do depend on the mechanism, and the residue identification is only by inference. The X-ray data indicate considerable mobility for His 119, and this residue might easily be involved in the isomerization reaction of the complexes although perhaps less clearly in the free enzyme. Myer and Schellman (469) have measured the binding of 5'-AMP by equilibrium dialysis. The data appeared to indicate two noninteracting binding sites characterized by a single dissociation constant. At pH 7.5 in 0.05 M tris buffer the value of the constant was 3mM a t 30". AH for the association reaction was -4.2 kcal/mole and A S = -2.0 eu. There is no evidence for two equivalent binding sites for any other nucleotide.

b. Simple Ions. Ribonuclease shows no tendency to bind chloride ions a t the isoionic point, pH 9.6. As protons are added during an acid titration, chloride ion binding increases. At pH 6.6 there is less than one chloride bound. At p H 4.5 the value has risen to about 2, and a t pH 2.6 the value is about 4.7 when the free chloride concentration is 35 mM. In the region acid to pH 4.5 one chloride ion appears to be bound for every two protons taken up. Saroff and his colleagues (474, 475) have interpreted these data in terms of the clustering of positive charges with carboxyl groups nearby. As the carboxyl groups become protonated and lose their negative charge, the high local electrostatic potential comes into effect and chloride binding occurs. Such clustering would lead to the abnormal carboxyl titration data discussed in Section V,A,10. The alkaline titration would appear normal since the carboxyl groups are ionized and the local net charge would not be high. The positive charge clusters were assumed to correspond to 3 basic residues each. Six such clusters were postulated to explain the data. 469. 470. 471. 472. 473. 474. 475.

Y. P. Myer and J . A. Schellman, BBA 55, 361 (1962). E. J. del Rosario and G . G . Hammes, JACS 92, 1750 (1970). R. E. Cathou and G. G . Hammes, JACS 87, 4674 (1965). J. E. Erman and G . G. Hammes, JACS 88, 5614 (1966). J. E. Erman and G . G . Hammes, JACS 88, 5607 (1966). H. A. Saroff and W. R. Carroll, JBC 237, 3384 (1962). G. I. Loeb and H. A. Saroff, Biochemistry 4, 1819 (1964).

770

F. M. RICHARDS A N D H. W. WYCKOFF

The three-dimensional structure does show regions of high positive charge, especially around the active site, but clustering into the discrete groups suggested by Saroff is not so clear; (Lys 7, Arg 10, Arg 39) and (His 12, Lys 41, His 119) form two such groups although they are all so close together that the division is somewhat arbitrary. All the other positive charges can only be put together in pairs a t most, for example, (Lys la, Lys l e ) , (Arg 85, Lys 98) or (Arg 85, Lys 66), (His 105, Lys 61) or (His 105, Lys 104). The variable activity of RNase toward different RNA preparations has been tracked down in part to the variable metal content of the substrates [see Wojnar and Roth ( 4 7 6 ) ,and earlier references quoted]. Takahashi et al. (477) have reported that MgZ+,Ca2+,and Mn2+ have little or no effect on step 1 or step 2 activity when these are assayed with low molecular weight substrates. However, Ca2+and Mg2+do interact with RNA and they inhibit the RNase-catalyzed reaction a t pH 7 because of this interaction with substrate (478). Eichhorn et al. (479) found activation by Mg2+and various transition metals a t p H 5. In any event it is clear that in general each metal can be expected to show different effects as a function of pH, ionic strength, specific buffer effects, etc. A substantial correlation of much of the data has been made by Alger (480) who studied RNA and C > p substrates over wide ranges of metal concentration. Activation appeared to involve predominantly metal-substrate interactions while inhibition occurred with direct enzyme-metal interaction. The most actively investigated cations are Zn2+ and Cuz+. Although these are common contaminants of RNA and undoubtedly interact with the nucleic acid, they also interact directly with the enzyme and inhibit step 1 and step 2 even with low molecular weight substrates. The greatest divergence of opinion concerns the maximum number of binding sites. This problem has been studied most intensively with copper where the binding has been studied by equilibrium dialysis, gel filtration, spectroscopy in the visible region, NMR, EPR, and magnetic proton relaxation rate. The results depend more on the laboratory of origin than the method. All seem to agree on a single strong site, but the number of secondary sites varies from one to four. Some of the data for copper are summarized in Table XXVIII ( 4 8 - 4 8 4 ) . The binding of zinc ions 476. 477. 478. 479. 480.

R. J. Wojnar and J. S. Roth, BBA 87, 17 (1964). T. Takahashi, M. h i e , and T. Ukita, J. Biochem. ( T o k y o ) 61, 669 (1967). G. A. Morrill and M. M. Reiss, BBA 179, 43 (1969). G. L. Eichhorn, P. Clark, and T. Edward, JBC 244, 937 (1969). T. D. Alger, Biochemistry 9, 3248 (1970).

24.

771


TABLE XXVIII BINDING OF Cu(I1) ~~

First site Preparation RNase-Ab l-CM-His-119RNase-Ab 3-CM-His-12RNase-A RNase-S &Protein

pH

Solvents

5.0 5.5 7.0 5.0 6.1 7.0 5.0

1 2 3 1

5.0 5.0

Secondary sites

K D (d) Ki (d) 7L. K D (d) 0.73 0.67

0.36

2 4 4 1 1 1

8 8.3 0.053 8 0.064 0.0012

0.60

2 1

7 4

0.002

1.0

4

4 1

0.62

1 1

0.65 0.61

Solvent and references: 1, (481);0.16 M NaC1,22", no buffer ions; K ; valuesmeasuredwithc > p as substrate. 0.11 M KCl, 0.05 M acetate, 25". Constants refer to total of (Cu*+ CuAc+) 2, (482); as the "free" metal ion concentration. 3, (482); 0.11 M KCl, 0.05 M acetate, 0.02 M N-ethyl morpholine-HC1, 25". 4, (483);Ionic strength 0.1 M maleate buffer 25", 8-alanine-Cu*+ metal ion buffer. Values for the dissociation constants for RNase-A reported by Saundry and Stein (484) tend to be about 10-50 times smaller than the values given here. The observations were made a t Z = 0.1, 25", in maleate, acetate pH buffers, and 8-alanine or a metal ion buffer. Evidence for one strong site and one weak site only was obtained. a

+

is similar, but dissociation constants are larger by factors of 25-100 than the corresponding values for copper. Ross et al. (485) reported evidence for the existence of a ternary complex of RNase, Zn2+,and 3'-CMP. They also implicated a histidine residue as one of the protein ligands. These observations have been confirmed in more recent studies with both Zn2+and Cu2+.These metals enhance the interaction of the enzyme with 3'-CMP and vice versa; however, 2'-CMP seems to compete with the strong Cu2+site (482, 486). The enhancement of the proton relaxation rate on copper binding, as , be used to infer details of the studied by Joyce and Cohn ( g l ) can protein environment a t the binding site. The value of the enhancement for the strong site in RNase-S was less than in RNase-A indicating a 481. 482. 483. 484. 485. 486.

B. K. Joyce and M. Cohn, JBC 244, 811 (1969). A. W. Girotti and E. Breslow, JBC 243, 216 (1968). R. H. Saundry and W. D. Stein, BJ 108,583 (1968). R. H. Saundry and W. D. Stein, BJ 105, 107 (1967). C. A. Ross, A. P. Mathias, and B. R. Rabin, BJ 85, 145 (1962). E. Breslow and A. W. Girotti, JBC 241, 5651 (1966).

772


more flexible structure. The changes in this factor in the various other derivatives were complex and not amenable to simple correlation. It is not clear why the number of binding sites should differ from the values of Girotti and Breslow (482). The binding of Znz+and Cu2+markedly lowers the stability of RNaseA to both thermal and urea denaturation (487).

c. Miscellaneous Inhibitors. Naturally occurring inhibitors of RNase have been isolated from a variety of tissues (see, e.g., references 488490). The significance of their inhibition of the bovine pancreatic enzyme is unclear. A diverse collection of substances has also been found to vitamin B I Z (492), inhibit the enzyme; for example, penicillin (PI), mercury hematoporphyrin (493), beryllium chloride (494), and putrescine (495).

E. STEADY STATEKINETICDATA 1. Michaelis Constants and Turnover Numbers

The first intensive investigation of the kinetics of step 2 was carried out by Herries et al. (496) in a study of C > p hydrolysis. The data gave linear double reciprocal plots and maximum velocities and Michaelis constants were measured as a function of pH. Similar studies on U > p have been carried out by others (497, @8), but these did not agree well with each other or with the later work of del Rosario and Hammes (499). I n one case no indication was given of substrate purity and 1/15 M sulfate was employed (497). I n the other product contamination was clearly a problem (498). Some data are shown in Fig. 26. The analysis has been carried out on the assumption of two ionizable groups in the enzyme.

487. 488. 489. 490. 491. 492. 493. (1962).

C. L. Hereig and C. C. Bigelow, BRRC 26, 645 (1967). K . Shortman, BBA 55, 88 (1962). J. St. L. Philpot and J . E. Stanier, BJ 87, 373 (1963). J. S. Roth and D. Hurley, BJ 101, 112 (1966). L. Massart, G. Peeters, and A. Vanhoucke, Ezperientia 3, 494 (1947). R. Llamas, Anales Inst. Biol. (Univ. Nacl. Mez.) 26, 1 (1955). Y. Miura, A. Fukuda, A. Nishimura, and S. Tanaka, Igaku N o Ayumi 40, 1

773

BOVINE PANCREATIC RIBONUCLEASE r

I

I

I

1

I

I

3

4

5

6

I

I

7

I

8

I

9

PH

(b)

- 2 1 " " " 4

5

,

6

7

I

,

0

PH (C)

FIQ.26. Dependence on p H of the steady state kinetic parameters for various substriitcs. (a) Hydrolysis of C > 11 at 25". I =0.2. ( 0 )log ( k . / K d ; (0) pk,; ( A ) log k. [reproduced from Herries et nl. (Q96)I. (b) Hydrolysis of: (0) U p; ( A ) C p (step 1). Cleavage of: Q UpU; 0 UpA; [ICpA (step 2). The solid curves were_ ciculated from Eq. (3) using: pK. = 5.4,ph'b = 6.4,and the following values of k , / K , in W ' sec-': U > p, 2940; C > 1'. 7540; UpU, 388 X 10'; UpA, 6.8 X W ;and CpA, 1.59 X 10'. (c) Hydrolysis of U > p: (0) log k.; ( 0 )pK.. The lines were calculated from equations based on the mechanism shown in Eq. (2). The following values were assumed: pK. = 5.4,pKb = 6.4, pK.' = 5.8,~ K L= ' 7.5, 6 = 4.9 sec", K. = 1.67 mM, and kr = 0.19 sec-', where kr is the turnover number for path A. [(b) and ( c ) are reproduced from del Rosario and Hammes

>

>

(499).I ~~~~

~~

~~~~~

494. G.Santacroce and F. Costabile, Boll. Soc. Ital. Biol. Sper. 42, 1023 (1966). 495. L. A. Nezgovorova and N. N. Borisova, Fiziol. Rust. 14, 644 (1967). 496. D.G.Herries, A. P. Mathias, and B. R. Rabin, BJ 85, 127 (1962). 497. E.N. Ramsden and K. J. Laidler, Can. J. Chem. 44, 2597 (1966). 498. C-C.S. Chcung and H. I. Abrash, Biochemistry 3, 1883 (1964). 499. E.J. del Rosario and G . G. Hammes, Biochemilty 8, 1884 (1969).

774


W.

WYCKOFF

For a single substrate-single product reaction with one path (assume path A is insignificant), the pH dependence of k,/K, reflects the ionization of groups on the free enzyme (K,, Kb) and free substrate and is not affected by any of the intermediates. The pH dependencies of k, and K, separately are more complicated and, as well as K, and Kb, involve K,' and Kb' which depend on the number of intermediates and steady state rate constants. Using the terminology of del Rosario and Hammes (499).

Ic. -L _ Ks Rs[1 -I- (H+>/Ko-I- Kb/(H+)]

(3)

where Ic, is the observed turnover number and K , the observed Michaelis and K, are the pH independent values of the same variables. constant ; is

The pH dependence of k,/Ks for a number of substrates is shown in Fig. 26b. The lines through the points have been calculated from Eq. (3) with a single pair of values of K , and Kb. The fit is quite good in all cases and indicates that the same ionizable groups on the free enzyme are involved in each case, and, in particular, that step 1 (UpU, UpA, and CpA) and step 2 ( U > p and C > p) appear to involve the same groups. TABLE XXIX AND pH INDEPENDENT KINETIC PARAMETERS IONIZATION CONSTANTS pH Independent kinetic parameters

Ionization constants

Substrate

c > pa c > pb u > pb UpUb UpAb CpAb

Free enzyme PKa PKb 5.2 5.4 5.4 5.4 5.4 5.4

6.8 6.4 6.4 6.4 6.4 6.4

ES complex(s) PKd

PKb'

6.3

8.1

5.8

7.5

-

-

5300 7540 2940 3 . 9 x 104 6 . 8 X 10' 1 . 6 x 107

21

3.9

4.9"

1.7

-

-

25", Z = 0.2 (496). b25", 0.1 M NaCl 0.1 M tris-acetate (499). c Better fit to pH dependence if, in addition] an acid path is assumed to operate in parallel with turnover number kr = 0.19 sec-1.

+

24.

775


STE.4DY STATE

Substrate Cp benayl Cp methyl CPA CpdA CPG Cp purine-9-riboside CpmsA Cpm2'A Cp3isoA CPC CPU CpmW CPT CPX" UPA UpO'A UPG UPC upo*c UpmaC UPU

TABLE XXXA KINETICPARAMETERS-STEP

Solvent'

ka (sec-l)

1 1 1 4 4 1 4 4 4 4 4 1 4

3 0.5 3000 2350 2350

500 220 600 90 40 400 240 160 27 18 15 3-60 1200 1000 14 69 40 26 2 20 11

11 4

4 4 4 1 4 4 4 1

4 4 4 11

4

1 Ka (d) 2 2 (assumed) 1.0 1.4

-

3.0 1.4 5.0 1.5 5.0 5.0 4.0 3.3 3.7

-

1.1 1.3-1.6 1.9 1.3 1.4 2.0 3.0 1.7

-

3.7

Ribose of second nucleotide replaced by alkyl chain, propyl through hexyl attached to N-9 of adenine as the base. Solvent 1, 0.1 M imidazole - HCl NaCl to give I = 0.2MI pH 7.0,27" (603). Solvent 2, 0.1 M dimethylglutaric acid - NaOH NaCl to give Z = 0.2 M, pH 7.0, 20" (604, 606). Solvent 3, 0.1M acetic acid - NaOH NaCl to give Z = 0.2 M, pH 5.8, 27" (6%

+

+

+

119).

Solvent 4, same as solvent 2 except 25" (606~).

I n Fig. 26c it is seen that the curves from this simple mechanism do not fit well in the acid region. If i t is assumed that additional paths are possible, A as well as N in Eq. (2), then a much better fit is obtained. The various constants derived from the data are listed in Table XXIX. 500. B. D.McLennan and B. G. Lane, Can. J. Biochem. 46, 93 (1968). 501. J. T.Bahr, R. E. Cathou, and G. G. Hammes, JBC 240, 3372 (1966). 502. P.W. Wigler, JBC 243, 3466 (1968). 503. H. Witzel and E. A. Barnard, BBRC 7, 289 (1962). 504. H.G. Gassen and H. Witzel, European J. Biochem. 1, 36 (1967).

776


TABLE XXXB

STEADY STATEKINETICPARAMETERS-STEP 2 Substrate

ac4C > p m4C > p m2C > p U>P m*U > p H > P ClSU > p Br6U > p ISU > p s4u > p mB4u > p *>P ml* > p mW > p mit% > p N-3-Uric > pb N-9-Uric > p” 8-BrG > p SCHISG > p 8-OxyG > p XXIXd

xxx.

XXXIl APC > P a

PH

Solvent0

7.0 7.0 5.8 7.0 5.8 7.0 7.0 7.0 7.0 5.8 7.0 7.0 7.0 7.0 7.0 7.0 7.0 7.0 7.0 7.0 7.0 7.0 7.0 7.0 7.0 7.0 7.0 7.0 7.0 7.0

k. (sec-1)

5.5 5.5 2.0 0.5 0.2 1.9 0.2 2.2 1.4 1.0 No inhibition 0.5 1.7 1.3 1.2 5.2 1.9 0.3 1.6

1 2 3 11 2

3 2 2 1 2 3 2 11 2

2 2 2 2 2 2 2 2 2 2 2 2 2 2 2 2 2 4

K* (d) 3.0 3.3 0.4 5.5 0.4 5.6 2.5 5.0 3.6 0.5 5.5 2.4 1.6 1.4 1.9 3.3 3.6 1.8

-

-

No inhibition 0.5 0.1

1.2 1.0

-

-

1.3 0.2

0.8 1.5

-

-

0.2 11

27.8 0.9

See footnote a, Table XXXA, for solvent designation.

* N-3-Uric acid riboside-2‘: 3’-phosphate.

N-%Uric acid riboside-2‘: 3’-phosphate. 1(j3-~-Ribofuranosyl)-2-pyridone-2’: 3’-phosphate. 1(~-~-Ribofuranosyl)-3-methyl-2-pyridone-2’ :3’-phosphate. 1(j3-~-Ribofuranosyl)-3,6-dioxypyridazine-2’ :3’-phosphate.

Table XXX contains some of the earlier data of Witzel. The marked rate difference between Py-Pu and Py-Py bonds is confirmed in partial digests of high polymers (600). The equilibria for the hydrolysis of the pyrimidine cyclic phosphates lie far in the direction of hydrolysis but have been measured. The 505. H. Witzel and E. A. Barnard,

BBRC 7, 295 (1962).

24.

BOVINE PANCREATIC

777

RIBONUCLEASE

TABLE XXXI CONSTANT-STEP

INHIBITION

Inhibitor 3'-UMP 3'-CMP

2'-CMP

2'(3')-HMP 5-F-Uridine3'-sulfate

NO*-

PH

Solvent

2 Ki

(pM)

5.5 7.0 6.1 7.0 8.1 4.0 5.0 6.6 7.0 7.0 7.5 7.3 5.6 5.8

50 0.1 M NaCl, 0.1 M tris-acetate, 25" 0.1 M NaCl, 0.1 M tris-acetate, 25" 450 I = 0.01 imidazole chloride, 25" 4 I = 0.01 imidazole chloride, 25" 57 Z = 0.01 imidazole chloride, 25" 430 I = 0.2, 25' 1000 I = 0.2, 25" 80 I = 0.2, 25" 365 I = 0.2, 25' 654 I = 0.2, 25" 148 I = 0.2, 25" 717 I = 0.1, tris, 30" 280 20" 1 0.1 M acetate, 25" 720 390 7.0 0.01 imidazole-HClJ25" 0.1 M

5.5

Ref. (499) (499) (602) (609) (60.9) (496) (4.96) (496) (4.96)

(496) (496) (467) (468) (460)

(606) (466)

hydrolysis of C > p to give 3'-CMP has an equilibrium constant of 360 120 at 25", pH 5, 0.1 M KNO, (501).The equivalent value for U > p e 3 ' - U M P is 440 110 a t 25", pH 5 , 0.1 M NaCl, 0.1 M trisacetate (499).With these values, the other kinetic parameters and the Haldane equation the turnover number for the reverse reactions can be calculated. Some values have been reported by del Rosario and Hammes (499) for 3'-UMP and by Wigler for 3'-CMP (502); see also Bahr et al. (501).Inhibition constants for these products and other related compounds are given in Table XXXI (455,457,458, &O, @6, 499,602, 606).

*

*

2. Influence of Ionic Strength With RNA as a substrate and using an assay based on release of acid soluble nucleotides, Kalnitsky et al. (607) showed the interdependence of pH and ionic strength on the measured activity. At pH 7.3 the optimum ionic strength (largely KCl) was about 0.07 M while a t pH 5.2 the value was 0.2 M . At constant ionic strength the pH optimum 505a. H. Follman, H. J. Wieker, and H. Witrel, European J. Biochem. 1, 243 (1967). 506. P. W. Wigler and H. U. Choi, JACS 86, 1636 (1964). 507. G. Kalnitsky, J. P. Hummel, and C. Dierks, JBC 234, 1512 (1959).

778


shifts from 7.5 a t I = 0.1 to 6.0 a t I = 1.0 (508).Dickman and Ring (509) found comparable effects and also showed the much higher ionic strength, about 0.35 M , for maximum activity in the C > p hydrolysis. The inhibitory effect of salts in the range 0.5-1.0 M appears to be correlated much more with the nature of the anion than with the cation, nitrate being a stronger inhibitor than fluoride and F > C1, I > Br. Although Irie (508) treated the effects as simple competitive inhibition. the general picture is much more complicated. A summary of the types of behavior found is given in Figs. 27a and b (510).The “zero” salt values actually refer to activities measured in the 0.1 M buffer included in all the solutions. The apparent activation seen a t high ionic strength depends on the salt and the substrate. Note that action on RNA effectively ceases above 2 M salt while activity toward C > p is enhanced. ’

r---

1

I

,

I

I 1 2 ‘ 3 / I

I

/

-0

I

3

2

I

I

4

Salt concentration (moles/liter)

I

I

1

I

2

3

Ammonium sulfate conc. (moles/liter)

(0)

(b)

>

FIG.27. Effects of specific salts and of ionic strength on the C p activity of RNase. (a) The activity is plotted as that observed relative to the value a t low ionic strength (0.1 M).All C p assays were carried out at pH 7. Insert: Substrates, (0-0) RNA (pH 5.0); ).-.( C p, salt, (NH4)&04. (b) Effect of ammonium sulfate on the activity of various enzyme preparations: (0) RNase-A, (0) RNase-A (dimer), ( A ) RNaseS‘, (A)S-protein incubated with substrate and then activated with S-peptide. [Reproduced from Winstead and Wold (610).1

>

>

508. M. hie, J . Biochem. (Tokyo) 57, 355 (1965). 509. S. R. Dickman and B. Ring, JBC 231, 741 (1958). 510. J. A. Winstead and F. Wold, JBC 240, 3694 (1965).

24.


779

3. Influence of Organic Solvents

Findlay et al. (611615) have reported extensive studies on the influence of organic solvents on RNase activity. Primary alcohols serve both as general components of the solvent and as substrates in the alcoholysis of cyclic phosphates. Dioxane and formamide are not potential substrates. Solvent polarity is expected to have a much larger effect on the ionization of a neutral buffer where charge separation is involved than on a cationic buffer where charge separation is not involved. Thus, the change in the pH profile of activity on the addition of a solvent such as dioxane in the presence of various buffers can be used to infer the charge type of the ionizing groups a t the active site. The activity is plotted against the pH of the solution before addition of the dioxane. If the induced change in pH resulting from the change in buffer pK is the same as the induced change in the pK of the protein group then no change in protein ionization will occur. Since this cancellation of effects occurred when cationic buffers were used the authors concluded (512) that both groups in the enzyme, assumed to explain the bellshaped curve, were cationic acids, i.e., histidine or lysine. This accords well with all the other evidence pointing to the involvement of two histidine residues. The differential influence of inert solvents on the rates of hydrolysis and alcoholysis of C > p led Findlay et al. (613) to postulate a water or alcohol binding site on the enzyme. Hydrolysis appears to be competitively inhibited by alcohols. A large portion of the effect can however be attributed to reduced water activity, the acidity of the alcohols being greater than the acidity of water and to the variability of acidity among the various alcohols. Although the data are interpreted as noncompetitive inhibition, a t least in the case of glycerol, classic competitive inhibition curves seem to fit the given data equally well. At a fixed water-alcohol ratio the relative rates of hydrolysis and alcoholysis are not affected by pH. Esters are formed by primary alcohols except for benzyl alcohol but not by secondary alcohols. Hydrolysis of C > p is favored over methanolysis in dioxane but the reverse is true in formamide. In the step l reaction of the methyl ester of cytidine-3’-phosphate, dioxane promotes the formation of C > p relative to the effect of formamide. These observations were used as support for the concept of a water (methanol) binding site. The conclusions are based on the absence of any conformational effects induced by the solvents. The lack 511. D. Findlay, A. P. Mathias, and B. R. Rabin, Nature 187, 601 (1960). 512. D. Findlay, A. P. Mathias, and B. R. Rabin, Biochem. J . 85, 139 (1962). 513. D. Findlay, A. P. Mathias, and B. R. Rabin, Biochem. J . 85, 134 (1962).

780


of activity of RNase-S in 40% dioxane indicates a loosening of structure. This probably also occurs in RNase-A, and the effects are not easily predictable.

F. MECHANISM OF CATALYSIS There are few subjects which generate as much heat as the intimate details of organic reaction mechanisms. The mechanism of ribonuclease action is no exception. 1. Mathias and Rabin et d.

The first detailed proposal for the mechanism of action of ribonuclease was put forward by Mathias and Rabin and their colleagues (614). An original diagram from their paper is shown in Fig. 28 (616, 616). It bears a remarkable similarity to the geometry of the active site as defined by the X-ray studies and shown in Fig. 23. For Step 1 the mechanism proposes (1) removal of the proton on the 2'-OH by an imidazole residue in the base form, (2) protonation of the 5' 0 of the leaving nucleoside by the other imidazole in the acid form, and (3) attack by the 2' alkoxide on the phosphorus atom to yield the cyclic phosphate. Hydrolysis or alcoholysis of the cyclic phosphate requires the reverse of each of these steps. At the start of step 1, one histidine is in the acidic form and one in the basic form. At the start of step 2 the roles of the two histidine residues are reversed. It is now thought that the two histidine residues are almost certainly His 12 and 119. The pH dependence of the reaction rate implies two pK values of about 5.4 and 6.4 (25",I = 0.2). The rate of loss of enzymic activity as a function of pH in the presence of iodoacetate also follows a bell-shaped curve and is known to involve both His 12 and 119 (see Section IV,B,5,b). However, the derived p K values are 4.8 and 5.5 (40", I = 0.02) (617).The shift in pK values is compatible with the ionic strength effects on the His pK values as reported by Witzel (see Table XIV) . The original mechanism was modified later to include Lys 41 and a water molecule hydrogen bonded to both C2 carbonyl oxygen of the 514. D. Findlay, D. G . Herries, A. P. Mathias, B. R. Rabin, and C. A. h a s , Nature 190, 781 (1961). 515. D. Findlay, D. G. Herries, A. P. Mathias, B. R. Rabin, and C . A. Ross, Biochem. J . 85, 152 (1962). 516. A. Deavin, A. P. Mathias, and B. R. Rabin, Bwchem. J . 101, 14c (1966). 517. M. P. Lamden, A. P. Mathias, and B. R. Rabin, BBRC 8, 209 (1962).

24.

781


I I-\

H

+

'

NH,

0

n FIG. 28. Mechanism of action of RNase according to (a) Findlay et al. (616) and (b) Deavin et al. (616).

pyrimidine and the N H of the His residue in the acid form Fig. 28b (616). The latter was an attempt to provide an alternate role for 02 as compared to that proposed by Witzel. 2. Witzel

One of the keys to the Witzel mechanism is the keto oxygen a t the 2 position on the pyrimidine (618, 619). He assumed that the only interaction with the enzyme occurs through the phosphate group and at 518. H. Witzel, Ann. Chem. 635, 191 (19aO). 519. H. Witzel, Progr. Nucleic Acid Ree. 2, 221 (1963)

782


that only with the dianionic form of the pentacoordinate intermediate as in Fig. 29. The enzyme alters the electrophilic character of the phosphorus atom by single or double protonation in the transition state but does not affect the nucleophilic character of the 2’ OH of the sugar. The latter is changed through the interaction of the 2’ OH and the keto oxygen, 02. The differing rates of the various substrates are then related to the electronic character of this particular oxygen atom. The polarizability of the base is suggested as the most relevant parameter but basicity and positioning are also important. For step 1, the large effects of the second ester group are related to a “preordering” of the relevant atoms through base stacking from the rough correlation with hypochromism in the dinucleoside phosphates. Again, any influence of the enzyme on binding the base is ruled out. The positive sites on the enzyme are assumed to be His 119 and Lys 41. Histidine 12 is assumed to interact with His 119 through a shared proton in order to maintain the proper geometry of the free enzyme and the proper electrophilicity of the proton. The bell-shaped pHactivity curve is still the result of the titration of His 119 and 12, but neither of these residues is assumed to have the acid-base role assigned by Mathias and Rabin. 3. Wang

The general concept of facilitated proton transfer can be applied to RNase (520) (see Fig. 30). A basic imidazole group removes the proton 0

0

(VI)

0

(V)

(IV)

FIG.29. Mechanism of action of RNase according to Gassen and Witzel (604). 520. J. H. Wang; Science 161, 328 (1968).

24.

783


I

I

(b

f0

‘

/N+

f0

3’

k

II

P

I

C0

H-N

(W

+

“1

Fro. 30. Mechanism of action of RNase wcording to Wang (620).

from the 2’ OH group. This same proton is then donated to the 5’ oxygen of the leaving nucleotide a t the end of the cyclization (step 1 ) . For alcoholysis or hydrolysis, step 2, the path is merely reversed. If this process is to occur efficiently no significant movement of any of the atoms can be involved. Thus the 2’ 0 atom and the 5’ 0 atom must both be about the same distance from the imidazole nitrogen prior to, during, and immediately following the formation of the pentacovalent intermediate. This requirement forces the 5‘ 0 t o be equatorial in the initial intermediate, and pseudorotation is a mandatory part of the mechanism. Protonation of a free equatorial oxygen facilitates attack and deprotonation. The other histidine or Lys 41 is involved in this function. A similar mechanism has been proposed by Hammes (4.66). 4. Usher

Usher (621) has classified all of the proposed mechanisms by the geometry of the displacement. The possibilities are the “in-line” and “adjacent” processes. The latter, which he favors for step 2 (622) require pseudorotation of an intermediate. The significance of the geom521. D. A. Usher, Proc. Natl. Acad. Sci. U. S. 62, 661 (1969). 522. M. R. Harris, D. A. Usher, H. P. Albrecht, G . H. Jones, and J. G . Moffatt, Proc. Natl. Acad. Sci. U. S. 63, 246 (1969).

784


W.

WYCKOFF

etry is based on the study of model compounds by Westheimer and his associates (see references in 691) and is discussed below and embodied in Fig. 31. The mechanisms of Witzel and Wang and Hammes require pseudorotation. The mechanism of Mathias and Rabin might or might not depending on the disposition of the acid and base groups. Eckstein (443) concluded on the basis of the phosphorothioate studies that pseudorotation does not occur in the enzyme-catalyzed step 2. The decision was based on the lack of sulfur exchange, but this does not rule out pseudorotation if the exchange processes are prevented or severely inhibited by binding to various groups on the enzyme or if mediated protonation of the leaving group is involved. The paths which are ruled out are indicated in Fig. 31. 5 . Roberts et al. From the geometry of the 3’-CMP complex derived from N M R and X-ray data and assuming that the dinucleotide substrate complex would be similar, Roberts et al. (523) concluded that a linear mechanism similar to that implied by the original Mathias and Rabin proposal is in fact correct for step 1. The two possible routes are shown in Fig. 31.

G. DISCUSSION OF THE MECHANISM AND SPECIFICITY Numerous closely related mechanisms for ribonuclease activity have been proposed based on an ever-growing body of knowledge concerning chemical and genetic or synthetic replacement modifications of various groups on the protein, preparation of S-peptide and S-protein, pH dependence of and comparisons between steady state (ks,K,, Ki, and stimulation) and dynamic kinetic parameters (reaction k, and isomerizations) of natural substrates and products and analogs, general organophosphorous chemistry, NMR chemical shifts of individual histidines, and the three-dimensional structure of the enzyme and ligand complexes with the enzyme as revealed by X-ray diffraction. The various proposals have each evolved with time, and the issues are numerous and intertwined. There is general agreement that the phosphorus is attacked by an oxygen atom made more nucleophilic catalytically. A catalytically stabilized pentacovalent intermediate (or activated state) is accepted. Specific catalytic protonation of the leaving group is involved. There is also agreement that a t least two of the three residues His 12, His 119, and Lys 41 are involved crucially in the mechanism. 523. G. C. K. Roberts, E. A. Dennis, D. H. Meadows, J. S. Cohen, and 0. Jardetzky, Proc. Natl. Acad. Sci. U.S. 62, 1151 (1969).

24.

BOVINE PANCREATlC RIBONUCLEASE

785

1. Structure

Our knowledge of the structures of substrate related compounds complexed with RNase-S in 3 M NH,SO,, 0.1 M AcO, at pH 5.5 as deduced from X-ray data (61, 62, 71,72b, 120,137, 469, 624, 626, 626) is summarized in Figs. 6a, 20, 23, and 32 (see also Figs. 2 and 4). Figure 20 is a diagrammatic representation of many of the findings, and Fig. 23 shows a detailed drawing of one dinucleotide complex. In Fig. 20, B,, R,, B,, and R, are the positions of the base and ribose components of the dinucleotide or independent pyrimidine and purine nucleotides, respectively. The phosphate position p, can be occupied by the 3‘,5“-diester (5” refers to the 5‘ position of R, in a diester) or the 3‘- and 5’-nucleotides, respectively. In the protein crystal a sulfate ion occupied this position in variable degree depending on the pH. Histidine 119 can be in any one of four or more positions depending on various factors. The second base might be in position B,’ when it is a pyrimidine. The phosphate of a cyclic substrate or pentacovalent intermediate may be a t p,’. The position labeled H 2 0 is the position of an isolated peak on the electron density map which is interpreted to be a water molecule, W,, present in the protein and in the complexes. The site B, has been established by three-dimensional electron density maps for UpcA, 3’-CMP, 3’-UMP, 2’-CMP, d-TpT, and 4-thio uridine. The site B, has similarly been demonstrated for the adenine in UpcA (626u),3’-AMP, B’-AMP, 3‘:5‘ A > p, and ATP. The two sites can be occupied simultaneously or independently by 3‘-AMP and 3’-CMP with no strong interaction. The B, site has also been demonstrated for 3’-CMP and 3’-UMP in the DNP-Lys 41 derivative. Electron density has appeared at the B,’ site in high concentrations of pyrimidines and in the inferred digestion products of UpU in the DNP-Lys 41 crystals. The interpretation of density a t p, is based on crystallographic “titration” of the sulfate in the protein crystal, the absence of a phosphate peak in difference maps for the mono- and dinucleotide, on arsenate and pyrophosphate binding, and by analogy with RNase-A where phosphate and arsenate peaks have been reported near His 119. The X-ray data show that His 119 is not in any one clear-cut position 524. H. W. Wyckoff, K. D. Hardman, N. M. Allewell, T. Inagami, D. Tsernoglou, L. N. Johnson, and F. M. Richards, JBC 242, 3749 (1967). 525. H. W. Wyckoff, F. M. Richards, M. Doscher, D. Tsernoglou, T. Inagami, L. N. Johnson, K. D. Hardman, and N. M. Allewell, 7 t h Intern. Congr. Bwchem. Abstr., Tokyo, Aug. 19-26 (1967). 526. W. Carlson, H. C. Freeman, A. W. Hansen, H. A. Heitsman, J. R. Knox, Y. Mitsui, D. Tsernoglou, and H. W. Wyckoff, unpublished data. 526a. UpcA is the phosphonate analog of UpA with a C& group replacing 05”.

786

F. M. RICHARDS AND H. W. WYCKOFF STEP 1

Opposite (05“)

Adjacent (05”)

(in-line)

(opposite YH)

(opposite XH)

C5’ \ 05” X

C5”

ox yo c5;

I/

HY-P

05#-P-02’

I

L2l 0 3\c& ’ \

‘

14 -02 ’ I \cz*

TC& \ \

\

+H

V

$.

C5”-05”-H Y X@\ ‘P-02)

’

C5”-05” \ x-P-02’

X

+

YO\

Y’

‘P-02’

‘R‘

OYCGL2’ \ \

\

Fm. 31. Possible pathways for RNase action. The “in-line” mechanism is -sored for step 1 by Roberts et al. (623). The “adjacent” mechanism requiring the intermediate step of pseudorotation is suggested by Usher (629) for step 2, the hydrolysis of the cyclic phosphate.

in the plain protein crystal. In the presence of bound adenine nucleotides in site B, definitive electron density appears in site IV and leaves site 111. I n the 3’-CMP complex the density in site I11 increases slightly showing that this position is somewhat stabilized. Position I1 contains electron density, but it may be a bound solvent molecule. Position I completely overlaps the sulfate site (pl), and in the absence of sulfate it could be a prominent site a t some pH values. At p H 8 there is no

24.

787


STEP 2

1

t

(-H) + H

0,

yo

Z(H) Yo

\/

HX-P-02‘

@X

Z(H)

I/

HY-P-02‘

\

Opposite ( 0 2 ’ ) (in line)

FIQ.31 (Continued).

electron density a t p,. In position I His 119 would contact His 12 and could hydrogen bond to it. I n 3’-CMP the C3’-03’-P03 moiety is twisted away from Lys 41 in such a way that the PO, rests between His 12 and 119, above the backbone N H of Phe 120, and next to the solvent molecule W,. If the hydrogen bonding of N1 of His 12 to the carbonyl group of Thr 45 is correct, then N3 is next to the phosphate. The amide side chain of Gln 11 is next to W, and probably hydrogen bonded to it. In position 111, His 119 is highly exposed to the solvent. 02’ of 3’-CMP is twisted toward Asn 44 and may hydrogen bond to the amide oxygen if the amide

788


NH, hydrogen bond to the backbone oxygen of Gln 11 is correct or to the NH, if the assignment is erroneous. The 2 oxygen of the base in the site B, accepts a bent hydrogen bond from the backbone N H of Thr 45. This oxygen is close to His 12, and if the atom assignments for the latter are correct 0 2 is in contact with C2-H. 0 2 appears relatively inaccessible to solvent although a space filling drawing or accessibility calculation has not been made yet. 0 2 is on the opposite side of C2‘ from 03’ in 3’-CMP and the 05” equivalent in UpcA. The B, ring partially overlaps the ring of Phe 120. The OH of Thr 45 provides and ambivalent acceptor or donor to hydrogen bond to N3; Asp 83 is distal to N 3 potentially bonding to Thr 45 in the free protein and in some complexes. I n the dinucleotide UpcA the B,, R,, p,, and W, situation is very similar to 3’-CMP, but the B,R, site may be shifted outward somewhat with 02’ somewhat further from Asn 44 and closer to P. The adenine ring in B, stacks against His 119 in position IV and Asp 120 may bond to the N3 of His 119. Glutamic acid 111 and Asn 71 may bond to the adenine. The B, site is not necessarily exclusively specific for pyrimidines, and specifically 3’-AMP and 3‘:5’ A > p appear to bind weakly in this position although the interpretation is not secure. I n the absence of sulfate 3’-AMP might prefer the B, site and Witzel (private communication) has evidence to this effect based on UV spectroscopy and enzyme kinetics. Lysine 41 is not in contact with the phosphate or sulfate in the p, site. It is nicely constrained by the structure so that i t cannot be in contact without a gross fluctuation (backbone motion) from the average structure derived in the X-ray analysis. Cyclization of 3’-CMP would require appreciable rotation around the C2’-C3’ bond, around the C3’-03’ bond, and around the 03’-P bond. The phosphorus must move approximately 2 A relative to the 02’-C2’-C3’-03’ configuration, and if B, and R, do not move then the PO, would move directly toward Lys 41 and potentially make contact. Figure 32 is a diagram of the distance of each a-carbon and the tip (T) of each side chain (see Section II1,B) from p,. Histidine 119 is in position IV and N1 of this residue would be much closer than T defined as halfway between C2 and N3. Lysine 41 is indicated a t two distances, one as it seems to be in DNP-Lys 41 and the other in its closest approach to p,. 2. Comparison of Transphosphorylation ( 1 ) and Hydrolysis ( 2 ) Most of the discussions have assumed that steps 1 and 2 are the reverse of each other with one of the prime reasons being simplicity. Usher et at.

24.


789

FIO.32. Diagram of the distance of each a-carbon and the tip of each side chain from the phosphate position, P. The angular position on the diagram has no meaning. The radial positions are calculated from coordinate set 6. (622) and Ward et al. (4@) have specifically noted that the two steps do not necessarily follow the same pathway. These provisos were based on general grounds, on the grounds that the small molecule chemistry did not rule out the possibility, on the basis of conflicting views with regard to the mechanism, and on specific results with formycin. The X-ray data do not yet, and cannot conclusively, provide an answer to this question. The current interpretation does distinctly allow for the possibility that transphosphorylation occurs by an opposite attack (see Section VI,G,3) and that hydrolysis occurs by an adjacent attack. The dinucleotide phosphonate as depicted in Fig. 23 seems poised for an while W, is in good position to make an attack of 02’ opposite C5”-CH2

790


attack on the product adjacent to 02’. Considerable motion occurs on cyclization as noted above and therefore the latter statement is admittedly conjectural. An additional set of data lends some plausible support to the suggestion that RO- (in the reverse of step 1) and OH- attack different faces of the cyclic phosphate. The presence of adenosine, which is a good acceptor in the synthesis of dinucleotides from C > p (S91), also stimulates hydrolysis of C > p as shown by Wieker and Witzel (527) instead of competing for the phosphorus as might be expected if the mechanisms were the same in detail. The pH dependence of K,/K, is similar for step 1 and step 2 reactions as shown in Fig. 26b, but this similarity in the p H curves indicate only that the same titratable groups on the free enzyme and/or free substrate are involved in the two steps. As discussed explicitly by Usher et al. (522) the roles of the two histidines could be reversed and this would make no difference since the ratio of HE:EH where these are the two singly protonated species is independent of pH. Similar k, and K , curves for the two steps would also fail to prove identical roles for the two histidines. Since a pentacovalent species-whether it is a transient activated complex or a more stable intermediate-is common to the various alternatives, pK shifts deduced from k, curves could be the same. Both substrates are monovalent anions with low pK values so that l/Km, whether interpreted as an equilibrium binding value or as a function of the kinetic parameters mirroring the total occupancy of all the stable intermediates, could also be the same for both steps. The values for the reverse of step 2 would behave differently since the pK of 3’-CMP, for example, is 5.9. It should also be noted that k,/K, curves should be and are ionic strength dependent (508) in the same way that the His 12 and His 119 pK values are as observed by NMR (280). Ward et al. (442) contended that digestion of poly formicin to F > p while F > p is not hydrolyzed shows that the two steps are somehow different. The difference could be in the substrates rather than the mechanism since they also reported the F > p was not an inhibitor. Even if it were an inhibitor nonproductive binding could explain the results invoking the rigidity of the cyclic phosphate ring in combination with the peculiarities of the base F. Bernfield’s (390) observation that S-protein can very slowly catalyze synthesis from C > p as compared to little observable catalysis of the hydrolysis does suggest that a minor pathway for step 1 not involving His 12 may exist. The digestion of poly A or ApApApA but little or no digestion of ApA or A > p (435, 519) may 527. H. J. Wieker and H. Witzel, European J . Biochem. 1, 251 (1967).

24.


791

be related. Specific differences in relative activities do not imply that mechanisms are different since specific base binding could account for the activity changes. Since 0 2 is sterically prevented from attacking opposite 03’ and can attack opposite 05’’ according to the preference rules given in the next section, an in-line mechanism is a simple proposal for step 1. On the other hand, in the hydrolysis the activated OH- attack must occur opposite either 02’ or 03’ and therefore must also be adjacent to one or the other as illustrated in Fig. 31. Pseudorotation allows such an attack to produce 3’-CMP and Usher’s cyclic phosphonate experiments lend support to this hypothesis. Since the two steps might be different with respect to pseudorotation and/or histidine involvement, each point must be proved separately for each step. One must also ask whether a given reaction such as methsnolysis of cyclic phosphates is similar to step 2 or the reverse of step 1. Is the “acid pathway” the same as the neutral pathway with respect to pseudorotation? Are the pathways for CpA and CpU the same? Usher et al. (622) have suggested a direct way of proving the point of attack based on reactions involving the resolved stereoisomers of cyclic phosphorothioates introduced by Eckstein (4.43). This must be applied to each reaction. In the case of methanolysis the product is stereospecific depending on the point of attack. Hydrolysis with I8O followed by neutron diffraction identification of the produce might be needed to test step 2. Step 1 involving a dinucleotide might be checked by using the S-protein to synthesize a stereospecific dinucleotide from the resolved cyclic phosphorothioate as Bernfield (390) used it to synthesize normal dinucleotides. 3. Opposite us. Adjacent Attack

The question of opposite vs. adjacent attack and the need for pseudorotation in the case of the latter but not the former has been brought to the forefront by Usher (621) and Dennis et al. (693)who both worked with Westheimer on the organic chemistry of phosphorus. The various possibilities relevant to the current discussion are illustrated in Fig. 31. A set of statements and preference rules summarizing the findings are as follows: (a) A nucleophilic attack on the tetrahedral phosphorus (bond angles 109’) produces a pentacovslent intermediate consisting of a trigonal biprism with the three basal atoms at 120” to each other and a t 90” to the apical atoms. (b) The attacking group enters a face of the tetrahedron assuming an apical position and forcing the three adjacent atoms to become basal

792


(equatorial) while the opposite atom becomes apical remaining in line with the entering group. (c) A leaving group leaves from an apical position. (d) The intermediate can rearrange with a minimum of distortion by increasing the angle between the two atoms in the basal plane displacing them 30” toward the third basal atom while displacing the two apical atoms 30” away from this atom to form a new basal plane which is 90” away from the original basal plane. This process is called pseudorotation. It is illustrated in the second column of Fig. 31 where the relative motions of 05’’ and 03’ are toward X while YH and 02’ move away from X. If 02’ and 03’ are held comparatively immobile, the same relative motions can be described as X moving 30” away from 02’ and 30” down toward 03‘ while YH moves forward 60” and away from X. At the same time 05 “ moved 60” toward the original X position and away from 03’. (e) Groups tend to be in the apical position with preferences in the order OR(H) > 0- > CH,. ( f ) A five-membered ring such as exists in the 2’:3’-cyclic phosphates spans an equatorial to basal position. Otherwise the ring would include a 120” angle a t the phosphorus while the COP angles also tend to be 120” and the average angle in a pentagon is 108” or less. With reference to Fig. 31 and step 1, one can thus postulate attack of 02’ opposite 05’’ or following or concomitant with protonation of X or Y the attack can be opposite one of these. Attack opposite 03’ is sterically impossible. Attack opposite 05’’ would allow immediate departure of 05’’ without pseudorotation or protonation of X or Y. Protonation or juxtaposed charges could facilitate attack by making P more positive and thus reduce the energy of the activated complex. Either would also likely stabilize the pentacovalent intermediate ES’ and might trap it for a finite time. Witzel (619) has emphasized both of these points without reference to pseudorotation. Protonation of X or Y coupled with pseudorotation could allow 03’ to leave forming the 2’,”’-diester, but such isomerization does not occur. Roberts et al. (693) emphasized that the in-line mechanism almost requires that two different groups deprotonate the 2’0H and protonate 05’ since they are on opposite sides of the basal plane. They are physically removed from each other with a negative P-0- between them to trap any shuttle. Since they observed 3’-CMP interactions with His 12 and 119 by NMR with the greater effect on His 119 and both histidines protonated and since the X-ray structure shows the phosphate between the two histidines, they believed that both histidines are directly involved. Furthermore, the interaction of His 119 is more sensitive to the specific

24.


793

dianion bound than His 12; therefore, they argued that the motion of the phosphorus during cyclixation is more likely to release the proton on His 119 than the proton on His 12 for protonation of the leaving group. The X-ray data show that His 12 is most likely to attack 02’. Thus the push-pull system of Rabin et al. (515)is favored and embellished with specific identification of the histidines. They backed this up with all of the evidence cited above and below to involve two histidines in an important role. They then essentially said that since the linear mechanism requires involvement of two groups and since two histidines are involved in the absence of any compelling arguments to the contrary, the simplest explanation is that the probable mechanism for step 1 is linear. The adjacent mechanism for step 1 would involve the 02‘ attack opposite X (or Y) and this would require protonation of X (or Y) so that it could be apical in the intermediate. Pseudorotation would then be required to allow 05” to become apical preparatory to leaving, and 03’ would also become apical. Either group could then leave and specific protonation of 05’’ would be required to explain the lack of 2’,5”-diester formation. The motion of X might facilitate its deprotonation, and conversely deprotonation would induce or facilitate motion to an equatorial position. Wang (520) specifically involves His 119 or Lys 41 in this role in an adjacent mechanism. Looking toward the P from 03‘ the clockwise sense of the other oxygens would be 02’,X,Y. If the atoms can be tagged this isomeric difference from the in-line mechanism would provide a definitive answer as suggested by Usher and discussed above. At present, the evidence in favor of the adjacent mechanism relates to step 2 only and is provided by Usher’s work on the phosphonate. Wang preferred the adjacent mechanism with His 12 acting as a shuttle by analogy with chymotrypsin and because of the nicety of the intermediate situation where the group protonating the leaving group is not trapped by bonding to an anion and is symmetrically between 02’ and 05”. Hydrogen bonding to an 0- or =O is greatly preferred to bonding to an ester oxygen. Facilitated transfer of a proton along a hydrogen bond would allow His 119 to protonate and deprotonate X (or Y ) , but using His 119 first to stabilize an intermediate and then to protonate the leaving group would be more complicated and less likely. Witzel’s mechanism (519) has much in common with these adjacent mechanisms since one group, the pyrimidine 02, is used to deprotonate the attacking group and protonate the leaving group. It also specifically invokes protonation of phosphoryl oxygens to facilitate attack opposite one of these and uses histidine in this role with a sensitive balance required between the ability to attract a proton from solution but not hold it so tightly that it cannot protonate the PO- or at

794


W.

WYCKOFF

least induce positive charge on the P. Hammes (466) preferred a shuttle mechanism for step 1 since it ends with protons in their original positions and avoids any rate problems that might ensue from switched position. Eckstein (443) has shown that the two isomers of uridine 2’:3’-0,Ocyclophosphorothioate (Le., either X or Y are sulfur) are both substrates for RNase and that no sulfur exchange takes place with the solvent. He concluded from the latter that pseudorotation cannot take place since sulfur would then become apical and be able to leave, thus producing a cyclic phosphate. Such exchange does take place in normal acid catalysis. Usher et al. (522) have countered that a requirement of specific catalytic protonation of the leaving group could prevent this exchange. This protonation would be in addition to the protonation which allowed the sulfur to become apical. The in-line mechanism for step 1 would not be ruled out by this argument since it does not involve this double protonation of the leaving atom. In summary the argument for or against in-line and adjacent (pseudorotation) mechanisms are inconclusive and may both be correct in the transphosphorylation and hydrolysis steps, respectively. 4. Stabilization of Intermediates

The NMR data in the presence of inhibitors such as 3’-CMP show His 12 and 119 to be protonated with apparent pK values of 8.0 and 7.4, respectively, in this specific case. The actual pK values may be higher since saturation was not necessarily maintained above pH 7. The implication is that the phosphate oxygens are not protonated although this was not determined. If a proton is between the histidine N and a phosphoryl oxygen it will spend part of its time on the oxygen depending on the relative basicities in this situation. The latter are not known except by analogy with the pK values, and certainly the basicities in the pentacovalent intermediate are not known. Since the jump rate in a good hydrogen bond is very rapid the question may be pedantic. Hummel and Witzel (456) showed that protons were released on binding 3’-CMP below pH 5.5 and taken up from solution above this pH as noted below. X-Ray data show the product phosphate to be in contact with both histidines although His 119 is poorly defined, partially stabilized in position 111, and probably in several positions. But where is the PO, in the intermediate or the cyclic substrate? After cyclization the PO, may contact Lys 41 and His 12 and could bond to Gln 11 and Asn 44. Histidine 119 may move to a position between I and IV and still contact the intermediate, and Asp 121 may move to contact His 119. Is this the situation in the active complex? Histidine 119 may be trapped in position I11 part of the time in an abortive complex. In the absence of phosphorus

24.


795

NMR data, one can only say that an intermediate is probably protonated part of the time and thus either an in-line or a pseudorotation mechanism is allowed. The total interaction can include other specific hydrogen bonds and the interaction with solvent is completely conjecture. There must be a balance between too much stabilization of any intermediate and too little stabilization of the activated complex following it in the pathway. Hammes and his colleagues have observed a number of relaxation times in temperature jump and stop flow experiments with various substrates and products a t various pH values. They concluded that there are a t least two detectable isomers for each substrate enzyme complex. The relaxation times between isomers is approximately sec and thus are quite fast compared to step 2 and somewhat faster than the highest overall rate for step 1, which is 2700/sec for CpA. Hammes suggested that the isomerization involves a cooperative conformational accommodation of the substrate which might involve only small and perhaps remote changes in the protein. Involvement of His 48 is suggested (468).Witzel preferred to consider this first significant complex to be the pentacovalent dianion stabilized by Lys 41 and a protonated histidine. The transition states are formed by double protonation of the phosphoryl oxygens by these two acids and protonation of the leaving group by a protonated 0 2 of the pyrimidine. The finding that deoxy CpC, for example, is not an inhibitor while 3’- (2’-deoxy) -CMP is an inhibitor indicates that the dianion is very important in binding. By analogy the postulated pentacovalent dianion intermediate would bind more strongly than the substrate. This depends on the energetics of the intermediate and cannot be settled without further information. Other authors do not attempt to assign relative lifetimes to specific complexes. 5. Proton Transfer and the Rate-Limiting Step (see Fig. 31) I n the in-line push-pull mechanisms of Rabin and Roberts, the highest energy transition state may be either the pentacovalent intermediate or the alkoxide (hydroxide) state with 02‘ or 05‘‘ deprotonated but not bonded to P. Incipient deprotonation of 02’ in an activated state is equivalent. Protonation of X or Y or nearby positive charge could stabilize the pentacovalent intermediate. Removal of either could facilitate formation of the alkoxide in the breakdown of the intermediate. Restoration of the initial state of the enzyme is required in this mechanism and could be rate limiting. I n the adjacent (pseudorotation) models of Witzel, Hammes, Usher, or Wang protonation of X or Y would be required to allow one of the two “pseudomers” to exist. In step 1 this requirement (and thus perhaps a rate limiting process) applies t o the attack by 02’. Deprotonation would force or facilitate reversal or pseudorotation to

796


W.

WYCKOFF

prepare 05” to leave. Specific protonation of the leaving group is required, and this might also be rate limiting. In step 2 the attacking OHis opposite 03‘, and protonation of X or Y is required to allow 02’ to prepare to leave. Deprotonation of the intermediate would facilitate or force departure or reversal. I n either step 1 or 2 deprotonation of the attacking group is required, protonation of the leaving group is required, protonation of the intermediate is required, and deprotonation of the intermediate is facilitative. Any of these individually or in combination can be rate limiting. It is clear how double protonation of PO, could facilitate attack, but i t is not clear how it could facilitate leaving as suggested by Witzel. It is clear how a hydrogen bond to 05” could facilitate leaving as proposed by Hammes, but it is not clear that such a hydrogen bond is probable, especially in the presence of a nearby 0-. Pseudorotation itself, following protonation, is not thought to be rate limiting but in the presence of the enzyme it could be. Since a large motion of -PO, relative to R, is required in cyclization or hydrolysis, constraint by the enzyme might make this process rate limiting. The mobility of His 119 might be related to this problem. On the other hand, the existence of three positive centers might prevent excessive stabilization and trapping of one intermediate by a kind of “rack” arrangement. One argument against the in-line mechanism is trapping of histidine protons by bonding to P-0-. It is possible that such a trap exists, but it could be symmetrical with His 12 biased toward 02’ and His 119 biased toward 05” such that protonation of P-0- by either would release the other for facilitated transfer to the leaving group. Another argument against a push-pull mechanism is the need for return to the original state as a separate operation which could be time-consuming. Motion of His 119 to contact His 12 directly, forming a hydrogen bond as proposed by Witzel, allowed by the structure, and suggested by the NMR data, could provide a quick, neat return mechanism. Insertion of a water molecule between the two histidines could allow a concerted, facilitated bond shift to affect the transfer. Thus neither the in-line nor adjacent mechanism is simple nor is it impossibly complicated. Neither provides a firm suggestion as to what the rate limiting step is or why the fastest hydrolysis [C > p stimulated by adenine a t 18/sec (527) and ApC > p a t Il/sec (505a)] should be so much slower than the fastest transphosphorylation (CpA a t 2400/sec). The fact that the pK of water is about 15.5 and the pK of ROH about 14.0 would account for a factor of about 30.

6 . Substrate Specificity and the Role of 0 2 Witzel and his colleagues are the main proponents of the direct involvement of the pyrimidine 0 2 in the proton transfer. Several aspects

24.


797

of their mechanism have been discussed above. The question of 0 2 involvement can only be discussed in connection with the question of specificity. Witzel and his collaborators have amassed an impressive array of comparative kinetic data on numerous compounds as given in Table XXX. They explained each comparison on the basis of the “nucleophilicity” of 0 2 as it would apply to the required deprotonation or protonation of 02’, 0 5 ” , or bound water. The term nucleophilicity is meant to include basicity (reaction with free water) polarizability (stability of a protonated intermediate state or transient state), and positional entropy of the functional activated state, i.e., the effectiveness of 0 2 in its catalytic role. Others explained the same facts in terms of subtle or gross steric effects on binding and on presumed critical relationships that allow productive or nonproductive binding. The arguments against involvement of 0 2 as a base are that (a) it is a weak base, (b) His 12 is available and a much better base, (c) the proposed induced changes in nucleophilicity are not obviously correct or sufficient, (d) the activated water in the hydrolysis step should attack opposite 02’ or 03’ and this access is not obvious, (e) substrates are known in which there is no 02, and (f) other explanations of the facts seem more plausible to the critic. The X-ray structure does show 0 2 to be near 02’ in product binding, but His 12 is between 02’ and the potential 05” sites. An adenine site has been established distal to 0 2 across the confluence of R,,pl, His 12, and the His 119-Phe 120 backbone. If this site is the active site in CpA digestion, it is nearly impossible to conceive of a fluctuation that would allow 0 2 to be involved in protonating 05”. Even if the hydrolysis step is the adjacent mechanism, it is difficult to see how 0 2 could compete with His 12 for a water molecule in position to attack opposite 03’ in a reasonable position for the cyclic phosphate. X-Ray data cannot be obtained from substrate bound to active enzyme since digestion takes place in the crystal and diffusion is relatively slow. Therefore absolute statements cannot be made. If 0 2 does not protonate the leaving group can it still be involved directly in deprotonating 02’? This is even more difficult to answer from the X-ray data. 0 2 is in contact with His 12 and may affect its orientation or electronic configuration and thus indirectly affect the action of His 12. The activity data relevant to the 0 2 question can be considered in several classes. First, it can be stated that all of the good substrates have 0 2 or a substitute nucleophile S or N in position on a base such that it can roughly occupy the proper position. I n addition to the normal substrate bases U and C this includes 2-thio U, pseudo U, 8-oxy purines, and formicin in poly F. The one exception is poly A and oligomers of A which are cleaved very slowly but definitely. Witzel agreed that this is real and thus that 0 2 is not absolutely essential. H e argued that the 2’ OH could

798

F. M. RICHARDS AND H .

W.

WYCKOFF

slowly attack the phosphorus without specific aid in deprotonation if the phosphorus were forced on it for long enough periods. Why is A > p or ApA not attacked or poly A not digested more rapidly if 0 2 is not directly involved? The counterargument has to be that A > p binds very weakly in the B, site (with A in the syn position) as evidenced by lack of inhibition. Furthermore, any binding could be abortive. A > p does bind in the B, site and stimulates C > p hydrolysis (527).ApA also stimulates C > p hydrolysis. 3’-AMP does inhibit a t low salt concentrations. If the 0 2 is so important in binding, why is this so? If the poor H bond to 0 2 proposed from the X-ray data is not so important, why is there not greater activity? Synthesis of polynucleotides and depolymeriaation by S-protein without His 12 is a counterpart to this argument since the same considerations then apply to His 12 whichever way one looks a t it. Poly F digestion can also be used to support either argument. If positioning is very critical, why does the placement of the N of the five-membered ring 1A away from the normal 0 2 position not prevent activity? If this N is acting as a nucleophile in place of 02, why does its position further from 02’ and 05’’ not prevent activity? The S of 2-thio U can accept a H bond, but being larger than 0 the bond is longer. Again displacement or electronic arguments explain the small loss of activity. The second group of arguments involved N3 or its equivalent. Protonation as in U or in C a t low pH is acceptable. Methylation blocks activity completely. Sterically interposing a group with a radius of 2 A, 1.4 A from N3 would be expected to lead to a nonproductive complex if any, since N3 or 3 N H are normally H bonded to Thr 45 ( a t an N-0 distance of 2.9A) in the bottom of a cleft. Electronically such a substitution would affect the mesomeric system that contains 02. Replacement of N3 with CH in 2-pyridone (504) does not destroy activity. I n fact, k , = 0.2 sec-I and K,,, decreases to 1.5 mM. The steric argument would be that the blockage is much less severe than N-CH, (by about 1.2A) and must be tolerable. The loss of the H bond is not so severe since the hydrophobic interactions would replace it. The overall kinetic constants for U > p under set conditions are k , = 1.4 sec-’, K,,, = 2.6 mM. Replacement of 0 4 with SCH, increases Km and k , only about 30% each. Substitution with S or NH, increases k , to 5.5 sec-I, while K,,, decreases 30% for S and increases 30% for NH,. Methylation of the NH, reduces k , to 1.8, while acetylation or dimethylation further reduces k , to 0.5 and 0.2, respectively. The K,,, values for the latter two are 5.5 and 2.5 mM. These examples show that k , can be changed by a factor of 25 and K m by a factor of 3 in a complex pattern by changing the hydrophobicity, the bulk, and the potential hydrogen bonding a t position 4. The resonant and potential tautomeric forms of

24.


799

the mesomeric system including 0 2 and N3 are simultaneously changed. Witzel claimed an activity correlation with properties of 02, while others dismissed the observations as too complex to explain and well within the expected variations based on nonproductive complexes and critical steric relationships. After all, catalysis by a factor of lo8 is being considered. It should be noted that 0 2 is in contact with His 12 and also that electronic interaction between the base and the phosphate is demonstrated by the hypochromicity of C > p relative to CpC or 3’-CMP. Modifications a t C5 are tolerated; for example, lc, for 5 Iodo U is 1.2 sec-’ and K , is decreased to 1.4. N-Methyl pseudo U is very similar while pseudo U has reduced turnover at 0.3 sec-’ and a K , of 3.6 mM. Position 5 is particularly exposed to solvent in the X-ray structure. Why should iodine in this position decrease K , a factor of 2? Bulky substituents in this position are also acceptable as in the 8-oxy purines or formicin. Hydrogenation of the 5,6 double bound is also acceptable. The tolerances at position 2 seem to be $0.5 A (S) to -1.0 A (poly F). The tolerance at position 3 is +0.8 A (CH) . The tolerance a t position 4 is very considerable and in position 5 essentially unlimited. Ward et al. (44.2) sharpened the focus on this situation in describing the rationale for activity against poly F. Meadows et al. (465) have presented the hydrogen bonding scheme to 2, 3, and 4 deduced from the RNase-S X-ray structure. Allewell (120) presented much of the evidence in her thesis. Richards et al. (469a) discussed the situation in a conference report. The transphosphorylation step has been demonstrated to have a much wider range of rates than the hydrolysis step, perhaps observed because hydrolysis is slow to begin with and certainly observed because the natural substrates exhibit much of this range. The reverse of this step is also observable and has even been used in synthesis of dinucleotides. The while the CpA reaction proceeds a t up to 3000 sec-l with K , 1.0 d , rate for CpU is 27 sec-’ and K , is 3.7 mM. Why are the rates so different? Witzel attributed the difference to base stacking which supposedly induces different nucleophilicity to 02 partly through direct coupling, partly through interactions with water, and partly through critical positioning of 05”. Others have invoked a separate binding site on the enzyme for Bz with A and U having somewhat different binding energies and very significantly different positions. Wieker and Witael (527) showed that binding of adenine, adenosine, 2’:3‘ A > p, and ApA all stimulate the hydrolysis of C > p with a change in both K , and K , as evidenced in parallel shifts of the Lineweaver-Burke plot. They attributed this to a ternary complex with the stimulator binding on top of the C > p in the presence of C > p and binding abortively in the absence of substrate. Rabin et al. (463) showed spectroscopically that AMP binds

800


and proposed that a positive group was nearby since the spectral shift resembled protonation. (Increasing a pK by juxtaposition of a carboxylate would serve equally well in this reviewer’s opinion.) X-Ray data have now shown an adenine nucleotide binding site as noted above. The evidence is strongly in favor of this being the active configuration or something very close to it. Secondary binding of UMP and related compounds does not occupy the adenine site, but rather the base seems to be above and to the left with little overlap with the adenine site. It is still an acceptable position for U in CpU. The electron density map for deoxy-TpT showed material in the B, and R1sites but no more, except for an intermolecular site far removed from the active center. The implication is that the second thymidine is moving about. Solubility problems and the intermolecular site limited this experiment. Histidine 119 was moved to position IV by 3’-AMP which stimulates, C > p hydrolysis (in these high sulfate conditions) and by UpcA which mimics UpA. The deduction is that this is probably the active site for His 119 in both steps 1 and 2. It comes very close to the carboxyl group of Asp 121 in this position. The implication of all of these observations regarding B, binding is that the kinetic differences result from variable positioning and configuring and dynamics of the ester linkage produced by specific interactions of the second base with the protein. The interactions are stronger with A than with U and the positions are quite different, U going part of the time to a subsite B,’ and rarely to B,. The increase in k, for C > p induced by A may result from stabilization of His 119 in the active position preventing it from getting trapped in the nonproductive positions I1 or 111. It may be more involved than this. The induced decrease in K , may likewise be this simple or more complex. Although the solvent contains 6 M NH,’, 3 M SO,2-, 0.1 M acetate, and is a t pH 5.5, stimulation by 3‘-AMP and inhibition by 5’-AMP was observed in a similar solution by Reed and Wyckoff (628). AMP inhibited CpA digestion. Thus, peculiarities of the crystal solvent cannot negate the conclusions. It remains to be seen whether all of the modifications of B, analyzed kinetically can be observed to bind in a range of subsites and whether the stimulator ApA reveals a further A site or stacking on the first A. Further X-ray observations on the more interesting of Witzel’s B, site compounds should also be attempted. In short, there are attractive alternate explanations to most, if not all, of the evidence cited by Witzel and his colleagues in support of the direct 628. S. Reed and H. W. Wyckoff, unpublished data.

24.

801


involvement of 0 2 in proton transfer. Many of the alternatives, pro and con, are hypothetical or essentially dodge the issue with vague suggestions of subtle conformational sensitivity as opposed to subtle changes in nucleophilicity. The reader is left to choose or not choose his preference.

7. L ~ 41 S Lysine 41 derivatives can have extremely low activity (if any) when properly purified (see Section IV,B,2 and Table VI) . There are, however, cases such as E-carboxyamidomethyl-Lys 41 (No. 26 in Table VI) with 3% residual activity toward C > p which raise doubts about the absolute requirement for Lys 41. One less secure observation is Marfey’s report that the cross-linked er’-DNPene-Lys 7-Lys 41 is 15% active. Goldstein reported step 1 activity for r-CM-Lys 41 and Allewell reported 1% activity for exhaustively guanidinated RNase-S. Ettinger and Hirs (371) have emphasized conformational distortion as part of the reason for loss of activity. Others generally have invoked a role for the charge on Lys 41 stabilizing a pentacovalent intermediate or transition state as discussed above. It was also suggested above that the constraint on Lys 41 and the motion of the phosphorus group during cyclization and hydrolysis might provide a more sophisticated involvement than previously proposed. It should be noted that PO4, known to bind a t His 119 and 12, inhibits reactions with Lys 41.

8. Variation of Rates and Equilibria with p H Equations (A) through (F) illustrate some of the prominent reactions proposed for the enzyme and CMP:

+ +

pH 4.5 EHz . C1 CMPH e EHz . CMP pH 5.5 EHi.4 CMPHo.6e EHz . CMP pH 7.0 E CMP 2H+ e EHz * CMP E+CMP+H+eEH.CMP

+

+ H+ + C1-

+

E E’ EHz . CMP e (EHz. CMP)’ E 2 H + e E H f H+=EH;+

+

+

(A) (B) (C)

(D)

(El (F)

The justification for (A) is manifold. NMR as interpreted by Jardetsky and his colleagues indicates that His 119 and 12 would be almost fully protonated at pH 4.5 in the protein and the complex. The pK of 3’-CMP is 5.7. Hummel and Witzel (466)showed that one proton is released when the complex is formed. Loeb and Saroff (476)detected chloride binding a t low pH. Irie showed KC1 to be a competitive inhibitor of C > p hydrolysi.~,and Riiterjans and Witzel showed that the pK values

802

F.

M . RICHARDS AND H. W. WYCKOFF

of His 12 and 119 were sensitive to ionic strength. The X-ray structure of the complex with SO, at pH 4.9 is consistent with this scheme which can be denoted His 119.H-S04.H.His 12, and the complex with 3'-CMP a t pH 5.5 is similar with the phosphate of CMP replacing the sulfate. At pH 5.5 EH2, EH, CMP, and CMPH species are present with fractional protonations of 12, 119, and CMP being approximately 0.8, 0.6, and 0.6 (pK values 6.1, 5.8, and 5.8). Since the pK values are sensitive to ionic strength, bound anions may be present in some free protein species but less than a t pH 4.5. The complex of EH1.4with CMPHo.,; would produce EH,.CMP with no release of protons as observed by Hummel and Witzel. There are several important protein species involved a t this pH and there is some controversy as to what they are and how significant. Witzel proposed that a hydrogen-bonded histidine system 119.H * 12 is important with the additional species (119-H, 12), (119, H-12), (119, 12), and (119-H, H-12) all present. The NMR results of Ruterjans and Witzel (280) support the presence of such a system and the X-ray structure permits this bonding (His 119 position I of Fig. 20). The amount of the singly protonated nonbonded species would be ionic strength dependent. The pK of the lower transition would be shifted by anion binding to the doubly protonated form. The relative apparent protonation of 12 and 119 in the bonded anion free complex would depend on the basicities of the histidines as influenced by the environment in general and the bonding to the other nitrogen in particular. The EH,.CMP complex consists of a t least two species as shown by the reaction kinetic studies of Hammes and his colleagues. Specifically they observed an isomerization of the initial complex formed on mixing the enzyme with either substrates or products and indeed an isomerization of the protein itself following a temperature-pH jump. The time constants for all of these isomerizations are similar, and a group with a p K of 5.86.0was involved in each. Since the pK values of His 12 and 119 are both > 7 as observed by NMR and are higher than 6.0 as deduced from turnover number variations with pH, they concluded that some other histidine is probably involved in the isomerizations. Histidine 48 is the logical candidate since the properties of His 105 seem quite insensitive to all variables. At pH 4.5 the kinetic constants for the first complex of UMP with protein agree with the equilibrium binding constant. At p H 7 the latter is 10 times the value predicted from the rate constant and therefore presumably 90% of the ES complex is in the second form. The E H , - CMP complex may also contain other significant isomeric forms. The X-ray data show that His 119 can be in many positions. There are two quite differentpositions both in contact with the phosphate.

24.


803

These are positions I11 and IV of Fig. 20. Position 111 seems to be preferred in the absence of adenine. It is very exposed to the solvent. In position IV, which is forced by AMP binding, His 119 is against the protein surface and potentially bonded to the carboxyl group of Asp 121. When adenine is present as AMP or in UpcA, His 119 is quite buried. Since these isomers directly involve an active histidine they may be more important than isomerization of His 48. If the shift is too rapid to have been observed it may be insignificant. If it is too slow to have been observed in the dynamic experiments the complex with His 119 in position 111, for example, may be a significant nonproductive complex affecting both K, and the turnover number. At pH 7 and above, His 12 and 119 and C M P are all largely deprotonated. Thus they would not be interacting with anions or with each other by ionic forces involving the histidines in the absence of additional protons from solution. An interaction with Lys 41 and the general cluster of positive charge would contribute to binding, and interactions of the pyrimidine with the protein would play a more dominant role. To the extent that the most stable complex EH,.CMP requires up to two protons from solution the binding would be second order in (H') and thus fall rapidly above pH 6 or 6.5 as observed. Relatively significant amounts of EHCMP and even ECMP probably exist a t these higher pH values, but E H 2 - C M Pmight still be the only productive complex in the reverse reaction. In this cyclization reaction a t high pH there is a net uptake of a proton, and this proton may enter the system in a rate limiting step leading to complex formation or in the final productive transition. The binding of 3'-CMP a t various p H values is thus fairly well documented, and a rather limited set of reasonabIe phenomena have been clearly postulated. The steady state kinetic data are more extensive but less complete, and the explanations are more controversial and less well defined. The rate of reaction at low substrate concentration is proportional to the saturation rate, k,, and concentration of the array of enzyme-substrate complexes, which is proportional to l/K,; thus, the rate is proportional to k,/K,. Since stabilization of the complex by changing pH, for example, increases the concentration of the complex at the same time as it decreases the probability of activation to a transition state, the net result is that k,/K, is related to the free energy of the activated complex relative to the unbound enzyme, an unbound substrate. The p H profile of k,/K,,, thus potentially reveals the pK values of groups on the free enzyme and free substrate that are involved significantly in rate limiting processes. This well-known relationship has been used to establish the pK values of 5.4 and 6.4 for groups on the enzyme in 0.2 M KCl for both

804


steps 1 and 2 in the overall reaction for a variety of substrates. Slightly different values reported in other conditions can in part be reconciled with the variation of histidine pK values with ionic strength or the specific salts present. (NO,-, SO,'-, and imidazole, for example, are all inhibitors.) The catalytic alkylation of His 119 with iodoacetate exhibits similar pK values when adjusted for ionic strength. The variations of the histidine pK values with ionic conditions may in turn result in large measure from ion binding and internal hydrogen bonding as discussed above. The pK values detected from k,/K,,, are macroscopic values, and in fact the microscopic values may be composite curves for each histidine. The NMR values of 5.8 and 6.1 are much closer to each other than the steady state kinetic values. The resolution of this question would not distinguish between involvement of the two histidines in the mechanism as a hydrogen-bonded system, in a push-pull linear mechanism, or in an adjacent mechanism with one histidine acting as a shuttle while the other protonates the phosphate in an intermediate or transition state. The only conclusion that can be drawn is that double protonation or zero protonation of the two histidines slows down a rate limiting function. The variation of the limiting rate Ic, or k, with pH, which can be obtained as an extrapolated value, does not relate to conditions in the free enzyme or free substrate but rather to pK values of intermediate complexes or transition states. For example, Herries et al. (496) determined values of 6.3 and 8.1 for complexes in C > p hydrolysis compared to 5.2 and 6.8 for the free enzyme. del Rosario and Hammes (499) deduced values of 5.8 and 7.5 for U > p hydrolysis. It is agreed that the increase in pK for the group acting in the protonated form is reasonable since the proton binding would be enhanced by the negative cyclic phosphate or a pentacovalent intermediate. The interpretation of the increased pK for the histidine which acts as a base is more controversial. Some argue that a dianionic intermediate must exist to explain the shift, while others point out that the proximity of two or three nonionic oxygen atoms could do equally well. The shift is not nearly as large as that observed by NMR for 2'-CMP binding where both pK values are 8.0 or higher. One must also question why the pl' values on the free enzyme are lower than the normal value of 6.7 which is observed for His 105. One explanation is that the cluster of positive charges repels protons from that region, and the ionic strength effects support this argument, C > p would then be expected to raise both pK values. The histidine hydrogen bonding scheme also would explain why one pK for each histidine on the free enzyme is low and thus why the average pK of each is low. Breaking this system with C > p could then allow both pK values to rise or not depending on whether the substrate sterically blocked

24.


805

one site. The specific accessibility of each histidine, the “dielectric constant” of the location, the hydrogen bonds to the other groups of the protein, the mobility of His 119 and immobility of His 12, and the detailed solvent structure in this area should all affect the pK of each histidine by affecting both the entropy and entholpy terms. Several of these might also directly affect the magnitude of the chemical shifts observed by NMR and the relative residence time of a given proton between two bases. Thus arguments from the titration data lead t o the conclusion that protonation of both histidines or deprotonation of both in the complex slow down the reaction and that protonation of both histidines is probably favored by complexing with the substrate. The lc, curve for C > p hydrolysis is not symmetrical, and a secondary “acid pathway’’ has been proposed to explain part of the activity a t pH 4.0 (499). Perhaps the protonation of both histidines produces enough polarization of the phosphate to permit direct attack by water or stabilizes the transitional state in the formation of the pentacovalent intermediate. With excess protons present, protonation of the leaving group would be easy. If histidine is not involved as a base, the k,/K, curve should also be distorted according to this view. This description would not seem to fit their model. In any case it would be interesting to see if this acid pathway is different from the normal pathway with respect to in-line vs. adjacent mechanisms. An alternate proposal is that the protonation of the acetate-tris buffer was distorting the curve. Another factor to consider is the chloride behavior as a competitive inhibitor, but this would not affect k, if it is strictly competitive. If the histidine titration curves are composite in the free protein as observed by Ruterjans and Witzel but not in the substrate complexes, the ks/Km curve should be distorted but not the k, curve. The opposite is observed. The variation of K , with pH depends on the pK values of free enzyme, the free substrate, and the complexes. The absolute value will also depend on other interactions including the ionic interaction with Lys 41. I n one of the simplest cases K, is equal to the overall equilibrium dissociation constant for all of the intermediates preceding a decisive rate limiting step. If this is not the case, K m is increased when the transition to the product is facilitated since the preceding intermediate state is occupied less of the time. K , is decreased when an intermediate is stabilized and also when a nonproductive isomer is stabilized. I n the simplest case the only intermediate state is the first complex formed, and this is in rapid equilibrium with the separate components. Hammes has shown that there is isomerization of the first formed complex, and whether or not this is on the reaction path the interpretation of the detailed kinetic

806


parameters is complicated. The simple situation for C > p is quite similar .to that already discussed for 3’-CMP. The most obvious difference is that the pK of the C > p is not in a range of direct concern. The monoanion could bind to the doubly protonated system without discharging one proton; therefore, one of the main explanations for loss of 3’-CMP binding at low pH does not hold for C > p. Irie (508) presented evidence that competitive binding of salt is important and favored this explanation. Witzel made the case for a prominent stabilized pentacoordinate intermediate and pH effects on the rate constants of formation and forward and reverse breakdown of this state. del Rosario and Hammes (499) involved a possible acid pathway to simultaneously explain the anomalously high values of k. and K, a t low pH.

Author Index Numbers in parentheses are reference numbers and indicate that an author’s work is referred to although his name is not cited in the text.

A Abderhalden, E., 130, 132, 136, 149(104) Abdulla, Y. H., 369 Abdumalikov, A. K.,103 Abendschein, P. A., 80, 82(6), 84(6), 93(6), 101, 107(2), 110(2), 126 Abraham, E. P., 23, 24, 27, 28(37), 30, 31(37), 34(29, 37), 38(31, 37, 80), 39 (29, 31, 32, 371,40(31), 42(31), 43 (31,37), 44(47) Abrash, H. I., 773 Abrosimova, N. M.,50 Abrosimovadmelyanchik, N.H., 245(137, 1381, 249 Abul-Fadl, M. A. M., 423, 451, 477, 478, 479, 480 Accorsi, A., 619, 620, 627, 628(35), 629 (40), 631(38, 39) Acharya, A. S.,731 Adams, J. M., 234, 237(107), 238(107) Adamson, R. H.,121 Adler, J., 316 Adunts, G.T.,802 Agren, G., 396,419,423 Ague, S. G.,442 Ahluwalia, G.,391 Ahmad, F.,44 Ahmed, Z.,436,440(159) Aiaazi, M., 609 Aida, K.,75, 149 Ainis, H.,103 Akinrimisi, E. O.,732 Akita, S., 124 138(9) Aksel’rod, V. D.,245(137, 138) Alberici, M.,367, W(49)

Albers, E., 451 Alben, H.,451 Albertsson, P.A., 650 Alberty, R. A., 88, 710 Albrecht, H.P.,783 Aldridge, W. N.,409, 410, 441, 605(273), 608 Alexander, H. E., 757 Akxander, M., 177(11), 178, 185(11), 193 (11) Alexianu, M., 605(286), 606 Alger, T.D., 770 Allan, R.,74 Allen, F.W., 364, 710,711(270) Allen, J. M.,485 Allende, J. E.,673,674(94),699(94),703 (94) Allewell. N. M., 196, 656, 679, 683(61), 697(120), 763, 785 Allfrey, V. G., B 5 Alpers, D.H.,422 Altenbern, R.A., 104 Altescu, E. J., 750,751(410) Altman, K.I., 275, 281 Alvarea, E.F.,436,437,441(1f33) Alvino, C.G.,755 Amberg, S.,48 Ambler, R. P.,31, 33(63), 35(83), 37, 42 (63), 514 Amirkhanova, 9. N., 750, 751(408) Anagnostopoulos, C.,427 Anagnostou-Kakaras, E.,419 Anai, M.,261 Ananta-Narayanan, P.,140 Anastasi, A., 128(57, 581, 129 807

AUTHOR INDEX

Andersen, J. A., 14 Anderson, C. E.,6Q5(304), 607 Anderson, C. M.,140 Anderson, D. G.,759 Anderson, E. A., 710 Anderson, L., 378, 384(44), 385(44), 386 (441, 393(44) Anderson, R. L.,639 Anderson, W.A., 606(329), 808 Anderson, W.B., 408, 580(141), 581, 587 (141),594(14), 598(178), 599 Ando, T., 221 Andrews, A., 8(43), 9. 10(43), 11(43), 12(43) Andrews, A. T.,110 Andrews, P., 55, 140 Anfinsen, C. B., 154(21, 22, 23), 155(5), 156(22), 160(22), 163(36, 37, 38), 166 (30), 172(29, 30, 32, 33), 173, 174 (28), 175(25), 177, 179(3), 180(20), 181(25, 26, 27), 182(21, 23, 2.8, 27)) 183(40), 184(47), 185(3, 20, 25), 186 (3, 46), 187(3, 40. 61, 62), 188(61), 189(61), 190(3, 61), 191(3). 192(61), 193(3, 63, 681, 195(48), 196(47, 48. 53), 197(47, 48, 53, 801, 1W47, 801, 199(80), 200(48, 801, 201(87, 881, 202 (85, 87). 203(88, 89, 901, 333, 334 (31), 648, 654, 669, 670(75), 671, 673, 675, 677, 678(105, ill), 681(111). 691, 693(169a, 179), 694, 695, 699 (89), 703(89), 705, 710, 715(296), 716 Angeletti, P. U., 477,484 Anraku, Y., 244(144, 145), 249, 333, 334 (331, 340, 349(16), 350(16), 356. 357 (2.3), 358, 359(2, 3), 360(2, 3), 361, 362(13), 363(11) Antonini, E., 71 Apgar, J., 48, 215, 222(29) Apirion, D.,243, 245(132), 376 Aposhian, H.V.,258 App, A. A., 641 Appella, E.,128(70). 129(70), 130, 272. 273, 274(12) Appelmans, 484 Appiani, G.,124 Applebury, M. L.,379(54), 380. 382(48). 387(48), 388, 389(48, 641, 40264). 403(48), 405(54, 129), 427, 444(10)

Appleman, M. M., 369 Appleyard, J., 472 Aqvist, S., 129 Arber, W.,263, 264(52) Archer, J. G.,435 Archibald, R.M.,132 Arens, A., 108, 109(65), 111, 112, 113(65), 114, 115, 116, 118(65) Arima, K., 213, 240, 244(119, 120, 121) Arima, T., 208(7), 209, 210(7), 211(7), 215, 227(30), 228(30), 231, 233(103), 234(30), 235(7, 30, 106), 236(30), 237 (30, 106), 238(30), 239(30), 248(7, 30, 106) Arion, W. J., 544, 546, 547, 549(37, 39. 401, 553(5, 371, 557, 558(37, 41, 93, 98), 560(93, 941, 563, 564, 566, 667 (35, 37, 40, 411, 568(37, 40, 41, 93, 94, 981, 569(98),570(41), 573(37, 40, 41, 981, 574(37, 40, 41, 981, 575(37, 41, 981, 576(37, 40, 41), 577(37, 40, 41), 579(40, 411, 580(37, 40, 411, 581 (371,583,685(37), 587(37, 39,40,41), 590(37, 40, 411, 597(37, 40, 411, 598 175, 121),599, 600(37) Armentrout, R. W.,127, 147(45), 148, 149(45, 46) Armstrong, A. R., 418 Arnaie, G.R.,367, 368(49) Arnon, D.I., 643 Arnone, A., 165, 156(31). 157(42). 164 (31), 173(31), 183, 187(49), 195(49) Amott, S., 158 Arora, B.S.,581(147), 582 Arora, K.L.,55 Arrison, R.,103 Arsenis, C.,343,351(31), 352(31) Asadourian, A. S.,713 Asakawa, K.,429 Aschaffenburg, R.,428 Ashcroft, S.J. H., 604 Ashmore, J., 544,548,552,557,568,580 (7). 581(7). 596(6), 597(169), 598(7), 600(211), 601, 606(332), 808, 613 Askari, A,, 66,67(142), 68(137, 140,1421, 69(137, 142) Aslanyan, I. G.,602 Assenhajm, D., 455,457(29) Atherly, A. G.,30.31(55),33(55) Atkinson. D.E.,48,613

AUTHOR INDEX

809

Atkinson, M. R., 64, 66(124, 148), 67 Banfield, J. E., 39 (IN), 68(148), 256, 257, 25809, 31) Barancik, M. B., 683, 699(139), 703(139) Baranowski, T.,96 Attias, J., 390 Barber, E. D., 71 Atwood, K.C.,639 Barber, M.,24 ArrbelSadron, G.,283 Barbour, S.D.,259 Auricchio, S.,600(187), 601 Barden, H.,604 Austrian, C.R.,48 Barendregt, T. J., 136 Avaeva, S. M.,394, 530, 531(7), 534 Bargellesi, A., 627 Avers, C.J., 609 Barka, T., 485 Avey, H.P.,656 Barkemeyer, H.,698 Avrameas, S., 428 Barker, G. R., 353,747,755 Axelrod, B., 451,465,472 Barkley, D.S.,366 Ayliffe, G.A. J., 42 Barman, T. E., 409, 410(146), 424, Azegami, M.,215(36), 216 437(83), 439(83), 443(83), 444(83), Azuma, Y.,246(161, 162), 250 566 B Barnard, E. A., 243, 648, 681, 682, 688. 690(135,136), 733,775 Babkina, G. T., 313,314(4) Barrel], B. G.,234, 237(107), 238(107), Babson, A. L.,433,456,457 240 Bach, D.,103 Barrera, H., 750, 751(415) Bach, M.K.,759 Barreto, A., 154 Bachrach, H.L.,292,750,751(413) Barrett, H., 397,407 Bachynsky, N.,118 Baer, H.P.,51,52(65a), 58(65a),59(65a), Barrett, K.,375, 379(21), 382(21), 385, 389(21),394(21), 395(21) 60(65a), 61,62,63(115),347, 351(68), Barrnett, R. J., 609 35268) Barron, E. S., 469 Baev, A. A., 321 Barron, K. D., 347 Baginski, E. S.,567 Ba.ny, J. M.,107 Baglioni, C.,130 Bartley, J. C.,600(188), 601 Bahr, J. T.,777 Bases, R.,496 Bailey, C.J., 5,9(25), lO(25) Basile, R.,605(301), 607 Bailey, J. L.,654,691 Bassham, J. A., 540,641,643 Bain, J. A., 750,751(397) Bastide, P., 603, 605(294), 606(335, 3371, Bajusz, E.,347 607,608 Baker, B. R., 77 Batchelor, F. R., 38(79), 39, 40(79), Baker, L.,614 41 (79),42(79) Baker, W.,27 Bates, C. J., 110 Bakwin, H.,420 Bauer, C.,58 Bakwin. R. M.,420 Bnldwin, R.L..308,310(65),504,708,709. Bauer, E., 525 Bauer. K.,108. 109(65), 111(65), 112(65). 727 113(65), 114(65), 115(65), 116(66), Balinsky, J. B., 50 118(65) Ball. E. G.,96 Bauer, R. J., 51, 52(65), 53(65), 59(65), Ballard, F. J., 600(197), 601 60(65), 61(65), 63(65) Baltscheffsky, H.,539 Bauer, W.,260 Baltscheffsky, M.,539 598(176), 599 Bnmann, E.,393, 418(17), 419, 446, 451 Baum, H., Bamman, B., 55, 56(92), 65(92), 70(92), Baumann, G.,602 Baumann, P.,640(93), 641 73(92), 74(92) Baumler, A., 553, 557 Bnnerjee, G.. 606(332), 608,645

810 Baust, P., 606(328), 808 Baxter-Grillo, D.L., 603 Bayev, A. A., 245(137), 249 Beams, J. W., 705,708(248), 707 Beaudreau, C.,630,631 Beaufay, H.,546, 553(26, 27), 554, 555, 557, 568(25), 577(26), 578(26), ssO(25, 261, 581(25, 261, 590(25, 26) Beaven, G. H., 697, 698(221) Beck, A. K.,741 Beck, J. V.,50, 76 Becker, A., 351 Becker, B. J. P., 605(314), 807 Becker, R. R., 677, 678(115, 117), 740 Becking, G.C.,309 Beckman, G., 454 Beckman, L.,454 Beecham, A. F.,131, 133 Beers, R.F.,Jr., 755 Beevers, H.J., 643 Behal, F. J., 245(149), 249, 345, 351(49), 356, 357, 359(4), 380(4) Behnke, O., 347 Behrendt, H., 477 Beintema, J. J., 654,655 Belaj, K.,548,605(52) Belding, M.E.,15,17(86) Belfanti, S.,419 Belfield, A.,567 Bell, E.J., 640 Bello, J., 656,657(60), m(69), 694, 717, 734 Benacerraf, B., 489 Benassi, C. A.,691 Bencovic, S.J., 616 Bender, M.L.,85,90,91(23),444 Bender, M.Z., 90,91(24) Bendich, A., 77 Benedetti, E.L.,344 Ben Hamida, F.,243,245(133) Bennett, N. G.,394, 395(107), 411(107), 442, 445(177) Bennett, V., 540 Benson, A. A.,641 Berenblum, I., 432 Berends, F.,471 Berg, P.,332,501(10),502 Bergel, F.,76,103,107(18), 117(18) Berger, A, 654 Bergman, M.,133

AUTHOR INDEX

Bergman, S., 454 Bergmann, F.,332,397,409(118) Bergmann, F.H., 501(10), 502 Bergmeyer, H. U.,5 Bernardi, A., 272, 275(10), 278(10), 283, 284,330, 331(11, 121, 332(11), 334(11, 35), 335(11), 493, 494(103), 495(103) Bernardi, G.,260, 272, 273, 274, 275(6, lo), 276(13), 277, 278(6, 9, 10, ll), 279(6), 280(16), 281(33), 282(6), 283, 284(19), 285(6), 287, 329,330(5), 332, 333(30), 334(35), 336, 493, 494(103), 495(103) Berne, R.M.,347 Berne, R. N.,63 Bernfield, M.R.,130, 141(76), 747 Bernheimer, A. W.,282 Berns, D.S.,14 Bernsohn, J., 347 Bernstein, I. A.,639 Bernt, E.,5 Berry, A. J., 51, 52(68), 64(68), 65(68), 66(68),67(68), W 6 8 ) Bertaccini. G., 128(58), 129 Berthet, J., 484, 545, 546(21), 548(24), 549(24), 553(26), 577(28), 578(26), 580(26), 581(26), 590(26), 600(21) Berthet, L., 484, 546. 548(24), 549(24), 600(24) Beskid, G.,40 Ressey, 0.A.,433 Bessman, M.D., 316 Bethke, R. M.,149 Bets, R.F.,49,50(37) Reychok, S.,694,721, 722(313) Bhalerao, V. R.,581(147),582 Bhargava, P.S.,59 Bhattacharyya, R. N.,631 Bhoornittra, D.,420 Bianchetti, R.,643 Bier, C.J., 155, 156(31), 157(42), 164(31), 173(31), 183, 187(49), 195(49) Bier, M., 427 Bigelow, C. C.,716, 717, 722, 723(320). 728,731,735.736,772 Biggs, M. L., 642(103),643 Bilimoria, M.H., 109 Billeter, M.A.,216 Bingham, E.W.,442

AUTHOR INDEX

Binkley, F., 94, 95(35), 96(35), 97, 423, 441, 540 Birkett, D. J., 428,443 Birnboim, H.C.,252 Bisaz, S., 429, 433(118), 439(118), 440, 443(118), 500 Bittman, R.,71 Bjork, W.,314,317(21),318,319,320(21), 326(13), 328(21),342,350(23),352(23) Bjorkman, N.,604 Black, A. L.,600(188), 601 Blain, J. A.,5 Blake, J., 133 Blakeley, R. L.,3,4, 5(13), 6(13), lO(13). 12, 15(59), 16(59), 17(59), 19(13) Blattler, D. P.,5, 7(27), 8(27), lO(42. 45), 18(27) Blauch, M.B., 49 Blears, D. J., 732 Bloch, A., 59 Block, W.D.,50 Blomback, B., 127, 128(44, 54,60,61,62). 129(44), 130(54) Blout, E.R.,505,713,739 Blumenthal, B. I., 539 Blumenthal, H., 376,497 Blunden, H.,149 Bobbitt, J. L., l a Bobrzecka, K.,469,470(60) Bock, R. M..48 Bodansky, A , , 418,422,433(13) Bodansky, O.,338, 343, 344, 345, 351(29). 419,420,438,442,496 Bode, V. C.,264 Boeck, L. D.,108 Boggust, W.A.,55, 56(91) Bohak, Z.,731 Eohn, H.,699,700(231) Boivin, A.,373 Boler, J.. 128(59), 129 Boles, M.O.,656 Bollard. E. G.,2 BollC, A,, 501 Bollum, F. J.. 290, 302(11), 307, 308, 321 Boman, H. G.,24, 29(43), 30. 31(43), 32(43), 33(43), 34(43), 39(4, 43). 40(43), 41(43),42(43),43(43),46(43), 316, 466 Bona, C., 605(302), 607 Bond, R.P.M., 90,91(23)

811 Bondi, A,, 38(78), 39, 40(78), 41(78), 42(78), 44 Bonner, D.M.,230,247(100) Bonner, J. T.,366 Bonsignore, A., 616,618(27) Bonu, G.,605(311),607 Boon, W.R..27 Borghese, E., 421 Borin, G.,671, 699(244), 701(244), 702 Borisova, N.N.,773 Bornig, H., 536,540(23) Bornstein, P.,128(72), 129(72), 130 Boroff, D.A.,686 Borosa, L., 392, 393(100), 394(100), 395(100) Borrelli, J., 457, 495 Bos, C.J., 344 Bose, S. K.,269 Boser, H.,581(152),582,594( 152) Bossenbroek, B., 605(320),607 Bot, G.,568,598(117) Botvinik, M.M.,394,534 Bouchilloux, S.,612 Boulter, D.,5,9(25), lO(25) Bourne, G.H., 347,368,605(281,282,288), 606(333),608,609 Bovcy, F.A.,716 Bowers, C. Y.,128(59), 129 Bowkiewicz-Surma, E.,77 Bowler, C.,43 Bowles, M.E.,318 Bowsen, K. E.K., 120 Boyd, J. W.,104 Boy de la Tour, E.,501 Boyer, P. D., 380,525, 526(61), 540, 583. 587(155), 591 Boyer, S. H.. 434,436(149) Boyle, R.D.,709 Royse. E. A., 103, 104, 109(39), llO(39). 117(39), 118, 120 Bradbury, J. H.,724 Bradley, A.,358 Drudshaw. M..602 Rrady, T. G., 49, 50, 54(81, 82), 55. 56(81, 82, 91). 57(42, 82), 58, 64 Braga, E.A..530,531(7) Branir. H.,366,368(41),369(41) Rrandts, J. F.,727,729,740,741 Branion, H.D.,435 Bratton, A. C.,148

812 Braun-Falco, O.,605(312), 607 Bray, H.G.,125 Bray, R.C.,76, 103,107(18), 117(18) Breckenridge, McL., 368 Breeae, K., 711 Breslow, E.,771 Breslow, R.,394 Brestkin, A. P.,443 Bridges, J. M., 103,105 Briggs, M. H.,15 Bright, H.J., 71 Brightwell, R.,484,489,490,540 Brindley, D.N.,603 Brinkely, S. B., 40 Brisbois, L., 314, 318 Brock, M. J., 433 Brocklehurst, K.,746 Brockman, R.W., 362,375,378(15) Brodey, R.S., 117 Broh-Kahn, R. H.,545, 580(17), 600(17) Broome, J. D.,102, 103, 106, 107, 109, 117(55), 118(98), 119, 120(59, 98), 121(59) Broomfield, C.A.,675(98,99), 676 Brons, D.,380 Bronstein, 5.B., 645 Broude, N. E.,249 Brouillard, J., 425, 435(91), 439(91). 445(91) Brown, D.M., 329,751 Brown, G . B., 49 Brown, G. W., Jr., 48 Brown, J., 597 Brown, J. E.,671 Brown, J. H.,318 Brown, J. R.,691,693(182) Brown, K.D.,281, 291 Brown, 0.E.,281 Browning, E. T., 644 Brownlee, G. C., 240 Brudenell-Woods, J., 420,443 Bruhl, G.,144 Brunel, C.,423 Brunner, R.,44 Bruno, R.,800(213), 601 Brushtein, E.A.,719 Brutlag, D.,256, 258 Buchanan, B. B., 643 Buchanan, J. M.,110 Budowski, E.I.,249,318,653

AUTHOB INDEX

Bueas, F. W., 313 Bukenberger, M. W.,124, 126, 134(8), 135(102), 136(102) Bulger, R. J., 24 Bull, H. B., 711 Bunton, C. A., 438, 445(167), 4&(167) Burch, H.B., 420 Burch, P. J., lm,118(62) Burck, P.J., 108 Burger, R.,66, 68(141), 69(141), 72(141) Burgos, M. H.,605(316), 607 Burk, D.J., 576, 577(135) Burlet, A.,605(289), 607 Burlet, C.,605(289), 607 Burlington, R.F.,600(205),601 Burma, D.P.,245(150), 249,641 Burman, L.G.,24 Burnett, F. F., 557, 559,560, 562,568(90), 578(90), 598(90), 605(291),607 Burnstein, Y., 692,693(184) Burson, G.,105, 107(43), 108(43), 109(43), 114(43) Burstone, M. S.,433,445(145), 496 Burton, A., 266 Burton, P. M., 506, 509(30), 510(30), 511(30), 512, 513, 514(34), 515, 516(45), 517(45), 518(45) Butcher, R. W., 366, 367(36), 368(36, 491, 369(36), 371(34, 35) Butcher, W.W.,454 Butler, G.C., 314, 317(15), 320(15) Butler, L., 530, 531(8), 532(8), 534(8, 91,535(9, 13), 536(9), 537(9), 538(9), 539(9), 540 Butler, L. G., 11, 586, 587(156), 591(156) Butler, R.,397, 407(121) Butterworth, P.J., 428,431 Butts, J. S.,149 Buszell, J. G.,710 Bykov, A., 375 Byrne, R.,378, 379(46), 382(46), 389(46), 424 Byrne, W. L., 544, 546, 555, 557(86), 563, 567(30), 579(87), 580(30), 582(29), 583, 586, 587(30), 591(30), 594(30), 596(30), 631

C Cahill, G. F., Jr., 552, 544, 596, 800(202), 601

813

AUTHOR INDEX

Cain, D. F., 72 Cabby, J. H.,72 Cdam, C. T.,27 Calcegno, M.,638 Calvet, F., 750,751( Calvin, M.,641 Cameron, E.,606(336),608 Cameron-Wood, J., 38(79), 39, 40(79),

41(79), 4!2(79) Cemmack, K.A., 116, 119(84) Cammarata, P.S.,126 Campbell, D. M.,419, 433(30), 435(30),

Cei, J. M., 128(58), 129 Center, M.8,246(149), 249, 265, 266(69), 345, 351(49), 356, 357, 359(4), 360(4) Cerami, A., 256 Cha, C. Y.,684,690(140,143) Chaiken, I. M., 200, 201(88), 202(85),

203(89) Chaikovskaya, S. M., 34(76), 39 Chain, E. B.,23, 27, 38(79), 39, 40(79),

41(79), 42(79), 432

Chakrabortty, H. C., 641 Chakraburtty, K., 245(150), 249 Chakravorty, M.,641 457 Campbell, H. A., 1Q3, 101, 105, 109, Chdkley, H.W., 50 llO(39, 411, 114, 117(31, 391, 120(41, Chance, E. K., 461 Chang, P.K.,110 87) Chang, 9.H., 216(33), 216,227 Canfield, R. E., 181 Chang, T.M.S., 12 Cannan, R.K.,125, 131 Chang, Y.Y.,366,368(39) Cantero, A., 581(153), 582,602 Cantoni, G. L., 207, 289, 319, 330, 331, Chanley, J. O.,446 Chappelet, D., 405,406(138), 413(138) 332 Charbonnier, A., 605(283),606 Canvin, D. T., 643 Chargaff, E., 473, 757 Caplow, M.,85 Charreau, E.H.,605(315), 607 Carbini, L., 605(299),607 Chassy, B. M., 51, 52(70), 53(70), 59(70), Carbone, P.P.,16,17(91) 60(70), 61(70) Carew, E.B.,708,709(257),727(257) Chen, C.-S., 313 Carey, M.J., 428 Chen, P. S.,432 Carlisle, C. H., 656 Chen, R.F.,184 Carlson, W., 785 Chen, S.Y.,59e(lSO), 599 Carpenter, F.H.,181 Chen, Y.,342,351(27),352(27) Carrara, M.,284 Chernitskii, E.A.,10 Carrier, W. L., 269,321 Chersi, A.,272,!275(10),27S(lO), '284, 330, Carrington, H.C., 27 493,494, 495(103) Carroll, W. R., 654,735,769 Cheung, C . 4 . S.,773 Carruthera, C., 553,557 Cheung, W. Y., 366, 367, 368(50, 55) Carter, C. E.,50,245(146), 249 369(53), 370(43) Carter, J. R.,691 Carty, R. P.,677,678(109. 1101,680(109, Chevalley, R.,601 Chevillard, L.,457 110) Chibata, I., 149 Carver, J. P.,505 Chibnall, A. C.. 105, 132 Caskie, M.,5 Chien, J. R.,252 Casula, A.,605(299), 607 Childs, B.,420 Cathala, G.,423 Chilson, 0.P., 56, 57(98), 58(98), 64. Cathou, R.E.,769,777 65(123). 69, 73(153b) Catley, B.J., 292, 293 Catlin, B. W.,177, 186(1), 252. 256(6) Chin, C. C., 4, 5(20). 7, 9(36), 10(20), Catsaras, M.,13 11. 20(56) Chio, K. S., 750, 751(415) Cavalien, L.F.,304 Chiquoine, A. D., 602 Cecil, R.,691 Choate, W.L., 651 Cedar, H., 109

814

AUTHOR INDEX

Choi, H. U., 777 Chopra, S. L.,11 Chou, C.-L., 697,698(218) Chou, J., 356, 362(10) Chow, K. Y., 149 Chung, S. T., 75 Chyka, G., 605(302), 607 Chytil, F., 366, 368(41), 369(41) Ciegler, A., 101, 105, 109(42), 117(37), 119(42)

Citri, N., 24, 25(2), 26(2), 28(38), 30(2),

Cohn, M., 430,519,538,771 Cohn, R. M., 544,596 Cole, D. F., 608(336), 608 Cole, P. W., 80 Cole, R. D., 512, 513(38), 691 Cole, T., 708, 709(257), 727(257) Coleman, C. M., 433 Coleman, D., 133 Coleman, J., 750, 751(398), 758 Coleman, J. E., 376, 378(27), 379(27, 5 4 ) , 380, 382(48), 387(48), 388, 389(27,

48, 54), 402(54), 403(48), 405(54), 31(2, 54), 32(2, 571, 33(2, 54), 34(38, 427, 444 (101) 75), 35(2), 38(77), 39(2, 38), 40(2, 38, 75, 771, 41'3, 75, 771, 52, Coleman, R., 344 54, 571, 43(2), 44(38, 75, 771, 45(2, Collins, P., 54(82), 55,56(82), 57(82) 38, 52, 57, 75, 77, 93, 101, 112, 1 1 5 ~ Collip, P. J., 567, 568(116), 598(116, 1801, 599, 800(116) 46(52, 74, 101, 115, 120) Colowick, S. P., 65, 66(127), 67(127), Clark, A. J., 254,259(15) 70(135), 207 Clark, D. A., 49 Concustell, E., 247(168, 169, 170), 250 Clark, H. E., 132 Cone, J. L., 154(23), 155, 179, 180, Clark, P., 303, 770 181(27), 182(23, 27) Clark, S. L., 421 Conn, J. B., 698 Clarke, A. M., 707 Connell, G. E., 121, 126(5), 142, 144(5) Clarke, D. A., 489 Conradie, A . R., 50 Clarke, D. D., 80 Contardi, A., 419 Clarke, H., 347 Contaxis, C. C., 2, 5, 7(27), 8(7, 27), Cleaves, D. W., 131 9(7), 18(27), 20(7) Clement, G. E., 85 Conway, E. J., 3,49,50(38) Clementi, A., 102 Conyers. R. A. J., 426,443 Cliffe, E. E., 125, 142 Cooke, J., 654, 677, 678(111), 681(111), Cloetens, R., 426 695 Clouse, J. A., 71(168), 73 Cooke, R., 49,50(38) Clubb, J. S., 428 Coon, M. J., 534 Clyman, M., 496 Cooney, D. A., 110 Cobbin, L. B., 63 Cooper, D. E., 40 Cochran, D. C., 72 Cooper, E. J., 291, 299(19, 20) Cocucci, E., 422 Coddington, A., 51, 52(66), 55(66), 62(86) Cooper, O., 96 Cooperman, B., 530, 532(6), 534(6). Coffey, D. S., 11 535(6), 537(6), 539 Cohen, C., 505 Copeland, P. L., 638 Cohen, E., 293, 651 Copenhaver, J. H., 432 Cohen, J., 380 Corcoran, C., 692 Cohen, J. A., 471 Cordonnier, C., 260, 270, 281, 285, 287 Cohen, J. S., 154,724, 784 Cori, C., 545, 567, 568(115), 578. 580(13). Cohen, L. A., 127 598(115), 600(115) Cohen, P. P., 48 Cori, C. F., 48 Cohen, R. B., 604 Cori, G. T., 545, 578, 580(13) Cohen, S. R., 402, 403 Corley, L. G., 183 Cohen, W., 427 Cornet, J., 117 Cohn, E. J., 504

AUTHOR INDEX

Cory, J. G., 51, 52(72), 55, 56(84), 57(84) Cory, S., 227, 232(92), 237(92) Costa, E., 367 Costabile, F., 773 Cotlove, E., 419, 423(29) Cottam, G., 403 Cotton, F. A., 155, 156(31), 157(42), 164(31), 173(31), 178, 180, 182(28), 183(19), 187(19, 49), 195(49), 201(19), 203(19) Cousin, M., 319 Covelli, I., 684, 689 Cowans, J . A., 19 Cowgill, R. M., 685 Cowgill, R. W., 718, 719, 727(3@3) Cox, B., 69,71(151, 1671, 72 Cox, D. J., 709 Cox, R. P., 422,429,442 Cozzani, I., 348 Cozzarelli, N. R., 255, 257(24) Crabtree, B., 635, 644 Craig, L. C., 7, 9(35), 650, 651, 669(28, 301, 670(28, 30) Craig. L. T., 650 Cramer, F., 222(75), 223, 755 Crane, R., 568,569 Crawford, K., 27, 34(29), 39(29) Creeth. J. M., 6, 7,9, 12, 708 Crestfield, A. M., 380, 425, 650, 653. 669(25). 670(25). 686, 687, 688(160), 690(161), 691, 710, 711(270), 744, 745, 746. 750, 751(399) Creutzfeldt, W., 605(307), 607 Crick, F. H. C., 158 Crocken, B. J., 420 Crocker, A. C., 496 Crompton, B., 27. 34(29), 39(29) Crook, E. M., 746,749 Crouse. H. V., 609 Csopak, A., 402,403 Cuatrecasas, P., 154, 155(5), 166. 172(29. 30, 32, 331, 173, 174(28, 33), 175(25). 177, 179(3), 182(21), 183, 185(3). 186(3, 46), 187(3, 61, 62), 188(61), 189, 190(3). 191(3), 192, 193(3. 63. a),195, 333, 334(31) Cuchillo, C. M.. 247(1M, 169, 170), 250 Cunningham, B. A., 69,128(66), 129 Cunningham, L., 177, 186(1), 252, 256(6),

272, 291, 299(21), 301(21), 308, 314, 316(10), 353, 476 Currie, R. D., 65, 66(127), 67(127), 76 Curtis, W. C., 103, 106(11), 107(11) Curzen, P., 606(323, 324), 608 Cusurnano, C. L., 154(22, 23), 155, 156(22), 160(22), 180, 181(27), 182(27) Cutler, A., 567 Cyr, K., 398,406(123), 407(123), 408(123), 409(123), 410(123)

D Dabich, D., 423, 424(76), 435(76), 436 (76), 437(76), 443(76), 444(76)

Dabrowska, W., 17,291,299(19) Dalcq, A. M., 606(330), 608 Dallner, G., 578 Dall’Orso. F., 548,605(53) Darnmkoehler, R. A., 523 Damodaran, M., 140 Danesino, V., 605(319), 607 Dang, H. C., 13 Danyluk, S. S., 732 D’Ari, L., 409 Daria, G. M., 103 Das, N. B., 14, 17(75) DasGupta, B. R., 686 Dastugue, G., 603, 606(335, 3371,808 Datta, N., 26, 29(25, 42), 30(25), 31(25), 32(25), 34(25), 38(25), 39(25), 40(25, 42), 41(25), 42(25) Davidson, B., 505 Davidson, B. E., 692 Dnvidson, H. M., 451, 459(38), 466 Davidson, W. D., 612 Davies. D. R.. 207. 289. Davies; R. E.; 14,15(&), 72 Davis, B. J., 503 Davis, F. F., 384 Davoll, J., 49 Dawkins, M . J. R., 600(198), 601, 602 Day, R. A., 44 Day. V. W., 156, 157(42) Dayan, J., 394, 395(105), 398(105). 406. 407(123), 408(123), 409(123), 410(123) Deak, J. E., 698 Dean, M. R., 498 Deavin, A., 763, 780 DeBoer, E. J., 758

AUTHOR INDEX

Dierick, W., 55 Dierks, C., 777 Dingle, J. T., 286 548(24), 549(24), 553(26, 27), 554(27), Dipietro, D. L., 347,596 555(27), 557(27), 562, 568(25), 578 Dirksen, M. L., 185 (261, 580(25, 26), 581(25, 26), 590 Dixon, G. H., 380 Dixon, M., 436, 574, 591 (25, 261, 600(21, 24) Dmochowski, A., 455, 457(29) DeEstrugo, S. F., 69, 71(151) Doege. S., 609 De Flora, A., 616, 618(27) Doherty, D. G., 87 Degkwitz, E., 600(193), 601 Dolapchiev, L. B., 320, 322 DeGroot, N., 103, 104 Dolowy, W. C., 117 Deig, E. F., 602 Done, J., 375 Deimling, 0.V., 602 Donohue, J., 158 de Jarnette, E., 694 Dekker, C. A., 128(56), 129, 177(12, 14, Donovan, L. G., 684 151, 178, 185(12, 14, 15), 186(12), Doolittle, R. F., 127, 128(61, 62, 63), 129,

de Boer, H. G. A., 348 Decker, M. L., 606(325), 608 De Duve, C., 484, 485, 545, 546(21),

234, 248(109, 110), 304, 307(53), 408

Delbrueck, A., 605(305), 607 Della Monica, E. S., 436 Delluva, A. M., 14, 15(85) Delori, P., 314 Delory, G. E.,435, 447 deLoze, C., 713 del Rossrio, E. J., 769, 773 Delsal, J.-L., 432 de Maine, M. M., 616 de Meuron-Landolt, M., 185 Denhardt, G. H., 501 Denkewalter, R. A., 196 Denkewalter, R. G., 694, 698 Dennis, E. A., 784 Depue, R. H., 38(78), 39, 40(78), 41(78), 42(78)

Derieux, J., 378 De Robertis, E., 367,368(49) Derr, I., 118 Desnuelle, P., 11 De Stevens, G., 758 Deutscher, M. P., 256,257(27) Dewey, R. S., 6% de Wulf, H., 594 Diamond, R., 157 di Bella, S., 548, 605(50) DiCarlo, F. J., 49 Dickerson, R. E., 656 Dickinson, M. J., 698 Dickman, S. R., 651, 697, 698(215), 750, 751(4021, 778

Dieckmann, M., 332,501(10), 502 Diederichs, K., 451

147(45), 148, 149(45), 513

Dorow, C., 553, 554(77), 600(77) Doscher, M. S., 657,671,680,683,785 Doskocil, J., 284 Dosta, G. A., 548,605(54,55,56) Doty, P., 505, 722 Dowben, R. M., 606(331), 608 Downing, M., 741 Downs, C. E., 464,465 Drappier, A., 390 Dreisbach, R. H., 432 Dreyer, W. J., 128(69), 129(69), 130, 181, 627

Drummey, G. D., 422 Drummond, G. I., 51, 52(65a), 58(65a), 59(65a), 60(65a), 61, 62, 63(115), 347, 351(68), 352(68), 364, 365(29). 366, 367(37, 441,368(37) Dryden, E. E., 50 D’Souza, L., 44 Du, Y.-T., W, 698(218) Dube, S. K., 755 Dubnau, D. A., 42 Duchateau, G., 49 Duerksen, J. D., 30 Diitting, D.. 218, 223(48) Dumford, 9. W., 18 Dumitrescu, M., 605(302, 3031, 607 Duncan, L., 347, 351(68),352(68) Dunkley, C. R., 50 Dunn, M. S., 149 Dupret, L., 545, 546(21), 600(21) Dutcher, J. D., 126 Duttera, S. M., 555, 557(86), 579(87)

817

AUTHOB INDEX

du Vair, G., 313,314(8), 316(8), 342 Dvorak, H.F.,339, 340(10), 349(16), 350 (101,362, 363031,375 Dyson, J. E. D., 580(141), 581, 587(141), 594(141)

E Eaker, D. L., 651,689(28, 30), 670(28, 30) Earle, A. S., 600(202), 601 Easwaran, C. V., 327 Eaton, R. H.,419(32), 420, 424, 429,431, 437( 116) Echols, H., 254, 377 Eckstein, F.,394,395(1CbS), 768 Edelhoch, H.,155, 166(30), 172(30), 183, 186(46), 750, 751(398), 758 Edelman, G. M.,128(68), 120, 330, 715 Edelstein, M.,454 Edgar, R.S.,501 Edman, P.,128(54, 60), 129, 130(54), 181 Edsall, J. T., 504,591 Edward, T.,770 Egami, F.,207,208(5, 6, 7), 209, 2106 7), 211(5, 7, 8), 212(5), 213(5, 12), 215(32, 35), 216(11, 18,27), 218, 219, 220(11), 22l(ll), 222(74), 223(5, 121, 224, 225, 226(27, 85), 227(30, 32, 35, 85), 228(30, 84), 230(6), 231(6, 81, 232(6, 8, 104). 233(103, 104), 234(7, 30), 235(7. 30, 106), 236(30, 35), 237 (30,106, lOS), 238(30, 108), 239(30). 248 241, 247(6, 8, 102, 105, 123, la), (7, 30, 106, 1731, 250, 048, 756, 757 Egan, T., 603 Eggermont, E., 605(Z71), 806, 617 Egorov, A. M.,530,531(7) Ehrenfeld, E., 85, 94(12), 97(12) Ehrlich, S.D.,277,284(19),332 Eichhorn, G.L.,303,770 Eifler, R.,531, 534(12) Eigen, M.,71 EIAsmar, F.,105 El-Badny, A. M.,540 Eldjarn, L., 623 Ellfolk, N.,125,126(17). 128(17) Elliot, A., 6,9(32) Elliott, W.H., 105(51), 108 Ellis, B. W., 126 Ellman, G.L.,512

Elsevier, E., 246(167), 250 Elson, E. L., 308,310(f35) Elsworth, R.,104, 116(38),119(38) Ely, J. O.,436 Emmelot, P.,344 Engel, A. C., 71 Englander, S. W.,713 Englhardt-Goelkel, A.,605(278), 606 English, A. R., 40 Englund, P. T.,258 Engstrom, L., 380,396, 397,398(114, 1151, 405(114), 419, 423(22), 424(2!& 71), 425, 427(22), 436(20), 437(20), 439 (71,88) Enser, M., 620, 622, 623(45), 624(45), 630(42), 631(42), 632(04), 633(63,041, 645(42, 63) Enzmann, F., 128(59), 129 Epand, R. M.,712,724(281) Epstein, C. J., 677, 678(116), 694, 695 ( 116) Epstein, R. H., 601 Epstein, S.,603 Ercoli, A., 419 Erdtman, H., 418 Erecinska, M.,327,336 Eremenko, V. V., 105 Erikason-Greenberg, K. G.,24, 39(4) Eriksson-Quensel, I.-B., 8, 10(40), 12(40) Erlander, S. R.,709 Erman, J. E., 769 Ernster, L., 553, 554, 558(70), 582, 569, 579(70), 600(70) Enpamer, V., 128(57, 5 8 ) , 129 Essner, E., 485 Estborn, B., 496 Eto, Y.,246(160), 247(171), 250 Ettinger, M.J., 739 Evans, E. A., Jr., 266 Evans, S. A., 658 Everett. G. A.. 48,215,222(29) Evseev, L.P., 105 Exton, J. H.,044 Eyring, E. J.,805(304),607 Eyring. E . M.,114 Evring. H., 59

F Fabro, S., 121 Fahey, P.F.,707

818

AUTHOR INDEX

Fahrney, D., 697, 698(222) Fairley, F. L.,353 Falco, E. A., 76 Falk, K. E.,403 Falkenheim, R.,275 Fallon, B. M.,654 Fan, D. P., 385 Fankuchen, I., 654 Fantl, P.,545, 580(16), soO(l6) Fareed, G.C.,304,306(56a) Farnararo, M., 422 Farnden, K.J. F., 105(51), 106 Farnsworth, J., 654 Fasman, G. D., 505 Faeaina, G., 177(6), 178, 185(6), "6) Faulkner, R. D.,228 Feageson, E.,446 Federman, M.,557, 559, 560(92) Fedorova, N. A., 3% Feldman, D., 423 Feldman, F., 586, 587(156), 591(156) Feldmann, H.,218, 223(48) Felicioli, R. A., 750,751(423) Felix, F., 318 Fell, H. B.,286 Fellig, J., 248(174), 250 Felsenfeld, G.,154, 185 Fendler, E. J., 438, 445(167), m(167) Fennelly, J. J., 426 Fenster, L.F., 603 Fenton, 481, 482(81) Ferioli, V.,602 Fernando, J., 614, 630, 632(54), 633(64), 634, 645(74) Fernley, H. N., 392, 393(99), 394, 404(99, IN), 4 m , 410, 429, 431(117), 433 (118), 435(143), 436(143), 437(117), 438, 439(118, 165), 440, 441(170), 442 (117, 1701,443(118) Ferrarese, E., 699,701(238) Ferreri, A. E., sOO(nO), 801,602 Ferry, R.,461 Fetherolf, K.,378, 380(38), 384(38), 385

(38) Feuer, G., 578 Fiaccadori, F., 602 Fidler, J., 117 Field, J. B.,603 Fiers, W., 326, 750, 751(403, 404, 42%

Fife, W. K., 404, 410(136) Filipowicz, B., 333, 334(34) Filmer, I. D.L., 707 Findlay, D.,779, 780 Finean, J. B.,344 Finland, M., 31,40,43(66) Finn, F. M.,196, 698, 699, 700(227, 229, 230, 232,2331, 701(233) Fischer, G. A., 110 Fischer, H.H.,140 Fishbein, W. N., 3, 4(9), 5, 6(9), 7(9, 28), ll(9, 391, 16, 17(91, 95), 18(95, 97), 19(97) Fisher, C. J., 567, 568(113), 598(113) Fisher, E. A.,645 Fisher, J. R.,55, 56(83, 961, 57033, 98), 58(83, 98) Fisher, R. A.,483 Fisher, R. C.,763 Fishman, W. H., 419, 420, 422, 423, 424 (81), 426(42), 428, 436(42), 437, 442 (26,42), 443(26), 451, 454, 457, 458 (38), 459(38), 466, 472(25), 473, 475 (25) Fiske, C. H., 432,487,518 Fittler, F., 755 Fitzgerald, M. X.,426 Fitzgerald, O.,3 Fitzgerald, P. L.,757 Fitzmaurice, M.A., 120 Flavin, M.,854 Fleisch, H., 421, 500 Fleming, J., 38(70), 39, 40(70), 41(70), 42(70), 46(70) Flinn, A. M.,609 Flint, M., 600(199), 601 Florescu, R.,602 Florey, H. W., 27 Florkin, M.,49 Foa, P.P.,567 Fodor, P.J., 96, 125, 142 Foldes, J., 24,39(4) Folk, J. E.,80 Folkers, K., 128(59), 129 Folley, S. J., 418(14), 419, 420, 421(14, a),428(35), 432(35), 44304) Follman, H., 775 Fontana, A., 691,699, 701(235, 237, 239) Forbes, M.,149 Ford, E.J. H., MO(189, 1951,601

819

AUTHOR INDEX

Foas, J. G., 733 Fossa, S., 548,605(53) Foasa, T.,579(145), 580(145), 582, 590 (1451,605(276), 606 Fossitt, D. D., 639 Foster, D. O., 645 Foster, J. F., 709 Fouquet, J. P.,606(327),608 Fowden, L., 133 Fox, L.E.,602 Foz, M.,548 Fraenkel, D.G., 614,639 Fraenkel-Conrat, H.,326, 327, 358, 476 Frank, B. H.,108, 113, 114, 118(62) Franklin, J. E.,Jr., 66, 68(137), 69(137) Frappez, G., 49 Fraser, P.E.,101, 120(1) Frederico, E.,177(16), 178, 185(16) Fredericq, E.,279,280(2l), 309 Frederiksen, S., 51, 52(71), 53(71, 74), 59(71), 60(71), 61(71), 63(74) Freedland, R. A., 600(1881,601 Freeman, G. G., 27 Freeman, H. C., 785 French, T. C., 724,766 Frensdorff, A., 675(102), 677, 678(113, 1141, 681(114) Frezal, J., 603 Fridovich, I., 53,58, 59 Fridovich, P.,54(80), 55 Friedberg, E.C., 264,265,268(56) Frieden, E.,800(219), 601 Friedenwald, J. S.,558(100), 559 Friedland, R. A.,645 Friedman, L.,733 Friedman, M.E.,684,690(144) Friedman, S.,49 Friedmann, B.,597 Friedrichs, B.,348, 351(80) Fritzon, P., 344, 315(36), 348(36), 351 (36) Fruchter, R. G., W, 745,746 Fruton, J. S., 90, 91(22, 26). 128(56). 129 Fuchs, E.,11 Fuchs, S., 154, 155, 172(29, 32, 33), 174 (28, 33). 177, 179(3), 182(21), 185 (3), 186(3), 187(3, 621, 190(3), 191 (3).193(3. 63). 195 Fujii, I., 216

Fujii, Y., 215 Fujimura, S.,317 Fujioka, H., 673,685,690(147) Fujiwara, T.,50 Fukami, M.H.,644 Fuke, J., 216 Fukuda, A., 772 Fuller, W.,757 Furuichi, Y.,222(76), 223, 240, 244(121) Furukawa, Y.,320 Futai, M.,317

G Gadsden, E. L., 600 Gale, G. R., 16,17(96) Galizaao, G., 682,683(131) Gall, W.E.,128(66), 129 Galletti, L.,602 Gally, J. A.,330, 718 Galton, V. A.,600(218), 601 Galy-Fajou, M.,64 Galzigna, L., 680 Gancedo, C., 614. 619(13), 640(13), 645 (13) Ganguli, N. C., 581(147), 582, 605(306), 607 Ganozs, M. C., 553,555,579(87) Ganther, H. E.,692 Garber, N.,31, 32(57), 38(77), 39,40(77), 41(77), 42(57), 44(77), 45(57, 77, 93, 112) Gardner, L. J., 348,351(75) Garen, A., 374, 376(4), 377(4), 378(4). 384, 385, 387(4), 389(4), 392(4), 393 (4), 394(4), 395(4), 406(4), 429 Garen, S.. 385 Garsky, V., 698 Gassen, H.G., 775 Gayle, R.,477 Gebhardt, L.P.,602 Gehring, A. W., 546 Geis, I., 658 Gelled, M., 267 Genchev, D.D.,320 Georgatsos, J. G., 314, 316(12), 328(12). 423 Gerfaux. G.. 423, 424(69a), 425(69a) Cerhards, E., 17 Gerisma. 5. Y..732 Gervasini, N., 548, 605(49)

820

AUTHOR INDEX

Gereeli, G., 605(292),007 Gesteland, R. F.,243, 245(131), 333, 334 (361,335 Ghiringhelli, F.,605(292), 007 Ghosh, N. K.,420, 423, 424, 426(42), 428, 436(42), 437(42), 442(42) Gibim, H.,17 Giblett, E.R.,480, 481, 483 Gibson, K. I., 578 Gifford, R. H., 354 Gigliotti, H. J., 94, 95 Gilbert, J. B., 132 Gilchrist, M.,85 Gilgan, M. W., 366,367(44) Gilham, P. T.,215(31), 210, 234, 237(31, la), 238(108), 321, 754 Gill, S.J., 741,744 Gillin, F.D.,30,31(56), 33(65) Gillis, J., 51, 52(65a), %(=a), 59(65a), 00(65a) Gilmour, D., 72 Gilsdorf, J. R.,553, 550(80), 557(80), 558 (80),560(80),574(80), 598(80), 599 (80),653, 55f3(80), 557(80), 558(80, 98), 580(80), 568(98), 509(98), 573 (98), 574(80, 981, 575(98), 598(80, 981,599(80, 98) Giner, A., 014, 019(13), 640(13), 045(13) Gingery, R.,254 Ginsburg, A., 673,735 Girotti, A. W., 771 Giunta, C.,000(213), 601 Glaesmer, R.,531, 534(12) Glasenapp, I. V.,144 Glaser, L.,339,340 Glazer, A. N., 721,723(312) Glende, E. A., Jr., 557, 558(93), 560(93). 568(93), 598(93), 599(93) Glick, D. M., 882 Glickman, R. M.,422 Glitz, D.G., 234,248(109, 110) Glogovsky, R.L.,744 Glowinski, M.,609 Goad, W., 307,308 Gockerman, J., 485 Goebel, W.F.,697, 698(217) Giirlich, M.,553, 557(75), 60005, 76,

78) Golberg, L., 578 Goldbarg, J. A., 148

Goldberg, A. F., 498 Goldberg, B.,605(317), 607 Goldberg, D.M., 507 Goldberg, M., 387,389(77) Goldberger, R.,677, 078(105, 110), 084, 091, 093(182), 694,095(110) Goldberger, R. F., 197 Goldblatt, P.J., eoO(lsS), 601 Goldman, S.S.,423,424(81) Goldner, M.,43,46 Goldstein, G., 498 Goldstein, J., 077,678(119).682(119), 735 Goldstein, M.,634 Goldthwait, D. A., 204, 20566). 288(50). 279,302,303(49), 307(49) Goloborod'ko, 0. P., 50. 66(51). 67(51) Gomori, G.. 433, 012, 013, 016, 017, 018. 629 Goodban, A. E., 120 Goodman, E.H., Jr., 597 Goodman, R.M.,609(354),610 Goodrich, R.,5 Goodwin, T.W.,296 Gorbunoff, M. J., 685 Gordon, J. J., 540 Gordon, M. P.,476 Goren, H.J., 682, 690(135, 136) Gorenigen, E.,307 Goria, M.,gOo(220),601 Gorin, G., 3, 4, 5(20), 7, 8,9(14, 36). 10 (14,20,42,45), 11,15,20(50) Gorr, G., 103 Goslar, H. G., W(3281, 608, 609(355). 610 Goulian, M., 255,250(22),258(22) Goto, Y.,240(160), 250 Cots, J. S.,49 Gotterer, G. S.,420 Gottesman, M.,392, 393(96), 394(96). 395(96), 404(98), 408(96), 409(96). 410(96), 411 (96) Gottlieb, A. J.. 424, 425(86), 426(80), 437 (85) Gottlieb, P.D.,1!28(68), 129 Graham, J. M.,344 Gralen. N..8,10(40), 12(40) Grassman, W.,103 Gratser, W.B.,697, 698(221) Gray, M.W.,320 Gray, W. R.,128(69), 129(69), 130. 182

821

AUTHOR INDEX

Grazi, E.,615, 017, 018(21), 619,020,027, 628(35), 629(40), 031(38, 39) Greco, A. E.,239,244(114, 1401,247(140), 249, 281, 334 Greeley, S.J., 433 Green, C., 344 Green, H.,419,472 Green, M.H.,433 Green, R. H.,432 Green, S.,419,442(20), 443(20) Greenberg. B.,253, 261(10) Greenberg. D.M., 94, 105 Greenberg, E.,642(103), 643 Greenberg, H., 471,472 Greenberg, L. D., 149 Greenberg, L. J., 433 Greene, L.J., 651 Greenfield, N..505 Greenfield, R. E., 80, 82(0), 84(6), 93 (0), 101,107(2), 110(2), 126 Greengard, O.,598(177, 1791,599 Greenquist, A. C.,112, 115 Greenstein. J. P.,132,291 Greenstein, J. P..49,50 Gregory, H.,128(64), 129 Griboff. G., 291, 304(24) GriffB, M.,272, 278(9), 281(9), 283(9), 333, 33400) Griffin. C. C.. 600(186), 601 Griffin. M. J.. 422,429.442 Grillo, L.,314. 318 Grillo, T.A. I., 604 Grinnan, E. L., 108 Grisolia, S.,451 Grist. K.L.,880 Gross. D..120 Gross E..180,071,682 Gross. G., 248(175), 250 Gross, H. J., 227 Gross. H..005(279), 606 Gross. W., 430 Grosser, P..418 Grossman. L..258, 209(32), 270(76) Grossowicz. N.. 104 Grover. C.E..105 Griinherger. D., 222(77). 223 Grunberg, E..40 Guerritore. A.. 422 Guha. S.R..55 Guilbault, G . G., 12

Gulick, 2. R., 758 Gundlach, H.G., 686,091 Gunter, C. R., 85 Gunther, T., 553, 554(77), 000(77) Guschlbauer, W.,333 Gustafson, T., 49 Gutfreund, H., 392, 393(98), 394, 395 (107). 404(98), 408(98), 409, 410(98, 146), 411(107), 424, 437(83), 439(83), 442, 443(83), 444(83), 445(177) Gutman. A. B.. 420,455,457(30) Gutman, E.B.,455,457(30) Gutte, B.,196.673,094,098

H Haas. D. J., 667 Haber. E.,091,093(179), 094,096 Hacha. R., 177(16), 178, 186(10), 279, 280(21), 309 Hachimori, Y.,517 Hadi, S. M., 264. 205(56), zsS(50) Hadjiolov, A. A., 320 Haessler, H.A.,314,316(10) Hagerty, G., 290, 314,310(11) Haggis, G. H.,713 Haitinger, L., 124 Halford, S. E.,394, 395(107), 411(107), 442, 445(177) Hall, D. C., 502, 503(13), 504(13), 505 (13), 508(13), 509(13), 510(13), 512, 513(34), 5 1 4 W Hall, E. M., 366 Hall, J. R., 31,32(56) Hall, M. E.,755 Hall, T.C.,60 Halle. M.. 598(180), 599 Halmann. M.,677, 078(108), SSO(l08) Halpern, Y. S., 104 Ham, J. S.,127, 149(40) Hamada, M.,247(172), 250 Hamadah, K.,369 Hamilton, L. D., 158 Hamilton, R. M., 132 Hamilton-Miller, J. M. T.. 24. 26, 27. 38(82, 831, 39, 40, 41(90), 42(90). 43 (94,95) Hammer. R. A., 81, 83(8) Hammes, G. G.. 084. 724, 759, 705, 786. 789. 773. 777 Han, K., 378

822

AUTHOB INDEX

3anabusa, K., 4 Hand, D.B.,6,9(33),10 Handler, P.,58, 93 Handschumacher, R. E.,110,121 Hanes, C.S.,96, 124, 128(5), 142(5), 144 (6)

Hanke, M. E., 49 Hannah, J., 726 Hansen, A. W.,785 Hanslian, R.,136, 149(104) Hansman, F.S.,450 Hanson, A. W.,163, 656, 657(62), 658 (621, 666(62), 667(62), 72403% Hanson, D.M.,353 Hanson, T.E.,639 Hanson, T.L.,547, 549(42), 556, 557, 558 (981, 559(42, 881, 561(88), 563(88), 567(42), 568(42, 98), 569(98), 570 (421, 573(98), 574(98), 575(98, 103). 576(42), 579(42, 88, 143, 144), 580 (1031, 582, 587(103), 590(42, 103, 134), 592(103, 1341, 594(103), 595 (42, 881, 597(88), 598(98, 99, 175), 599(42, 98,99) Hara, L., 66, 67(142), 68(142), 69(142) Harada, F., 228, 227(86), 228(86), 230 (86,94), 240 Hardman, J. G.,370 Hardman, K. D., 196, 656, 683(61), 785 Hardonk, M. J., 348, 605(320), 607 Hardy, P.M.,128(64), 129 Harington, C.R.,133 Harker, D.,656, 657(60), 694 Harkness, D.,332, 376, 392(28), 393(28, 97), 394(28), 401(28), 419, 423(23), 424(23), 425(23), 427, 429(97), 441 (23),430(101a), 438(101a),442(101a), 443( 101a) Harper, A. E., 566, 645 Harrington, W. F., 675, 691, 693(169a), 705, 710, 715(296), 716 Harris, A. Z., 641 Harris, C.A.,50 Harris, H., 422, 423, 424, 477, 480(76), 481, 482(83), 483, 484 Harris, J., 182, 422 Harris, M.,376,378(27),379(27), 389(27) Harris, M.R.,783 Harris, T.N.,130 Hartenstein, R.C.,51

Hartley, B.S., 90, 91(18), 380 Hartley, R. W., Jr., 239, 244(114), 244 (1401,247(140), 249 Hartman, F.C., 696,720(212) Hartman, S. C.,81, 82(7), 83(8), 84(7, 9), 85(7), 86, 88(9), 90(9), 92(9) Hartsuck, J. A., 90,91(21) H e , J., 14(93), 16, 17(90, 93, 94), 20 (94) Hasegawa, S., 317 Hash, J. H., 246(167), 250 Hashimoto, J., 215(35), 216, 227(35), 236 (35) Hashimoto, S.,226 Hass, L. F., 546, 563, 567(30), 5W30), 582(29), 583, 586, 587(30, 155), 591 (30), 594(30), 596(30) Hasselberg, I., 49 Hastings, A. B.,544,552, 596(6) Hasunuma, K.,230, 247(101) Hatch, B.,304 Hathaway, J. A., 613 Hattori, T.,246(154), 249 Hauenstein, J. D., 651, 710,711, 715(274), 716(274) Hausamen, T.-U., 436 Hausmann, W.,531 Hayashi, I., 757 Hayashi, M.,356, 357(6, 71, 359(6, 7), 360(6), 361 (7) Hayashi, H., 216,222(74), 2!?3 Hayciahi, T.T.,72 Hayatsu, H., 240, 244(121), 752 Hazen, E. E.,Jr., 155, 156(31), 157(42), 164(31), 173(31), 178, 180, 182(28). 183(19), 187(19, 491, 195(49). 201 (191,202(19), 203(19) Heard, C. R.C., 600(204),601 Hearn, R. H., 740 Heath, D. F.,599 Heber, U.,643 Hedrick, J. L.,114 Hegarty, V. J., 50 Heidland, A., 602 Heinemann, B.,116 Heiney, R. E.,697, 698(214), 720(214) Heinrikson, R. L.,491, 492, 493, 677,678 (118), 684(ll8). 687, 688(118), 690 (161) Heins, J. N., 182, 183(40), 187(40)

AUTHOR INDEX

Heise, E., 553, 557(75), 600(75, 76, 78) Heitrman, H.A., 785 Helger, R.,436 Hellerman, L., 11 Hellerstrom, C.,604 Hellman, B.. 604 Henion, W. F.,367 Hennessey, T. D., 27, 29(45), 30, 31 (45), 33(45), 34(45), 39(45), 40(45) Henningsrn, I., 258 Henry, H., 64, 65(123), 66(123), 68(123). 69(123), 73(153b) Henry, R. J., 39, 43(71) Henson, D.,117 Hepp, K. D.,367, 368(45) Heppel, L. A,, 53, 54(76), 177(11), 178. 185(11), 193(11), 314(26), 315, 329. 330, 331, 332, 337, 338, 339, 340, 342, 349(16), 350(16), 361, 322(17, 23), 363, 374, 375, 376, 378(15), 392(28), 393(28), 394(28), 404, 426, 429(97), 521, 530, 534, 535(15), 538(5), 639, 746, 747 Herbert, D.. 104, 116(38), 119(38), 545 Hrrbut, P. A., 103, loS(17) Hercules, K.,268 Herman, R.H., 544,596(11) Hermans, J.. Jr., 711, 714, 727, 729(277, 337), 730(277, 337). 732 Herr, E. B.,446 Herries. D.G.,773,780 Hers, H. G., 545, 546(21), 548, 549. 553 (26), 577(26), 578(26), 580(26), 581 (26),590(26), 594, 600(21), 605(271). 606. 612, 617 Hrrsh, L.B., 147 Hersh. R.T.,11 Hrrskovits, T.T..717, 722, 723(318) Hrrzig. C. L.,772 Hrss, G.P.,90,91(19) Hrssel, B.. 128(54), 129. 130(54) Hemrr, P.E.,709 Hrymann, H., 758 Hrymann. W.. 428 Hiatt. H. H.. &?4 Hickmott, J. T.,Jr.. 708, 709(257), 727 (257) Higgins, J. A.. 344 Hilf. R.,605(291),607 Hill, H. D.,433

823 Hill, J., 713 Hill, J. M., 105, 107(43), 108(43), 109 (43), 114(43) Hill, R. M., 6,9(32) Hillborg, P.-O., 496 Hillman, G.,136 Hillman-Elies, A., 136 Hilmoe, R.,314(26), 315, 329, 330, 332, 334(14), 335, 376, 392(28), 393(28, 97), 394(28), 401(28), 426, 429(97), 521, 530, 534, 535(15), 538(15) Hilmoe, R. J., 746 Himmelhoch, S. R.,602 Hinds, J. A,, 12, 15(59), 16(59), 17(59) Hinsch, G.W., 605(313), 607 Hirahashi, T.,261 Hirarnaru, M.,215(32), 216, 227(32), 241, 247(123, 124) Hirchmann, R.,196 Hird, F.J. R.,96,142,692 Hirs, C. H. W.,148, 181, 514, 648, 650, 651, 653, 654, 655(56), 669(34), 670 (34), 671, 677, 678(107, 108, 109, 110), W(108, 109, 110), 691, 693 (169),712, 739 Hirschmann, R., 694,698 Hitchings, G.H., 76 Hiwatashi, O., 148 Ho, N. W. Y.,215(31), 216. 234, 237(31. lm),238(108), 321 Ho, P. P. K., 105, 108, 113(80), 114(78), 118(62) Hoagland, V. D.,Jr., 55, 56(83), 57(83). 58(83) Hoard, D.E., 307, 308 Hoare, D. G.. 112 Hodes, M.E., 283 Hodson, A. W., 419 Hoffman, M.,369 Hofrnan, K.. 698, 700(227) Hofmann, K.,196,699, 700(229, 230. 231. 232, 233), 701(233) Hofstee, B.H. J.,426,441 (99) Hogeboom. G.H., 126,132(24) Holcomh. D.N..706,707,729 Hollander, V. P.,450, 484, 485, 486. 487. 488, 489, 490 Holley, R. W.. 48. 215. 222(29). 227, 230 (93), 325 Hollingworth, B. R.,244(143), 249

8% ,Holly, F. W., 196,094,098 Holmes, E.,545 Holmqukt, N. D., 103 Holmstedt, B.,126 Holohan, P., 14 Holg, A., 218, 222(77), 223, 228, 754 Holzer, M.E.,573(131), 574 Honda, F.,368,309(54) Honjo, M., 320 Hood, L.,128(09), 129(68), 130 Hopkina, F. G.,49,133 Hopkinson, D. A.,477,480, 481,482,483, 484 Hoppe, W.,126 Horchrein, H., 802 Horecker, B., 53, 64(70), 361, 374, 377 (391, 378(6), 384(39), 387(39), 389 (39), 014, 017, 618(15), 019, 020, 021 (15,101, 022(15, 441, 623(45), 024(15, 45), 025, 026(51), 627, 630(42), 031 (30, 421, 632(64), 033(03, W ,634, 035, 036(80), 637(80), 639, 640(94, 95, 971, 641, 645(42, 63, 74, 94) Hori, A., 349 Horiniahi, H., 517,685,689 Horinouchi, R., 805(280),606 Horitau, H., 244(142), 246(101, 102), 249,

AUTHOR INDEX

Huang, S. M., 577,579(136) Hubbard, L.,140 Huberman, J. A.,257,258(31) Hubert, E.,620,030(43),031(43) Hubscher, G.,603 Hudson, P. B.,454, 459(23), 460(40), 466, 469(23), 481 Hughes, D. E., 87,95(10) Hughes, R. B., 19 Hulsmann, W.C.,614 Humeres, E.,438, 445(107), 440(107) Hummel, J. P., 627, 048, 732, 733, 739, 758, 777 Hunt, L.,727, 729 Hunt, V. M., 423 Hunter, A., 464, 465 Hunter, D., 418 Hunter, J. R.,55 Hurley, D., 772 Hurst, R.O.,309 Hurwitz, J., 53, 54(70), 177(11), 178, 185 (ll), 193(11), 250, 267(68), 268(68). 354 Husler, J., 418 Hvidt, A., 712,713

I

Ichihara, M., 429 250 Horiuchi, S., 374, 392, 393(95), 404(95), Igarashi, M.,450, 484(4), 485, 486, 487, 488, 489, 490 420 Igarashi, S., 354 Horiuchi, T., 374,420 Ihara, N., 804 Horn, A., 536, 640(23) Horne, R. N., 553, 556(80), 557(80), 558 Iida, S., 213,219(19), 221(19) (80, 981, 560(80), 588(98), 569(98), Ikehara, M.,757 573(98), 574030, 981, 575(98), 598 Ikenaka, T.,128(71), 129(71), 130 Il’in, V. S.,800(200), 801 (80,98, 178),599(80,98) Illingworth, B., 567, 568(115), 598(115), Hornichter, R. D., 697 800(116) Horowitz, B., 118 Imamura, H., 368,369(64) Horwitz, J. P.,327,433 Imanishi, A.. 094 Hoshino, O.,769 Imazawa, M., 229,246(97) Hotchkies, D.,426 Imsande, J., 30, 31 (551, 3365) Houck, J. C.,750,751(409) Housewright, R. D.,39(71), 43(71), 104 Imura, N., 756 Inagami, T.,196,656,683(01),785 Houssay, A. B.,605(315), 607 Inciardi, N. F.,634 Howard, A. J., 116 Ingbar, S. H., 000(218),601 Howell, B.A.,71(107, 1681,72 Inglis, N.I., 419,442061,443(26) Howell, L.G.,54(80), 55 Ingram, P.,348,351(75) Hsu, K.-C., 697,698(218) Ingram, V. M.,755 Hsu, L.L.,630 Ingwall, R.T.,734 Huang, N. J., 40

825

AUTHOR INDEX

Inoue, Y., 217,218 Inouye, K., 90,91(26) Ionescu, V., 605(302, 303), 607 Ipata, P. L., 58, 346, 348, 351(57), 352 (57), 750, 751(423)

Ire, M., 757 hie, M., 216, 217(41),

220(41), 229, 246(97, 163, 164, 165, 166), 247(172), 250, 536, 540(24), 648, 672, 758, 759, 761, 770, 778 Irie, N., 756 hie, S., 213, 215, 216(18, 271, 228(27) Irion, E., 108, 109(65), 111(65), 112(65), 113(65), 114(65), 115(65), 116(65), 118(65) Isernura, T., 694 Isherwood, F. A., 98,142 Ishida, Y., 51, 52(63), 53(63), 77(63) Ishii, F., 763 Ishii, J., 129 Ishii, K., 240, 244(122) Ishikawa, T., 230,247(101) Ishikura, H., 227, 230(89) Isselbacher, K. J., 422 Itagaki, K., 216 Itaya, K., 432 Ito, S., 421 Itoh, R., 343, 344, 345(32, 33), 348(33) Itoh, T., 213, 216(18) Ivemark, B., &I2 Iwai, K., 215(36), 216 Iwanaga, S., 128(65), 129 Iwanoga, C., 313 Iwert, M. E., 13 Iyer, N. T., 364

J Jnckson, L. J., 43, 44(102) Jackson, R. C., 110 Jackson, R. L., 654,655 Jacob, T. A,, 1%, 694 Jacobs, G., 281 Jacobs, S., 31, 33(62) Jacobson, E., 418(16), 419 Jacquemin-Sablon, A., 282, 304, 306(56a) Jagendorf, A., 641 Jago, M., 27, 34(29), 38(31), 39(29, 31), 40(31), 42(31), 43(31)

Jakobsson, S. V., 578 James, J., 565, 603(106)

James, S. P., 126 Jameson, E., 103 Janes, J. O., 118 Janssens, P. A., 600(216), 601 Jansz, H. S., 380,471 Jardetsky, C. D., 723 Jardetzky, O., 154, 155(18), 712, 723, 724(278, 281), 725, 732, 763, 784

Jasmin, G., 347 Jasrnin, R., 581(150), 582 Jayararn, H. N., 104, 117(30) Jencks, W. P., 85, 445 Jenkins, S. R., 698 Jenkins, W. T., 409 Jenner, H. D., 418 Jenrette, W. V., 291 Jensen, R. B., 643 Jeppessen, P. G. N., 234, 237(107), 238(107)

Jervis, H. R., 603 Jirgensons, B., 720,

721(311), 722, 723(319), 736 Johansson, S., 348 Johns, P. T., 556, 559(88), 561(88), 563(88), 575(103), 577(119, 120), 878(119), 579(88, 119, 1201, 580(103, 119, 120), 581(119, 120), 587(103, 119, 120), 590(103, 134), 592(103, 134), 594(103), 595(88), 597(88) Johnson, B., 379(54), 380, 389(54), 402(54), 405(54) Johnson, D. V., 50 Johnson, E. J., 639 Johnson, F. H., 59 Johnson, K. D., 573(131), 574 Johnson, L. J., 12 Johnson, L. N., 196, 656,683(61), 785 Johnson, M. K., 539 Johnson, P. H., 315, 328(31)

Johnson, P. L., 603 Johnson, R. N., 59 Johnson, W., 581(150), 582 Johnston, R. E., 368 Jonadet, M., 605(294), 606(335), 607, 608 Jonek. J., 600(201), 601 Jones, D. S., 128(64), 129 Jones, G.. 547,597,604(48) Jones, G. H., 783 Jones, H. W., 605(317), 607 Jones, L., 105

826

AUTHOR INDEX

Jones, L. C., 569 Jones, W., 48,649 Jorgensen, S. E., 255, 288 Jori, G., 682, 683(131) Jos, J., 803 Josan, V., 51, 76(60), 7760) Josefsson, L., 648,696, 750,751 (420) Joshua, H., 698 Josse, J., 178, 501, 502, 503(13), 504(13), 505(13), 506(13), 509(13, 301, 510(12, 13,30), 511(30), 512,513(34), 514(34), 515, 516(45), 517(45), 518(12, 45). 519, 520(12), 521(12), 523(54), 521(54), 525(54), 527(54), 536, 537(!22), 538(22) Jovin, T. M.,258 Joyce, B. K., 451,771

Juchnowicz, E., 302 Jung, G., 451 Junge, J., 433

K Kadner, R. J., 420 Kaesberg, P., 707 Kagan, H. M., 82 Kaiser, A. D., 178, 254, 264 Kakimoto, Y.,144 Kakinuma, A., 354 Kalbener, P. P., 643 Kalckar, H. M., 51,76(61) Kaldor, G., 70 Kaletta, U., 316 Kalkstein, A., 28(38), 30, 34(38), 38(77), 39(38), 40(38, 77), 41(77), 44(38, 77), w38, 77)

Kallen, R. G., 85 Kallio, R. E., 14, 18(77), 19(77) Kalnitsky, G., 048, 691, 732, 777 Kaltwasser, H., 13 Kalyankar, G. D., 76 Kaminski, Z. C., 44 Kanai, Y., 320 Kanazawa, A., 144 Kaney, A. R., 639 Kang, A. H.. 128(72), 129(72), 130 Kaplan, A., 5, 12(24) Kaplan, J. C., 258, 289(32) Kaplan, N. O., 74, 207,353,623 Kappas, A., 344 Kara-Murza, 9. N., 394,534

Karnovsky, M. J., 602 Kartha, G., 656, 657(60) Karu, A. E., 540 Kasai, H., 221 Kasai, K., 219, 231, 247(105) Kashnig, D. M., 549 Kasper, C. B., 549 Katchalski, E., 12, 392,

393(100),

394(100), 395(100) Kato, H., 128(65), 129 Kato, I., 196,695, 705 Katz, A., 181 Katz, I., 394 Katz, S. A., 5, 19

Kauffman, D. L., 380 Kaufman, J., 51, 52(67), 59(67), 60(67), 62(67), 63(67)

Kaufmann, W., 108, 109(65), 111(65), 112(65), 113(65), 114(65), 115(65), 116(65), 118(05) Kautzch, K., 130 Kauzmann, W., 694,706 Kawahara, K., 509, 710,728(287) Kawai, Y., 149 Kawaee, S., 216 Kawaahi, S., 313 Kawashima, H., 221 Kay, H. D., 418(14), 419, 420, 421(14. 351, 428(34, 351, 432(35), 440(51), 443(14) Kay, L. D., 641 Kazenko, A., 292 Keir, H. M., 245(146), 249

Keith, J., 364 Kellen, J., 548, 605(52) Kellenberger, E., 501 Keller, E. B., 319 Keller, P. J., 293, 651 Kelly, D. M., 883 Kelly, R. B., 255, 257(24), 258 Kelly, T. J., Jr., 263 Kemp, C. M., 763 Kemp, R. G., 309 Kendall, E. C., 133 KendrickJones, J., 72 Kenkare, U. W.. 686, 699(153), 703(153) Kenner, G. W.,128(64), 129 Kent, T. H., 803 Kettman, M. S., 740 Keppie, J., 104, 116(38), 119(38)

AUTHOR INDEX

827

Klecekowski, K., 17 Kerkhoff, J. F., 426 Klee, C. B.,332 Kern, F.,15, 17(86) Klee, W. A., 669, 670(74), 671, 672, 677, Kerr, I. M.,252 678(1031,727,730,749 Kersten, W., 581(149), 582 Kleiber, M.,600(196), 601 Kessel, R.G.,606(325),608 Klein, A. J., 136 KBedy, F.,85,404,410(135),444 Klein, W.,349 Khoarovchahi, H., 605(293), 607 Khorana, H. G.,188, 189, 299, 308, 309, Kleinschuster, J. J.,616 310, 314(27, 28), 316, 318, 319. Klenow, H.,258 320(17), 321(41), 322(28), 326. Klett, R. P., 256 328(17), 330, 331(15), 332(15), Klingman, J. D.,93 Klotz, I. M.,697, 698(214, 216), 720(214) 334(15), 356, 364 Kluetsch, K., 602 Khouvine, Y., 64 Kluge, H., 50, 70,73(155) Kibrick, A. C.,131 Iilunker, G.,18, lQ(104) Kida, M.,51,52(63), 53(63), 77(63) Knight, C. A., 177(12, 14, 151, 178, Kidd, J. G.,102, 119 185(12, 14, 151, 186(12), 755 Kiesow, L.,609 Knights, E.M., Jr., 77 Kiessling, K., 500 Knitsch, K. W., 581(151, 152), 582, Kilsheimer, G.S.,465,600(211),601 594( 152) Kim, J. H.,110 Knorre, D.G.,333 Kim, S.,692 Knox, J. R.,163, 656, 657(62), 658(62), Kimio, O.,148 666(62), 667(62), 724(62), 785 Kimura, F.,227,240 Knoa, R., 24,38(83),39 Kimura, K., 536,540(24) KO, S.H., 404,410(135) Kind, P.R.N., 433 King, E. J., 418, 419, 423, 433(30), Kobashi, K., 14(93), 16, 17(90, 93, 94), 20(94) 435(30), 436, 440(159), 447, 451, 469, Kobayashi, M., 227 477(18),478,479,480 Koch, A. L.,49,55(33) King, J. A,. 131 King, T.P.,650,651,669(28, 30), 670(28. Koch, F. C.,49 Kochman, M.,430,442(120) 30),889 Koeppe, R.E.,134 Kinne, R.,420, 421(50) Koerner, J. F., 254, 255(19), 268, 291, Kinne-Saffran, E.,420,421(50) 308, 317 Kinoshita, S.,124, 138(9) Kofoed. J. A.,605(315),607 Kirby, W.M.M., 24 Kogan, G.L.,394 Kirchheimer, W.F.,104 Kogl. F.,136 Kirk, M.,643 Kogut, M.,28(36),30,31(36),32(36) Kirkwood, J. G.,723 Kohn, J., 338 Kirsch, J. F.,59,61(113) Koide, H.,544, 548 Kirschbaum, J., 113, 114(77) Koike, T.,217,231,232,233(103. 104) Kirschner, K., 71 Komai, T.,16,17(94), 20(94) Kishi. K., 605(309),607 Komkova, A. I., 398 Kislina, D.S.,321 Konarev, V. G.,750,751(408) Kitagawa. S.,72 Konchetkov, N.K.. 249 Kitai, R.,129 Konev, S. V..10 Kitasato, T.,429 Kizer. D. E., 65, 66(128), 68(128), Konijn, T.M.,366 Kontornichalou, P., 29(42),30,40(40) 69(128), 71(151, 167,la), 72 Kool. M.,605(290), 607 Klain, G.J., 600(u)5), 601 Kopac, M.J., 600(217),601 Klebanova, L. M.,249

828

AUTHOR INDEX

Kopecka, H., 279 Kopko, F., 14 Kornberg, A., 178, 253, 255, Z56(22, 26, 271, 257(24, 27), 258(22, 29, 31), 315, 316,330,501,529,539,572

Kornberg, H. L., 643 Kornberg, S. R., 521 Kornfeld, D. S., goO(196), 601 Koshland, D. E., 112, 430, 445, 446, 454, 471, 746

Kosinski, P., 651 Koszalka, T. R., 275,281 Koudstaal, J., 348, 605(320), 607 Kovacs, A. L., 71 Kovd, E. Z., 609 Kowalczyk, L. S., 327 Kowal~ky,A., 724 Kowlessar, 0. D., 281 Koyama, J., 218 Kozloff, L. M.,259, 282 Kozlowska, K., 603 Krainick, H. G., 140 Kramer, M.,30 Kramer, P. I., 103 Kratowich, N.,631 Krause, S., 708 Kraushaar, A., 44 Krausz, L.M., 706 Krawczynski, J., 77 Krebs, E. G., 48 Krebs, H. A., 103, 106(15), 107(15), 632, 633(72), 634(72)

Krenitsky, T. A., 717, 731 Kresheck, G. C., 741 Kretovich, W. L.,105(50), 106 Krisch, K., 581(148, 149), 582 Kdshnan, P. S., 51, 76(60), 77(60, 189) Krishnamamy, P. R., 124, 126(6, 71, 137(6, 71, 150

Kropf, R. B., 697,6!38(215) Krulwich, T. A., 620, 630(42), 631(421, 645(42)

Krumey, F., 451, 501 Kuehl. F. A., Jr., 369 Kuentzel, H., 755 Kuff, E. L., 126, 13204) Kuhnel, W., 606(326), 608 Kuhns, J. G., 8w Kumagai, K., 244(142), 249 Kumar, S., 51,76(60), 77(60, 189)

Kunitz, M.,291, 292(18), 529, 530(3), 531, 534(3), 535(3), 649, 749

Kunze, H. E., 12, 15(59), 16(59), 17(59) Kuo, M., 376 KUO,M.-H., 497 Kupke, D. W., 705,706(248), 707,709 Kurihara, K., 517 Kurihara, T., 364,385 Kuriyama, Y., 216, 218 Kurnick, N., 289 Kurtz, H. M., 103 Kusaba, F., 246(159), 249 Kusaka, T., 612 Kushmerick, M. J., 72 Kushnarev, V., 375 Kushner, D. J., 24, 46(16) Kushner, S. R., 258,269(32), 270(76) Kutcher, W., 450,451,453(3), 455 Kuwabara, S., 27, 28(37), 30, 31(37), 34(37), 38(37, 80),39(32, 371, 43(37) Kuwano, M., 243, 245(132) Kycia, J. H., 677,678(108), 680(108) Kvam, D. C., 645

1 Laboureur, P., 121 Lacks, S., 253, 261(10) Laden, K., 144 Laga, E., 437 Lagemtedt, S., 648, 750,751(420) Laidler, K. J., 19(109), 20,444, 773 Laiken, S. L., 714 Lamar, C., Jr., 93, 95(28) Lambert, S. M., 523,535 Lsmden, M. P., 780 Lampen, J. O., 24 Lamy, M.,603 Landin, L. M., 117, 119, 120 Landing, B. H., 496 Lane, B. G., 313, 314(6, 7). 320, 776 Lang, K., 451 Langdon, R. G., 553,600(74) Lange. K., 602 Langman, M. J. S., 422 Lanza, T., 698 Lapanje, S., 509,710, 726(267) Lardy, H. A., 540,546,568,645 Lareau, J., 634 Lanen, M., 535 Larsen, S., 156

AUTHOR INDEX

Larson, A. D., 14, 18(77), 19(77) Larsson, S.,605(297),607 Laskowski, M.,Jr., 301 Laskowski, M.,Sr., 153, 154(1), 155(9), 156(8), 174(12), 177(10, 13), 178, 179, 185(10, 13), 186(10, 13, 22, 59), 193(22), 251, 260, 272, 281, 284, 285 (1, 5), 290, 291(10), 292, 297(10, 26, 27, a),299(10, 19, 20, 21), 301(21), 302(10), 303(6, lo), 307. 308(10, 26, 27, 30), 309(42), 3 W 6 . 7, 41), 313, 31403, 25, 29), 315(16), 316(8, 11, 12), 317(19), 318(30, 35). 319(35, 40), 320(16, 19, 20. 30, 35), 322(20, 251, 323(29, 50), 324, 325, 326(13, 29), 328(12, 19, 20, 31), 329, 333, 334(32, 34), 342, 350(23), 352 (23), 476, 717 Laster, L., 420, 603 Laszlo, J., 281 Lathe, G.H., 600(199),601 Latner, A. L., 419 Laurila, U.-R., 290, 299, 308, 3O9(42). 314, 316(11), 476 Laursen, R. A., 757 Laval, J., 282 Lavallee, W.F.,467 Law, A. S.,117, 119(86) Lawley, P. D.,265 Lazarus, L. H., 248(176), 250 Lazarus, S.S.,604 Lazdunski, C., 378, 379(43, 52), 380. 384(43), 385, 386, 392, 393(94), 394(94), 395(94, 106), 401(52), 403, 404(94, 106), 405, 406, 408(94), 409(94, 106), 413 Lazdunski, M.. 378, 379(43, 52), 380, 384(43), 385, 386, 392, 393(94). 394(94), 395(94, 106), 401(52), 403(52), 404(94, l06), 405, 406(138). 408(94), 409(94, lM),413(138), 425. 428, 435(91), 436, 437, 438(162), 439(91) , 442( 107), 444( 162), 445(91 162) Lea, M. A., 600(191, 192), 601, 603(191. 192),604(191, 192) Leach, S.J., 713 Leber, P. D.,516 Leblanc, D.,378 Lecocq, J., 408

829 Leder, P., 230 Lederberg, J., 501 Ledoux, L., 690 Lee, B., 163, 656, 657(62), 658(62), 666(62), 667(62), 724(62) Lee, C.W.,553,578(71),599(71) Lee, C.Y., 301 Lee, E.R.,440 Lee, J. F.,364,366(29) Lee, M. B., 103, 106 Lee, Y.-P., 48(20), 49, 50(20), 64, 65, 66(122, 130),67(122),69(130), 70(130) Lees, E.M.,105(51), 106 Legait, E.,605(289), 607 Legait, H., 605(289), 607 Legler, B., 561, 568(101), 580(101), 602(101) Lehman. I. R., 252,253,254,255,256(25), 257, 259, 260(7), 261, 266, 267(62), 272,282,285(3),315,316, 330 Leibach, 94, 95(35), 96(35), 97 Leibson, L. G.,600(207), 601 Leidy, G.,757 Leive, L., 374 Lennox, F. G.,49 Lentz, T. L., 609 Leonard, N. J., 51,52(69), 62(69), 767 Leonhardi, G.,144 LePage, G.A.,59,60(112), 61(112) LePecq, J. B.,260, 282 Lepoutre, L., 750, 751(405) Le Quesne, W.J., 126,133(25) Lerique, J., 72 Lerner, F.A.,457 Lesca, P.,282 Le Talaer, J.-Y., 282 Leuthardt, F.,50, 125,612 Leuthold, E.,422 Leienberg, B.,14,94,95,110 Levene, P.A., 418 I,evin, D.H., 215(37), 216 Levin. S.J., 343,351(29) Levine, D., 377,392(30),393(30),394(30), 398. 408(30), 410(30) Levine, R., 50 Levinthal. C., 374, 375, 376(4), 377(4), 378(4), 379(36), 380(38), 382(65). 384(38), 385(20, 38, 69). 387(4). 389(4), 392(4),393(4), 394(4), 395(4). 402(36), 406(4), 429

830

AUTHOR INDEX

Levintow, L., SO, 82(6), 84(6), 9303, 101, 107(2), 110(2), 126, 132(24) Levitt, M., 157 Levy, H.M., 516 Lewis, H.B.,149 Li, c. 75 Li, C. H., 182,711 Li, L.-K., 716 Libonati, M ., 242,245 ( 130) Lichtenstein, N.,103, 104, 126 Lie, S. Y.,10s Lieben, F., 14 Lielausis, A.,501 Lifson, S.,157 Lightstone, P.J., 434 Liguori, G.,605(299), 607 Limetti, M.,699,700(230) Lin, M.C.,673,688,690(163) Lindahl, P.E.,500 Lindahl, T.,330 Lindberg, U.,291, 292, 293, 296, 297(34), 298, 29!N22, 231, 300 Linderstrom-Lang, R., 712 Lindqvist, C., 31,39(59), 42(59) Lindstrom, E. B., 24, 29(43), 30, 31(43), 32(43), 33(43), 34(43), 39(4, 431, 40(43), 41(43), 42(43), 43(43), 46(43) Lineweaver, H.,576,577(135) Link, T.P.,682,690(133) Linn, S.,263,264-32) Linneweh, F.,605(279), 606 Lipmann, F., 126, 377, 378(35), 380, 396(35), 398(35), 405(35), 425 Lipscomb, W., 90,91(2l) Lipsett, M.N.,218 Limwski, J., 346,351(54),352(54) Lister, A. J., 14,17(76), 19(76) Little, C.,623 Little, J., 696 Little, J. W., 254,267 Litwin, J., 548,605(51,272, 3001,606,607 Liu, T.-Y., 292, 293(35), 295(35), 296(35), 297(35) Live, T.R.,304,305(56), 306(56a) Llamas, R.,772 Lo, T.,342,351(27), 352(27) Lobitz, W.C.,605(310), 607 Loeb, G.I., 769 Lohr, G.W.,605(279), 606 Logan, D.M., 245(147), 249

c.,

Logue, A. D.,738 Lohman, C. L., 66(145a), 67, 68(145a), 70(145a) Loke, J., 375 London, M.,466 Lopez, J. A.,432 Lora-Tamayo, M.,436,437(163), 441(163) Loring, H.S.,353 Losada, M.,643 Losert, W., 369 Loveless, J. L., 120 Lovig, C.A., 69,71(151) Lowe, M.J., 722 Lowenstein, J. M.,65, 66(129, 1311, 68(129, 131, 138, 1411, 69(131, 141), 70(129), 72(141), 73(131, 138) Lowry, D. L., 26,43,44(102) Lowry, 0. H., 432,433 Lubmann, A. J., 44 Lucas, Z.J., 255,256(22), 258(22) Ludowieg, J., 605(304), 607 Lueck, J. D.,547, 549(42, 43). 559(42, 4 3 , 567(42, 43), 568(42, 43), 570(42, 431, 573(43), 574(43), 576(42, 431, 579(42,43,143),582,585(43), 587(42, 4 3 , 590(42, 431, 595(42, 43), 599(42) Luew, H., 451 Luffman,J. E.,422,481,482(83), 483 Luft, D.,600(193),601 Luisada-Opper, A. V.,697.698(219) Lukton, A.,391 Lund, G.,353 Lundblad, R.L., 364 Lundgren, E., 454 Luppis, B., 614, 615, 616(22), 618(15), 621(15, 161, 622(15, 44), 624(15), 627(22) Lutsenko, M. T., 602 Lutwak-Mann, C.,49 Lygre, D. G.,547, 550(46), 551(46, 47), 553(46), 559(46), 561, 565, 567(46), 568(46, 114), 571, 572, 573(46), 574. 575(103, 118, 132), 577(46), 578(46), 579(46, 118), 580(46, 103, 1181, 581 ( 118), 585(46), 586 (1 181, 587(46, 103, 118),590(46. 103, 118), 592(103), 593(118), 594(103), 598(118), 599 (118),603(46,114) Lynn, K. R., 3,4, 18,19,20(12)

831

AUTHOR INDEX

Lyster, R. L. J., 426 Lyubimova, M . N., 66(146), 67, 68(146)

Mager, J., 66(143), 67, 68(143), 69(143) Maguire, M. H., 55, 56(86), 57(86), 63

M

Mahler, I., 258, 269(32) Mahy, B. W. J., 120 Mainiteri, L., 605(274), 606 Makarewicz, W., 49, 50, 64, 65(126), 66

Ma, P. F., 55,56(96), 57 Macbeth, G., 641, 642(100) McBride, T. J., 40 McBride, R. A., 600(210), 601 McCallum, G. H., 446 McCaman, M. W., 71 McCaman, R. E., 71 McCann, W. P., 602 McCarthy, J. R., Jr., 59 McCarty, M., 291 McClure, W. O., 90, 91(25) McCoy, T. A., 117 McCraw, E. F., 596,597(169) MacDonald, X., 491 McDonald, M. R., 649 McElroy, W. D., 51 McEvoy, F., 55 Macfarlane, M. G., 420 McGeeney, K., 3, 426 McGilvery. R. W., 612, 613, 616(4), 617, 618, 631, 644, 645(25)

McGrath, T., 86 MacHattie, L. A., 279 McHugh. R., 466 MacIntyre, R. J., 498 McKenzie, B. F., 133 Mackey. L. D., 734 McLaren, A. D., 15, 19 McLennan, B. D., 313, 314(6, 71, 353. 776 MacLeod, R. M., 605(296), 607 McLeod, S., 77 McManus, D. K., 49 McMeekin, T. L., 706 McMillan, F. H., 131 Macon, J. B., 51, 52(67), 53(67), 59(67). 60(67), 62(67), 63(67)

Macpherson, H. T., 128 McQuarrie. E. B., 44 Madison, J. T.. 48, 215. 222(29), 227. 230(93), 325

Madras, B. K., 118 Madsen. N. B., 48 Maeda, S., 603 Maengwyn-Davies, G. D., 558(100). 559 Magana-Plaza, I., 14

(861, 64(86)

(126), 67(126), 68(126), 69(126), 70 ( 126)

Malamed, S., 557, 559, 560(92) Malamy, M., 361, 374, 377(39), 378(6), 384(39), 387(39), 389(39)

Malato, M., 606(322), 608 Malet, P., 600 Maley, F., 651 Maley, G. F., 568 Malor, R., 550, 556(67) Malseh, L., 451 Malveaux, F. J., 451, 498(12a) Malysheva, M. K., 50, 55, 66(54), 67 (541, 68(54)

Mamiya, G., 3, 9, 10, 13 Mancini, R. E., 605(287), 606 Mandel, L. R., 369 Mandel, M., 724 Mandel, P., 365 Mandelkern, L., 505 Manery, J. F., 50 Mangiarotti, G., 375 Mangiarotti, M., 616, 618(27), 619 Manhouri, H., 432 Manjeshwar, R., 593(163), 594 Manjula, B. N., 731 Mann, T., 457,500 Manners, D. J., 544 Manning, G. B., 104, 117(31) Manning, L. R., 82 Manson, E. E. D., 43, 44(98) Mansoor, M., 76 Mansour. T. E., 48 Marble, S. J.. 85, 94(12), 97(12) Marcker, K. A.. 227, 232(92), 237(92), 755

Marchiori. F., 671, 695, 699(242, 243, 244). 701(235. 236, 237, 238, 239, 240, 241, 242, 243, 244). 702 Marcus. F.. 620, 630(43), 631(43) Mardashev, S. R., 105, 106 Marfey, P. S., 696 Marino, C., 548, 605(,53)

832

AUTHOR INDEX

Markham, R., 316, 321(41), 329, 363, 476, 746

Markley, J. L., 154, 155(18), 724 Markley, K., 14,15(85) Marquardt, H., 121 Marquisee, M., 48, 215, 222(29) Marshall, E. K., Jr., 148 Marshall, K., 706 Martin, A. J. P., 650 Martin, S. P., 602 Martin-Esteve, J., 750, 751(411) Martland, M., 418(15), 419, 450,477 Marullo, N., 291, 304(24) Maruo, B., 240, 244(118), 240 Maruyama, H., 363 Maruyama, Y., 148, 149(134) Marvin, D. A., 158 Marzotto, A., 680, 682, 683(131), 699, 701 (236, 241)

Masamune, Y., 253, 25403) Mashburn, L. T., 102, 104, 105, 106, 107, 109, 110(39), 114, 117(39). 119, 120 (41, 87) Mason, H. L., 133 Mason, H. S., 572 Mason, S. F., 51, 53(73)

Mason, T. W., 366 Maasart, L., 772 Massey, K. L., 366, 368(38), 369(38) Massoulie, J., 333, 755 Mastarlerz, P., 430, 442(120) Mathews, H., 600(219), 601 Mathias, A . P., 749, 763, 771, 773, 779, 780

Mathies, J. C., 419, 423(21), 427(21) Mathog, R. H., 245(146), 249 Matlina, E. Sh., 66(146), 67, 68(146) Matsubara, H., 317 Matsui, K., 245(148), zQ9 Matsuo, I., 10 Mattila, S., 468 Mauck, J., 340 Maver, M. E., 281, 334 Maximilian, S., 605(277), 606 Mayer, J., 600 Mayer, K., 451 Mayer, R. L., 758 Mayr, O., 103 Mazhul, V. M., 10 Mazur, R. H., 126

Meadows, D. H., 154, 712, 724(278), 732, 763, 784

Meadway, R. J., 31, 33(63, 641, 35(63,

W, 37, 4 2 ( W Meagher, J. G., 421 Mebs, D., 313 Medhat, P., 53 Medicus, R., 630 Medigreceanu, F., 418 Mehler, A., 640(95), 641 Mehta, S. L., 14, 17(75) Meijer, A. E. F. H., 489 Meisel, E., 421 Meisenheimer, M., 451 Meister, A., 80, 82(6), 84, 85(3), 93(6), 94(12), 96(32), 97(12), 101, 102, 106, 107(2, 52), 110(2), 118, 120(2), 124, 125, 126(6, 7), 127(13), 132(24), 133, 134(8), 135, 136(102), 137(6, 71, 141 (13), 142, 143(13, 1201, 144(13), 145 (13), 146, 147(127, 1291, 150, 151(154) Mekanik, G., 605(296), 607 Melani, F., 422 Melchior, W. B., 697, 698(222) Melgar, E., 265, 279, 302 Melnick, I., 110 Melo, A., 339 Melville, J., 132 Melzer, M. S., 281 Menahan, L. A., 367, 368(45), 626 Mendelsohn, S., 268 Mendicino, J., 66, 68(139), 615, 630(20), 631 Menozzi, A., 124 Menten, M. L., 433 Mentendiek, M. A., 645 Merrifield, R. B., 196, 199, 673, 694, 698 Merrill, S. H., 48, 215, 222(29) Meselson. M.. 263, 264 Mesrobeanu, L., 373 Messer, M., 94, 125, 130, 139, 140(10, 751, 141(116) Methfessel, F., 136 Meusers, P. J., 605(295), 607 Meyerhof, O., 419, 472 Michelson, A. M., 217, 230, 333, 755, 757 Micu, D., 605(277), 606 Mihailescu, E.. 605(277), 606 Miles, P. L., 431 Milkowski, J. D., 698

AUTHOR INDEX

833

Molemans, F., 750, 751(422) (22), 193(22), 291, 315, 318(30), 320 Moller, K. M., 750, 751(403) Money, C., 217 (30), 326 Monny, C., 757 Milikin, E. B., 108, 112(80), 114 Miller, A., 96, 125, 142(22) Monod, J.. 387, 389(77) Monroe, J. F., 602 Miller, D. L., 534 Montague, M. D., 747 Miller, D., 266 Montalvo, J. G., Jr., 12 Miller, J. A,, 136 Montibeller, J., 699, 700(230) Miller, R. J., 18 Monty, K. J., 6 Milner, W. A., 602 Milstein, C., 380, 397, 398(119, 120), 405 Moog, F., 421 Moor, S., 220 (120), 425 Moore, B. W., 484 Milstein, S.W., 750, 751(401) Moore, G. E., 132 Milton, J. M.. 11 Moore, L., 254 Minami, I., 244(141), 249 Minato, S., 51, 53(75), 73(179), 74(75). Moore, S., 129, 148, 181, 292, 293(35, 38, 39, 39a), 295(35, 39), 296(35), 297 213 (35, 37), 364, 512, 513(35, 37, 381, Mira, E., 605(292), 607 650, 653(22, 25), 654, 655(55), 669 Mirsky, A. E., 285 (25), 670(25), 673, 679, 682, 683(130), Mirsky, I. A., 545, 580(17), 600(580) 686, 688( 1601, 690( 158, 159, 161, 1631, Mirrabekova, A. D., 321 691, 744 Mitchell, H. K., 49 Moor, E. D., 118 Mitchell, P., 374, 609 Mitsuhashi, S., 29(44), 30, 31(44), 34 Mora, P. T., 758 (441, 38(44), 39(44), 40(44), 41(44), MorBvek, L.. 155, 163(38), 179, 182(23), 184, 196(53), 197(53) 42(44), 43(44) Mitsui, A., 343, 344, 345(32, 33), 348(33) Morgan, E. J., 49 Morgenstern, S. W., 423 Mitsui, Y., 785 Morita, R. Y., 677, 678(117), 740 Miura, K., 216 Moroder, L., 699(242, 243, 2441, 701(235, Miura, Y., 605(310), 607, 772 237, 238, 239, 240, 241, 242, 243, 2441, Miyaki. M., 51, 53(75), 7405) 702, 721 Miyaraki, M., 226, 227, 230(90), 232(90), Morrill, G. A., 651, 770 236(90), 237(90), 756 Morris, S. J., 656 Miyoshi, Y., 432 Mizuno, D., 244(144, 145). 249, 358, 363 Morton, M., 550, 556(67) Morton, R. A., 296 (111, 374, 420 Morton, R. K., 397, 406, 420, 423, 425, Mizuno, Y., 320 427, 429, 431, 434, 436, 437, 439(123). Moat, A. G., 38(78), 39, 40(78), 41(78), 440(90, 1231, 441, 442(113), 443(100, 42(78), 44 113), 549 Moav, B., 130 Moe, 0. A., 530, 534, 535(9), 536(9), 537 Moskvitina, T. A., 653 Moss, D. W., 419(32), 420, 424, 425, 428, (91, 538(9), 539 429, 431, 433(30), 435(30), 437(116), Moffatt, J. G., 783 441. 442( 174), 457, 469 Moffitt, W., 505,722 Moss, R. J., 747, 755 Mohan, R. R., 14 Mothes, K., 103 Mohn, G., 140 Mohnike, G., 581(151, 152). 582, 594(152) Motohashi, N., 125, 138(14, 15), 139(14) Motzok, I., 435, 436(152) Mohr, S. C., 222(78), 223, 233(78) Moudrianakis, E. N., 540 Mokrasch, L. C., 612, 616, 645(25) Mross, G. A., 128(63), 129 Moldoveanu, N., 72 Mikulski, A. J., 154, 174(12), 179, 186

834

AUTHOR INDEX

Narita, K., 129 Naylor, A. W., 2, 3, 5(15), 6, 9(34) Naylor, R., 321 Nayudu, P. R. V., 431 ( 10) Neale, F. C., 428, 443 Mulczyk, M., 148, 149(135) Neale, S., 390 Mundry, K. W., 750,751(414) Nechiporenko, Z. Yu., 50, 66(51), 67(51) Munier, R.L., 389, 390 Neet, K. E., 445 Munske, K., 369 Negi, T., 540 Munts, J. A., 66,68(139) Negrea, F., 602 Munro, J. L., 268 Neidle, A., 80 Murachi, R., 149 Murakami, M., 227, 230(W, 232(90), Neims, A. H., 11 Nekhorocheff, J., 72 236(90), 237(90) Nelson, C. A., 732, 733 Murao, K., 227, 230(89, 941,757 Nelson, N. S., 13 Murdock, A. L., 680 Nelson, P., 800(194), 601 Murison, G. L., 50 Murphy, P. M., 54(82), 55, 56(82, 91), Nemeth, A. M., 600(190), 601 Nesbett, F. B., 557 57 (82) Murray, A. W., 64,66(124), 66(148), 67 Nestle, M.,252 Nestor, L., 130, 141(76) (124), 68(148), 348, 351(80) Neu, H.-C., 24, 29(8), 31, 338, 339(9), 340, Mustafa, S. J., 49 349(9, 131, 350(9, 131, 356, 357(9), Mycek, M. J., 80 359(9), 360(9), 361, 362(9, 101, 374, Myer, Y. P., 769

Mueller, E., 17 Mukadda, A. J., 640 Mukai, J.-I., 177(10), 178, 185(10), 186

378, 639

N Nachmansohn, D., 397, 409(118), 471, 472 Nagana, B., 540 Nagano, M., 802 Naidoo, D., 347 Naik, M. S., 14, 17(75) Nair, K. G., 366, 367(42), 388(42), 369 (42)

Nakajima, Y., 605(309), 607 Nakashima, K., 617, 624, 625, 626(51), 631(30)

Nakamura, K., 207,648 Nakamura, M., 604 Nakamura, S., 246(154), 249 Nakanishi, K., 51, 53(75), 74(75), 213, 341, 350(21)

Nakano, E., 758 Nakao, Y., 246(155, 156, 157), 249 Nakatsu, K., 347 Nakayama, H. S.,269, 270(75) Nally, R., 13, 14(69), 17(69) Nandy, K., 347,606(333), 6Cb3 Naoi-Tada, M., 223 Nnra, S., 66(144), 67, 68(144), 69(136), 70(136)

Narayanan, R., 605(306), 607

Neuberger, A., 132 Neuhaus, 0. W., 423, 424(76), 435(76), 436(76), 437(76), 443(76), 444(76)

Neuman, R. A., 117 Neumann, H., 392, 393(100), 394(100), 395(100, 1091, 396, 398, 406(124), 407, 430, 452, 453, 691, 693(182) Neumann, R. P., 682, 683 Neumann, W. F., 421 Neurath, A., 90,91(25), 291 Neurath, H., 293, 380, 651 Neville, D. M., Jr., 344 Newmark, M. Z., 497 Newsholme, E. A., 633,635,644 Newton, G. G. F., 27, 34(29), 39(29, 32) Nezgovorova, L. A., 773 Nguyen-van-Thoai, 420, 423 Nichol, L. W.. 6, 7, 9, 12 Niebroj, T. K., 609 Nielsen, S. O., 712 Nigam, V. N., 454, 457, 458, 459(38), 472 (25). 473, 475(25) Nihei, T., 319, 332 Nikai, M., 244(141), 249 Nikiforuk, G., 65, 66(127), 67(127), 70 (135)

835

AUTHOR INDEX

Nikolaev, A. Y., 103,105 Nikolskaya, I. I.,316,318,321 Nilsson, T.,605(297), 607 Ning Kwan, C.,243,245(132) Nirenberg, M.,230 Nishikawa, A. H., 677, 678(117), 740 Nishimoto, T.,269, 270(75) Nishimura, H., 218 Nishimura, S., 226, 227(86), 228(86), 230 (86,89, 94), 239, 240, 244(112, 118), 757, 772 Nisselbaum, J. S., 345 Niwaguchi, T.,125, 138(14, 15, 16), 139 (14) Noda, H., 600(214), 601 Nodes, J. T.,750,751(419) Noguchi, J., 243 Nohle, E. G.,14 Noltenius, H.,602 Nomoto, M.,231, 247(105) Nomura, M.,148, 149(134) Noonan. M.,54(82). 55, 56(82), 57(82) Nord, F. F.,427 Nordlie, R. C.,408, 540, 544, 546, 547, 549(10, 37, 40, 42, 43, 451, 550(46), 551(46), 552, 553(5, 37, 46), 555, 556 (80,82). 557(80, 89), 558(37, 41, 45. 89, 93, 98). 559(9, 10,42, 43, 46, 88. 89), 560(80, 93,94,951,561(9. 10.88). 563(9, 10, 88, 89), 564, 565, 566, 567 (10,:3 37, 40, 41, 42,43, 45, 46), 568 (37,40, 41, 42, 43, 45, 46, 93, 94, 95. 98. 101, 114). 669(98), 570(41. 42,431, 571,572, 573(9, 37,40. 41,46, 89,98). 574(9, 10, 37, 40, 41, 43. 45, 80, 98). 575(37, 41, 98, 103, 118, 132),576(37. 40, 41, 42, 43, 45), 577(37, 40, 41, 45, 46, 119, 1201, 578(46, 66, 119), 579 (9,10, 40, 41, 42, 43, 46, 66, 82, 88. 118, 119, 120, 126, 143), 580(10, 37. 41. 46, 89. 95, 97. 101, 103, 104. 118. 119, 120, 141), 581(37, 118, 119. 120). 582, 583, 584. 585(37, 40, 41. 43, 45. 46). 586(10, 118), 587(37, 40, 41, 42. 43. 45, 46, 103, 104. 118, 119. 120. 141). 590(37, 41, 42, 43, 45, 46. 103. 118. 134), 591(104), 592(9. 10, 103). 593(9. 118). 594(9, 48, 103, 104, 141). 595(9. 10. 42, 43. 88, 104), 596(9, 10, 40), 597(37, 40, 41, 881, 598(40, 80,

93, 94, 95, 98, 99, 118, 121, 175, 178), 599(9, 10, 40, 80, 93, 95, 98, 99, 118), 600(37), 602(45, 101), 603(46, 1141, 645 Nordstrom, K., 24, 31, 39(59), 42(59) Norris, E.R.,51,53(62) North, A. C.T., 116,119(84), 156 Nossal, G.W.,332,338 Nossal, N. G.,241, 244(127), 255, 361, 374 Notkins, A. L.,120 Novelli, G. D., 623 Novick, R. P.,24,28,40,41(89),43(89) Novikoff, A. B.,485 Novikova, N. V.,436 Novitskaya, V. A., 436 Xovoa, W. B.,580(146), 581(146), 582, 602(146) Nowak, H.,696 Nowoswiat, E.F.,657,888(69) Nozaki, Y.,726 Nussbaum, A. I,., 266,287(62) Nussbaum, J. L.,365 Nutt, R.F.,196,694,698 Nyc, J. I?., 420 Nystrom, L.,126

0 Obrenovitch, A., 283 Ockerman, P.A., 603 O’Connell. W.,54(81), 55, 56(81) O’Connor, M.L.,30 Ochoa, S.,640(97), 641 Oda, K..240 Oda, T.,544, 548 Odin, E.,58 O’Donovan, C. I.,49,57(42) Ofengand, E.J.. 332, 501(10), 502 Ogata, K., 94, 246(155, 156, 157), 249 Oginsky, E. L..541 Ogorodnikova, L. G.,600(207, 22l), 601 Ogston. A. G.,31,32(56) O’Hara, K., 29(44), 30, 31(44), 34(44). 38(44). 39(44), 40(44), 41(44), 42 (44). 43(44) Ohmura. E.,246(159), 249 Ohnuma, T..103. 107(18), 117(18) Ohsaka, A,, 177(10), 178, 185(10), 1% (10) Ohtaka. Y., 246(158), 249 Oi, T.,670, 673, 674(77), 727(77)

836 Oishi, M., 259 Ojamiie, M., 423 Okada, N., 394 Okada, S., 281 Okamoto, K., 246(162), 250 Okasaki, R., 252 Okuaki, T., 252, 253 O’Keefe, G., 111, 366 O’Konski, C. T., 708 Okubo, M., 269, 270(75) Okuyama, T., 515 Olafson, R. W., 364, 365(29) Old, L. J., 103, 110, 117, 118, 120(87), 489 Oleson, A. E., 254,255(19) Olislaegers, P., 55 Olitsky, P. K., 697,698(217) Oliveira, R. J., 740 Oliver, I. T., 600(197), 601 Olkowski, Z., 600(20l), 601 Olmstead, P. S., 72 Olsen, R., 376, 378 Olason, B., 605(297), 607 Omenn, G. S., 186,201 O’Neal, C., 230 Ono, K., 548, 609(61) Ontjes, D. A., 186, 200, 201(87), 202(87) Ooi, T., 213, 219(19), 221(19) Oosterbaan, R. A., 471 Opie, L. H., 633,635 O’Reilly, K. T., 14 Orjamiie, M., 396 Orkand, P. M.,600(215), 601 Orlowski, M., 94, 96(32), 125, 127, 137, 141(13), 142, 143(13, 120), 144(13), 145, 150, 151(154) Orr, G., 118 Ortanderl, F., 314 Ortiz, J., 421 Orton, W. L., 497 Orulresu, M., 600(203), 601 Oshinsky, C. K., 287 Ostern, P., 545 Ostrovskii, Y. M., 548, 605(54, 56) Ostrowski, W., 245(151, 1521, 249, 466. 467, 469(51), 470(60), 476 O’Sullivan, M., 55, 58 Otani, T. T., 146 Oth, A., 279, 280(21) Ottesen, M., 94, 130, 140(75). 141(116), 670

AUTHOR INDEX

Ouellet, L., 425, 428, 435(91), 436, 437, 438(162), 439(91), 442, 444(162), 445 (91, 162) Ove, P., 281 Overman, A. R., 18 Owens, H. S., 1% Oyaert, W., 605(29S), 607 Ozawa, H., 697, 720(213)

P Padykula, H. A., 606(321), 607 Paek, S., 605(308), 6%’ Page, J., 654 Paik, W. K., 692 Pajetta, P., 680 Pakdaman, P., 602 Palade, G. E., 553, 554(70), 556(70), 562 (701, 579(70), WWO), 651 Paleveda, W. J., 698 Pallansch, M. J., 126 Palm, L., 268 Palmer, R. A., 656 Pamiljans, V., 124, 126(6, 71, 137(6, 7) Pan, S. C., 126 Pancholy, M., 11 Pandhi, P. N., 598(176), 599 Panje, W. R., 606(325), 608 Pany, J., 451 Paoletti, C., 260, 282 Parikh, I., 200, 203(90) Park, C. R., 644 Park, D. C., 50 Parker, S., 651 Parks, J. M., 683, 699(139), 703(139) Parks, R. E., Jr., 645 Parr, C. W., 120 Parsons, M. A., 747 Parvin, R., 547, 567(44), 568(44), 570(44), 573(44), 574(44), 586, 587(44), 590 (44) Pascher, G., 144 Paasonneau, J. V., 432 Paterson, A. R. P., 349 Patrick, C., 641, 642(100) Patterson, L. M. B., 420 Patterson, M. K., 118 Paul, A. V., 261 Paul, R., 339 Pauling, L., 448 Paw, R. J., 553, 556(80), 557(80), 558

837

AUTHOR INDEX

(go), 560(SO), 574(80), 579(142), 581, 598(80), 599(80) Pausescu, E., 602 Pavlic, M.,377, 400(31), 405(31) Pavlov, R. A.,605(275), 606 Pedersen, K.O.,504,509(18) Pedersen, S., 149 Pedersen, T.A.,643 Pederson, C. S.,128 Peer, P., 13, 14(69), 17(69) Peeters, G.,772 Pekar, A. H.,113, 114(78) Peller, L.,88 Pel’ttser, A. S., 14 Penalver, M. D.,436, 437(163), 441(163) Pennington, R.J., 50,71 Penswick, J. H.,48 Penswick, J. R.,215, 222(29) Perez del Cerro, M. I., 605(287), 806 Perham, R. N., 128(70), 129(70), 130 Perlmann, G. E.,451, 476 Perrin, D.,387,389(77) Perrot-Yee, S., 368, 367(37), 368(37) Ferry, J. D.,434 Perry, S. V., 72 Pesce, A.,501, 502 Peters, E. L., 103, 106(11). lOir(l1) Peters, P. C., 118 Peterson, M.J., 596,597(169) Peterson, R. E.,104. 105, 109(42), 117 (371, 119(42) Petitclerc, C.. 379(52), 380, 401(52), 403 (521, 405, 406(138), 413(138) Petkov, P., 604 Fetreseu, A., 6Q5(286), 806 Petrie, S., 38(70), 39, 40(70). 41(70). 42 (701, W 7 0 ) Petronio, L., 806.(274), 606 Peyrot, A.,60!?, Pfeiffer, U., 600(193), 601 Pfleiderer, G.,314 Pflumm, M. N.,694, 721, 722(313) Pfrogner, N..55, 56(85), 57(85), 358 Phelan, J., 55 Philips, F. S., 49 Phillips, B. M.,347 Phillips. G.E.,433,456(35), 457 Philpott, D.E.,606(331), 608 Philpot, J. St. L., 772 Pianotti, R. S.,14

Pichierri, U., 548, 605(50) Pierce, J. E.,419(32), 420 Pier, K. A., 128(72), 129(72), 130 Piggot, P. J., 128(55), 129, 130(55) Pigretti, M. M.,397, 398(120), 405(120) Pihl, A., 623 Pilcher, C. W.,343 Pillet, J., 177(6), 178, 185(6), 186(6) Pincus, J. H.,80 Pinkham, C.,18 Piperno, G.,284 Pitot, H. C., 667 Plapp, B. V., 691 Platt, J. R.,127, 149(40) Plimmer, R.H.A., 418 Plisetskaya, E. M.,600(207, 221), 601 Plocke, D.J., 378, 379(36), 401, 4O2(50), 406(50) Plotch, S.,391 Plummer, T.H.,Jr., 651,669(34),670(34) Pochon, F., 177(6), 178, 185(6), 186(6), 755 Podder, S. K.,218 Popell. B. M., 48, 488, 612, 613, 614, 616 (41, 617(26), 618, 619(14), 622(14), 628,631 Poirier, M., 567 Pol, E.H., 316,321(41) Polatnick. J., 292, 750, 751(413) Polgar, P., 128 Pollak, J., 375, 550, 556, soO(zo8), 601 Pollock, M. R.,24, 25(2), 26(2, 3, 111, 28(36, 39, 40), 29(40), 30(2, 231, 31 (2, 36, 39. 40), 32(2, 36, 39, 401, 33 (2, 401, 34(23, 40), 35(2, 3, 231, 38 (20,40, 46, 70), 39(2, 20, 23, 40, 461, 40(2. 40. 70). 41(2, 701, 42(2, 23, 40, 70), 43(2. 201,44(98), 45(2), 46(70) Polyakova, N. M., 50, 66(54), 67(54), 68 (54)

Pongs, O., 220,221(55) Pontremoli. S., 600(203), 601, 614, 615, 616(22), 617,618(15, 21,27), 619,620, 621(15, 16), 622(15, 44), 623, 624(15, 30. 45), 625, 628(51), 627(22), 628 (35),629(40),630,631(30,38,39),632 (64). 633(64), 634, 638, 639, 645(74) Porath, J., 476, 493 Porteous, J. W.,421

838

AUTHOR INDEX

Porter, R. R., 128(55, 67), 129(67), 130 (551, 650 Portmann, P., 423, 424(69a), 425(69a), 429(69), 440(69)

Posen, S., 420, 426, 443 Posternak, Th., 367 Posthumus, C., 380 Potter, C. S., 71 Potter, J. L., 281, 291, 292, 299, 309(42), 318

Potts, J. T., 654, 671, 699(89), 703(89), 710, 717, 738

Pover, W. F. R., 603 Powell, C. A., 367 Powell, J. F., 55 Prager, M. D., 118 Prasannan, K. G., 605(284), 606 Pratt, E. A., 259, 282 Preiss, J., 332, 642(103), 643, 747 Presa, E. M., 128(55, 67), 129(67), 130

Rabinovitch-Mahler, N., 314 Rabinowitz, J. C., 332 Rabinowitz, J. L., 696 Rabinowitz, K. W., 48 Racker, E., 638, 641(85), 642, 643 Radding, C. M., 254 Radloff, R., 260 Raffan, I. M., 125 Rafter, G. W., 546, 568(32), 580(32) Raftery, M. A., 691 Ragade, I. S., 51, 52(69), 62(69) Raggi, A., 48(20), 49, 50(20), 66(147), 67, 68(147), 69(147), 70(147), 71W a ) , 73( 147, 153a)

Raggi, F., 600(196), 601 Raine, L., 419 Rajagopal, D. R., 150 Rajagopolan, K. V., 58 RajBhandary, U. L., 215(33), 216, 227, 228

Rakosky, J., Jr., 50, 76 Price, P. A., 292,293,295, 296, 297(35, 37) Ralph, R. K., 308,310 Rall, T. W., 366, 367(32), 371(33) Price, V. E., 132 Ramachandran, G . N., 669 Primosigh, J., 374 Ramadan, M. E. A., 94, 105 Printz, M. P., 714 Privat de Garilhe, M., 154, 177(6, 71, Ramakrishna, M., 126, 150 178, 185, 186(1, 6), 207, 243, 252, 256 Ramakrishnan, T., 104, 117(30) (6), 272, 285(2), 290, 308, 314(25), Ramiah, A. J., 613 315, 317(22, 23), 320(22, 23), 322 Ramponi, G., 422 Ramsden, E. N., 773 (25), 328(22, 231, 476 Randall, S. S., 133 Proctor, C. M., 697,648(215) Randle, P. J., 604 Prohaska, E., 44 Rands, D. G., 771, 715(274), 716(274) Prokof’eva, E. G., 436 Ranieri, M., 70, 71(154a), 348 Pucher, G . W., 125, 132 Raniero, R., 671 Puig-Muset, P., 750, 751(411) Ransome, 0. J., 12 Purzycka, J., 49, 50(40) Purzycka-Preis, J., 64, 65(125), 66(125), Rao, J. G. S., 629 Rao, S. N., 66, 67(142), 68(142), 69(142) 67(125), 68(125), 70(125) Pustoshilova, N. M., 333 Rao, V. N. V., 14 Putnam, F. W., 128(68), 129(68), 130, Rapoport, S., 531, 534(12) 266 Rashba, 0. Y.,609 Putter, I., 154 Raskova, N. V., 394 Pynes, G . D., 540 Ratner, S., 125, 133 Ratych, 0. T., 113, 114(77) Q Rauenbusch, E., 24, 26(9), 43(9), 44, 108, Quigley, J. P., 420 (55)

R Raacke, I. D., 711 Rabin, B. R., 71, 749, 763, 771, 773, 779, 780

109(65), 111(65), 112(65), 113(65), 114(65), 115(65), 116(65), 118(65), 281 Ray, T. K., 345 Razin, A., 66(143), 67, 68(143), 69(143)

AUTHOR INDEX

Raaaell, W. E., 272, 285(4), 314(27, 281, 315, 317(18), 318, 319, 320(18), 322, 328(18), 329, 330(3), 331(3, 151, 332 333, 336(3) Read, P. A., 456(35), 457 Reading, E. H., 750, 751(397a) Reddi, K . K., 177(8, 9), 185(4, 5, 8, 9), 186(4), 201(9) Redfield, R. R., 654 Reed, S., 800 Reese, C. B., 215(34). 216 Reeves, J. Y., 109 Reich, E., 256, 757 Reid, A. F., 432 Reid, T. W., 377, 392(30), 393(30), 394 (30), 398(30). 400, 405(31), 408(30). 410(30), 413 Reindel, F., 126 Reiner, E. J., 366,367(44) Reiner, J. M., 459,460(40) Reinhart, F. E., 124, 142 Reis. J. L., 338 Reim, M. M., 770 Reith, A., 491 Reithel, F. J., 2, 3, 5(6), 7(27), 8(6, 27, 43), 9(6, 14), lO(6, 14, 431, 11(6, 43), 12(43), 18(27), 498 Remington, L., 755 Renold, A. E., 544, 596(6) Repaske, R., 338, 361 Resnick, H., 691, 732 Ressler, N., 55 Revel, J. P., 96 Rey, J., 603 Reynolds, J., 378, 379(47, 531, 380, 382, 383(49), 384(67), 387(47, 53), 389 (671, 403(49), 405(53), 406(49) Reynolds, J. H., 677, 678(104) Reaaonico, A., 548, 605(49) Rhodes, H. K., 46 Ricciardi, I., 605(318, 319), 606(322), 607, 608 Rice, A. C., 128 Rich, L. D., 114 Richards, E. G., 279, 301, 503, 504(16), 509( 16) Richards, F. M., 157, 163, 196, 656, 657 (62), 658(62), 666(62), 667(62), 669, 670(73), 671(73), 672, 673, 674(94), 675, 676(97), 677, 678(103), 683(61),

839 686, 688, 699(94), 703(94, 2341, 704 (97), 724 (62), 738, 750, 751 (4161, 763, 785 Richards, G. M., 313, 314(8, 29), 315, 316 ( 8 ) , 318(35), 319(35), 320, 323(29, 50), 324, 325, 326(29), 342 Richardson, C. C., 253, 254(8), 255, 256 (25), 265, 266(59), 304, 305(56), 306 (56a), 315, 330 Richardson, D., 606(321), 607 Richardson, D. C., 156, 156(31), 157(42), 160, 164(31), 173(31), 180, 182(28), 183, 187(49), 195(49) Richardson, J. S.,155, 156(31), 157(42), 160, 164(31), 173(31), 183, 187(49), 195(49) Richardson, K. E., 481,482(81) Richetta, G., 548, 605(50) Richman, P. G., 94, 125, 127(13), 141(13), 143(13), 144(13), 145(13) Richmond, M. H., 24, 26(18), 29(25, 41, 45), 30(25), 31(25, 41, 45), 32(25, 41, 581, 33(41, 45, 58, 62a), 34(25, 41, 45, 58), 38(25, 41, 58, 81), 39(25, 41, 45, 58), 40(25, 45, 811, 41(25, 81), 42(25), 44, 46(41), 383, 390(68) Richter, L., 553, 554(77), 600(77) Rick, W., 436 Ricketts, T. R., 600(199), 601, 581(154), 582 Ridlington, J., 530, 531(8), 532(8), 533 (13), 534(8), 535(13) Riedel, E., 418(17), 419 Riehm, J. P., 675(98, 99, 100, 101), 676, 677, 678(106), 716 Riesel, E., 12 Rigillo, N., 600(187), 601 Riley, V., 120 Rimai, L., 708, 709, 727 Rinaudo, M . T., 600(209, 213), 601, 602 Ring, B., 778 Riordan, J. F., 173, 619 Rippa, M., 621, 622(44) Ritchie, S., 602 Robbins, E. A., 525, 526(61), 540 Robbins J. E., 2, 3, 5(6), 8(6), 9(6, 14), lO(6, 14), 11(6) Robbins, P. W., 529 Roberts, E., 80, 87(1) Roberts, J., 105, 107, 108(43), 109, 114

AUTHOB INDEX

Roberts, G. C. K., 712, 724, 725, 732, 763,784 Roberts, W.K.,177(12, 14, 15), 178, 185 (12, 14, 15), 186(12), 252 Roberts, W. M.,418 Robertson, H. D., 241, 2420291, 245(128, 129) Robertson, J. M.,446 Robins, M.J., 59 Robins, R. K.,51, 52(65), 53(65), 59(65), 80(65),61(651,63(65) Robinson, J. C., 419(32), 420 Robinson, N.,347 Robinson, R., 27 Robinson, R., 450, 477 Robison, G.A., 366,371(34,36) Robison, R., 418(15), 419, 420, 421(33) Roblin, R., 326 Robmn, E. B.,422,423,424 Rocchi, R., 671, 695, 699(242, 243, 2441, 701(235, 236,237, 238, 239, 240, 241, 242, 243, 2441, 702 Roche, J., 420,423,429,612 Roche, M.J., 450 Rockwell, M.,55, 56(86), 57(86), 63(86), 64(86) Rogers, D., 498 Rolinson, G. N.,38(79), 39, 40(79), 41 (79), 42(79) Rolon, C.I.,484 Rome, N.M.,545,580(16), SOO(l6) Ronca, G., 48(20, 49), 60(20), 58, 59, 66(147), 67,68(147), 69(147),70(147), 71(154a), 73(147, 153a) Ronca-Testoni, S., 48(20), 49, 50(20), 58, 59(106), 66(147), 67, 68(147), 69 (147), 70(147), 71(154a), 73(147, 153a) Ron-Zenziper, E., 30, 31(54), 33(54), 42(54) Ronzio, R. A., 150 Roon, R. J., 14 Rooney, S., 55 Rosemeyer, M.A., 711 Rosen, 0.M.,619, 629, 635,636(80), 637 (SO), 638, 640(92, 941, 641, 645(94) Rosen, S. M.,619, 629,635,636(80), 637 (so),638, 640(94),641, 645(94) Rosenbaum, R. M.,484

Roeenberg, E., 757 Rosenbluth, R., 280 Rosenkrantr, H., 467 Rosenthal, R. L.,496 Rosevear, J. W.,71 Ross, C.A.,771, 780 Ross, M.H.,436 Rossi, C.A., 58,70,71(154a) Rostgaard, J., 347 Roth, J. S.,750, 751(40), 770,772 Rothe, W.,39 Rothen, A., 705 Rothman, F.,378, 379(46), 382(46), 384, 389(46), 424 Rothman, U.,604 Rothschild, J., 451,498(12b, c) Rotman, B.,454 Rottman, F.,230,747 Rounbehler, D.,560, 566(65), 568(65), 570(65), 573(65), 574(65) Rousch, A. H., 49, 50(37), 51(28), 53(62) Roussos, G.G.,259,282 Rovainen, C.M.,596 Rovery, M.,11 Rowe, H.J., 222(79), 223 Rowley, B.,104, 116(35), 119(35) Roy, J. E.,50, 76(68) Roy, K.L.,76 Royce, A.,43 Russell, F.E.,313 Russell, R. G. G., 421 Rust, J. H.,13 Rutenburg, A. M.,148 Rutishauser, U.,128(66), 129 Ryan, E. M.,516 Ryan, R. M.,103 Ryan, W.L.,119 Rybamka, J., 467,469(51), 470(60) Rydon, H.N.,126, 127(30) Ryle, A. P.,654 Rzhekhina, N.I., 435,436,443(154) Riiterjans, H., 220,221(55), 712, 724(281) Ruffier, N. K.,282 Ruiz-Herrera, J., 14 Rumbaugh, H.L.,541 Rumke, P. H.,344 Rumley, M.K.,179,180(20), 185(20) Rupley, J. A., 157, 648,657, 670, 673(76), 674

841

AUTHOR INDEX

Rusch, H.P., 750,751(397) Rush, E. A., 150 Rushizky, G. W.,177(12, 14, 151, 178, 185(12, 14, 151, 186(12), 212, 223, 229, 239, 244(114, 140), 247(140), 249, 755

S Sabath, L. D., 27, 30,31, 38(31), 39(31), 40(31), 42(31), 43(31, 66),44(47) Sachs, H.,133 Sadowski, P.D., 266,267(66), 268(66) Sadron, C., 279, 281(24) Saeed, M.A., 49 Saenr, A. C., 697,698(217) Sage, H.J., 715 Saidel, L.J., 127 Saintot, M.,423 Saito, M.,240, 244(121) Sakabe, K.,252 Sakai, T., 246(158), 249 Sakaki, T.,216 Sakaquchi, K.,758 Saksena, T.K.,11 Salanito, J., 386 Salas, J., 593(165), 594, 632, 633(71), 634

(71) Salas, M. C., 593(164, 165), 594, 596,614, 619(13), 632, 633(71), 634, 640(13), 645(13) Salganicoff, L., 367,368(50) Salkowski, E.,649 Sallusto, A.,605 (318,319), 606(322),607, 6M

Salnikow, J., 292, 293(39, 39a), 295(39) Salomon, L.L.,603 Sammons, D.W., 69,73(153b) San Clemente, C. L.,451,498(12a) Sandeen, G.,292,750,751(421) Sandoval, A,, 671, 681 Saneyoshi, M.,226, 227(86), 228(86), 230(86, 89, 94) Sanger, F.,129, 234, 237(107), 238(107), 240, 380 Sanger, K.C. S., 51,76(60), 77(60) Sanner, T.,623 Sanno, Y.,320 Sano, I., 144 Sansom, B. F.,107

Santacroce, G., 773 Santo, R.E., 50 Sapico, V., 639 Sarfare, P. S., 722,723(320), 736 Sargeant, K.,104, ll6(38), 119(38) Sarkar, N.R.,281 Sarkar, P.H., 606 Sarngadharan, M. G.,617,628 Saroff, H.A., 769 Sarrosin, G.,389 Sartori, S.,805(274), 606 Sasisekharan, V.,889 Satake, K.,515 Satirana, M.L.,643 Sato, K.,212,213(12),215,223(12) Sato, S.,218,220,224,228(84) Sato-Asano, K.,215, 222(73), 223, 233(73) Satta, M. A,, 609 Saunders, H. G.,27 Saunders, M.,723 Saundry, R. H.,771 Sliwada, F.,761 Sawai, T.,29(44), 30, 31(44), 34(44), 38(44), 39(44), 40(44), 41(44),42(44), 43(44) Saxena, R. P., 55 Saz, A. K.,26,43,44(102) Scala, J., 641,642 Scalturin, A., 721 Scatturin, A,, 629, 671, 695, 699(243), 701(236, 2431, 702 Schachman, H. K., 279, 304,307(53),503, 504(16), 505(20), 509(16), 530, 531, 673 Schaefer, E.-A,, 755 Schaeffner, A., 451,501 Schaeffer, H.J., 58,59 Schaller, H., 268 Schally, A. V., 128(59), 129 Schechter, A. N.,155,184 Schechter, E.,505 Scheele, C., 120 Scheetz, R. W.,104, 117(40),119(40) Scheffler, I. E., 3 B ,310(65) Schellman, J. A., 705, 710, 722, 733, 769 Scheraga, H. A,, 505, 648, 670, 673(76), 674(77), 675(98, 99, 100, 101), 676, 677, 678(106), 684, 690(140, 143, 144, 147), 699(153),703(153), 711,712,713,

842 714, 716, 724(281), 727(77), 729(277, 337), 730(277,337),732,734,741 Scherbaum, 0. H.,248( 1761,250 Schick, L.,540 Schildkraut, C. L., 713 Schito, G.C.,501, 502 Schlamowitz, M.,419 Schlegel, H.G.,13 Schlesinger, M.J., 375, 376, 378, 379(47, 53), 380, 382(65), 383(49), 384(44, 67), 385(44), 386(44), 387(47, 53), 389(66, 67), 390, 393(44), 403(49), 405(53), 406(49), 534 Schlesinger, S., 376, 390 Schlessinger, D., 243, 245(132), 375, 379 (211, 382(21), 385(20), 389(21), 394 (21), 395(21) Schlutz, G. A., 127 Schmalfuas, K.,103 Schmetz, F.J.,623 Schmid, K.,128(71), 129(71), 130 Schmidt, C.L.A.,149 Schmidt, G.,48(19), 49, 51(13, 14), 329, 443,459, 545 Schneider, E.G.,739 Schneiderman, H.,420 Schoenewaldt, E.F.,698 Scholtan, W.,108 Schonheyder, F.,454 Schreiber, W., 553, 554(77), 600(77) Schreier, K.,281 Schrier, E.E.,732,734 Schroeder, E. A. R., 638, 641(85), 642, 643 Schull, G. M., 40 Schulman, M. P.,48(18), 49 Schultz, G.,369 Schulz, A. S.,49 Schulz, D.W., 432 Schulz, R.,644 Schumaker, V. N.,279,304 Schutte, E.,553,554(77), 600(77) Schwartz, B. S.,14 Schwartz, D.P.,126 Schwartz, J., 377, 378(35), 380, 396(35), 397, 398(35, 116), 405(35, ll6), 425 Schwartz, J. H., 109,118,120(67) Schwartz, M.A.,58 Schwartz, M.K.,338,343,496 Schwarze, P.,393, 446

AUTHOR INDEX

Schwender, C. F., 59 Scoffone, E., 671, 680, 682, 683(131), 691, 695, 699(242, 243, 244), 701(235, 236, 237, 238, 239, 240, 241,242, 243, 2441, 702 Scott, D. B. M., 547,597,604(48) Scott, E.,14,480 Scott, R. A.,727 Scott, T.G.,343, 347 Scrutton, M.C.,72,516, 594 Scurzi, W.,7, ll(39) Scutt, P.B.,424,425 Seal, U.S.,540 Sealock, R. W.,301 Secchi, G.C.,548,605(49) Segal, H.L., 546, 553, 557, 560(72), 563, 567(31), 574, 575(72), 578(71), 582 (31), 583, 585(31), 586, 587(31), 590 (72), 594(31), 596(31), 598(72), 599 (71) Segal, L., 15 Sehgal, P.P.,2,3,5(15), 6,9(34) Seibles, T.S.,685 Seidel, M.K.,291 Sekiguchi, M.,269,270(75) Sekine, H.,758 Sekita, K.,13 Sekiya, T.,218, 222(76), 223 Sela, M.,31, 32(57), 42(57), 45(57), 675 (102), 677, 678(111, 112, 113, 1141, 681(111, 112, 114), 691, 693(169a, 182),694,695,710,715,716,758 Sellin, H.,117 Seneca, H., 13, 14(69), 17(69) Senft, G.,369 Senji, U.,148 Seno, T.,227 Setlow, B.,65, 66(129, 131), 68(129, 131, 138, 141), 69(131, 141), 70(129), 72 (1411,73(131,138) Setlow, P.,256,258(29), 269 Setlow, R.B.,321 Seto, T.A., 40 Sett, R.,369 Sevag, M.G.,149 Shack, J., 281 Shada, J. D.,48 Shadaksharaswamy, M.,6 Shah, N.S.,602 Shalina, N. M., 316, 318,321

843

AUTHOR INDEX

Shall, S., 656, 681 Shannon, C., 567 Shanta, T. R., 368 Shanygina, K. I., 600(206), 601 Shao-Khua, V., 106 Shapira, R., 650, 651, 750, 751(407) Shapiro, S.,622, 623(45), 624(45), 630,

Siegel, L. M., 6 Siekevitr, P., 553, 554(70), 556(70), 562 (70), 579(70), 600(70)

Sierakowska, H., 327, 328, 336 Signer, E., 254, 378, 380(38), 384(38), 385(38, 69), 386(71)

Siler, W. M., 120 Silman, G., soO(199), 601 633(63), 645(63) Shapot, V. S., 314, 317(24), 320(24), 328 Silman, H. I., 677, 678(112), 681(112) Sim, G. A,, 446 (24) Simbonis, S.S.,600(210), 601 Sharma, C., 593(163), 594, 596 Sharma, N. N., 605(285), 608(334), 608, Simmons, N. S., 505, 540, 721, 723(312) Simon, J., 318 809 Sharpless, T. K., 51, 52(69), 62(69), 74 Simon, L. N., 51, 52(65), 53(65), 59(65), 60(65), 61(65), 63 Shaw, D. C., 380 Simonart, P., 149 Shaw, J. G., 497 Simons, E. R., 739 Shenr, M. J., 758 Simpson, R. T., 378, 379(51), 380(41), 384 Sheinin, R., 30, 45(48) (41), 387(41), 389(41), 392, 393(96), Sheit, K. H., 222(75), 223 394(96), 395(96), 401, 402(51), 403 Sheppard, R. C., 128(64), 129 (51), 404(96), 405(134), 406, 408(96), Sherwood, L. M., 717, 738 409(96), 410(96), 411(96), 723 Shibata, K., 517, 685, 689 Sims, E. S.,316 Shiio, I., 240, 244(122) Singer, B., 327, 476 Shima, T., 317 Singer, M. F., 241, 244(126, 1271, 245 Shimada, K., 269, 270(75) (147), 249, 314(26), 315, 332 Shimada, K., 356, 357(8), 359(8), 360(8) Singer, S.J., 715 Shimizu, B., 51, 53(75), 74(75) Singhal, R. L., 645 Shimizu. S., 240, 244(122) Sinsheimer, R. L., 266, 279, 291, 308, 317, Shindo. Y., 536, 540(24) 321 Shinoda, T., 128(68), 129(68), 130 Sisini, A., 609 Shiobara, Y., 216, 221 Sjoquist, J., 181 Shiozawa, C., 222 Shirafuji, H., 51, 52(63), 53(63), 77(63) Skoczylas, B., 248(175), 250 Skoog, L., 299 Shirley, B. C., 71(167, 168), 72 Skujins, J. J., 15, 19 Shooter, E. M., 711 Slater, J. S.,128 Shorey, C., 375,600(208), 601 Nor, H., 282, 283 Short, E. C., 255 Small, C. W., 426 Shortman, K., 213, 772 Small, P. A., 419, 423(29) Shrago, E., 645 Shugar, D., 327, 328, 336, 714, 750, 751 Smiley, K. L., Jr., 51, 52(68), 64(68), 65(68), 6668, 145, 145a), 67(68), (408) 68(68, 145. 145a), 69(145), 70(145. Shull, K. H., 600 145a) Shumaker, V. N., 709 Smillie, R. M., 642 Shuster, L., 74, 353, 364 Smirnovs, T., 375 Sia, C. L., 627 Smith, A., 114 Sicard, P. J., 283 Smith, D. L., 609 Siddons, R. C., 617 Smith, E. L., 90, 91(20) Smith, H. O., 263 Siebert. G., 451 Smith, I., 434, 725 Siedler, A . J., 14

844 Smith, J. D., 476 Smith, J. K.,419(32), 420, 429, 437(116) Smith, J. T.,24, 31, 38(82, 83), 39, 42 (651,43(94), 44(65) Smith, L. D., 65, 66(128), 68(128), 69 (128) Smith, M., 328, 366, 367(44) Smith, M. A.,222(79), 223 Smith, P.W.G.,126, 127(30) Smith, R. A., 308, 310, 547, 567(44), 568 (441, 570(44), 573(44, 130, 574(44), 586, 587(44), 590(44) Smith, R.H., 274(14), 275 Smith, R.L.,605(296), 607 Smithers, M.J.,698,700(227) Smithies, O.,503 Smyth, D.G.,129,512,654,655 Snoke, R. E.,555, 556(82, 831, 557, 560 (951, 568(95), 579(82), 580(95, 971, 598(95), 599(95) Snustad, D. P., 268 Soames, K. M., 418 Soave, C.,277,284,332,333 Sober, H.A., 101, 120(1), 212, 223, 229, 239, 244(114, 1401, 247(140), 249, 755 Sober, H.H., 589,590(157) Sobin, L. H.,119 Sobotka, H., 697,698(219) Soda, K.,94 Sodja, A., 154(21), 155, 163(36), 180, 181 (26), 182(26), 183, 184(47), 196(47). 197(47), 198(47) Sokawa, J., 215(32), 216,227(32) Sokolovsky, M.,173,517 Sols, A.,593(164, 165), 594, 596, 614, 619 (13), 632, 633(71), 634(71), 640(13), 645(13) Somers, G.F.,2 Sommer, A.,466 Sondey, J., 698 Sonenberg, M., 735 Song, C.S.,344,345 Sonnenschein, N.,600(217),601 Soodsma, J. F., 547, 549(45), 556, 557 (89), 558(45, 89), 561, 563(89), 567 (45), 568(45, 1011, 5730391, 574(45), 576(45), 577(45), 580(89, 1011, 585 (45), 587(45), 590(45), 602(45, 101) Sorm, F., 218, 222(77), 223, 228, 284, 754 Sornson, H.L, 119

AUTHOR INDEX

Spackman, D. H., 181, 512, 513(36), 654 Spahr, P. F.,241, 243, 244(125, 143), 245 (1311,249,333,334(36), 335 Spears, C. L., 7, ll(39) Spencer, B.,50 Spencer, J. H., 302 Spencer, N.,477, 480(76) Sperling, R.,692, 693(183, 1841, 750, 751 (418) Sperow, J., 535 Spicer, S.S.,362, 375 Spiro, D.,609(354), 610 Spiro, T.G.,404 Spitrer, R.,144 Springell, P.H., 142,713 Springer, K.,14 Springhorn, S.S.,746 Squires, R. W.,108 Sripathi, C.E.,540 Srivastava, S. K.,645 Stadtman, E.R.,147,544 Stadtman, T.C., 420,435(38) Staehelin, M.,215 Stahlmann, C.,553,554,600(77) Stamm, N. B.,645 Stanescu, V.,605(302,303), 607 Stanier, J. E.,772 Stark, G.R.,512, 516, 679, 882, 686, 690 (133, 158)

Stark, J. B., 128 Starr, J. L., 150 Start, C., 644 Stasiuk, L., 154, 174(12), 179, 186(22), 193(22), 291,328 Staub, F.B., 695 Staudinger, H.,581(149), 582, 600(193), 601 Steele, B. B., 29(43), 30, 31(43), 32(43), 33(43), 34(43), 39(43), 40(43), 41 (431,42(43), 43(43), 46(43) Steenback, H.,149 Steensholt, G.,103 Steens-Lievens, A., 466 Steers, E.,Jr., 128(73), 129(73), 130 Stein, A. A., 602 Stein, S. S., 430,454 Stein, W.D.,886, 771 Stein, W.H.,129, 148, 181, 220, 292, 293 35, 38, 39, 39a), 295(35, 39), 296(35). 297(35, 371, 512, 513(35, 38), 648,

845

AUTHOR INDEX

650, 651, 653(22, 25), 654, 655(55), 669(25), 670(25), 679, 682, 683(130), 686, 688(160), 690(158, 159, 161, 1631, 691, 744 Steinberg, C. M., 501 Steinberg, I. Z., 691, 692, 693(182, 183, 184), 750, 751(418) Stellwagen, E. C., 697,698(216) Stepanova, N. G., 600(200), 601 Stephenson, M., 49 Sternback, H., 394, 395(108) Stetten, D., Jr., 501, 572 Stetten, M. R., 546, 547, 550, 557, 559, 580(92), 562, 566(65), 567(36, 381, 568(65, 90, 112, 113), 570, 571, 573 (65),574(38, 65, 112, 125), 577(112), 578(90), 579(125), 581(38), 587(38), 590(38), 598(90, 1131, 800(38) Stewart, C. P., 49 Stewart, D. J., 14 Stewart, K . K., 7, 9(35) Stockert, E., 118 Stockx, J., 55, 750, 751(405, 417) Stone, D., 128(56), 129 Stowell, R. E., 600(186), 601 Strachen, R. G., 698 Stracher, A., 713 Strachman, N., 442 Strassburger, M., 126, 134, 135(102), 136 (102) Straumann, F., 421 Strauss, B. S., 261, 269 Strecker, H. J., 125, 138(14, 15, 16), 139 (14) Strejan, G., 46

Stripati, C. E., 64 Strittmatter, P., 420 Stryker, U. H., 697, 698(216) Stuart, A., 215(33), 216, 228 Stubbins, S. E., 19 Studier, F. W., 280, 265, 288(59), 279 Stulberg, M. P., 525, 526(61), 540 Stumpf, P. K., 80 Sturtevant, J. M., 740 SU,C.-C., 313 Su, J. C., 75 Subbarow, Y.,432, 487,518 Subrahmanyam, K., 605(284), 608 Suelter, C. H., 51, 52(68), 64(68), 65 (68), W68, 133, 145, 1454, 67(68),

68(68, 145, 145~11, 69(133, 1451, 70 (133, 145, 145a), 71(133), 73(133)

Sugimura, T., 317, 326 Sugino, Y., 356, 357(8), 359(8), 360(8), 432

Suhadolnik, R. J., 51, 52(70, 72), 53(70), 55, 56(84), 57(84), 59(70), 60(70), 61(70) Suhara, I., 246(159), 249 Suld, H. M., 103, 106(17) Sulkowski, E., 154, 155(9), 156(8), 174 (12), 177(13), 178, 179, 185(13), 186 13, 22, 59), 193(22), 290, 291, 310(7), 314, 318, 326(13), 342, 350(23), 352 (23) Sullivan, D., 377, 400(31), 405(31) Sulston, J. E., 215(34), 216 Summer, G. K., 422,433 Summ, H. D., 605(307), 607 Sumner, J. B., 2, 4(2), 8, 10(40), 12(40), 13(2) Sun, Y. K., 697, 698(218) Sung, S.-C., 185, 280,307,316,319(40) Sur, B. K., 469 Surand, G., 12 Suriano, J. R., 182, 183(40), 187(40) Suskind, S. R., 230, 247(100) Susman, M., 501 Sussman, H. H., 419, 423(29), 424, 425 (86), 426(86), 437(85) Sutherland, E. W., 366, 367(32, 36), 368 (36, 49), 369(36), 370, 371(33, 34) Suzuki, I., 75 Suzuki, T., 128(65), 129 Svedberg, T., 504, 509(18) Svensson, H., 144 Swanson, M. A., 544, 545, 546(19), 566, 580(19), 581(18, 191, 800(19) Swenson, C. A., 733 Swenson, M. R., 283 Sykes, G., 43 Synge, R. L. M., 125, 126(17), 128(17) Szekely, M., 670 Szemplinska, H., 327 Szent-Gyorgyi, A. G., 505 Szentkiralyi, E. M., 72 Szewczuk, A.. 96, 128, 148, 149(135) Szmigielski, S.,548, 605(51, 272, 300).

606,607 Szpirer, J., 254

846

AUTHOR INDEX

f Taborsky, G., 650,653,697,698(220) Tada, M.,228 T d j e d d , I. B.,604 Taft, H.L., 567, 568(112), 574(112), 577 (112) Tagawa, T., 51,53(75), 74(75), 213 Tagnon, H.J., 466 Tait, G., 378, 379(41), 380(41), 384(41), 387(41), 389(41), 391, 408(91), 409 (91) Takagi, Y., 261,269, 270(75) Takahashi, F.,356, 357(7), 359(7), 361(7) Takahashi, K.,208(5), 209, 210(5), 211 (51,212(5), 213(5), 216(11), 218, 219, 220(11), 221(11), 222,223(5), 689 Takahashi, S., 127 Takahashi, T., 672,770 Takai, N.,2OS(6), 209, 210, 211(8), 230 (6), 231(6, 81, 232(6, 81, 247(6, 8, 102) Takakura, K.,498 Takayi, M.,757 Takazi, M.,756 Takei, S.,341,350(17, 18, 19,20,21) Takeishi, K.,240,244(121) Takemura, S.,226, 227, 230(90),232(90), 236(90), 237(90), 756,757 Takenaka, O.,685,689 Taketa, K.,48, 488, 614, 618, 619(14), 622(14) Talsky, G.,18, 19(104) Talwar, G.P.,76 Tamburro, A. M.,629,671, 695,699(743), 701(236,241,743),702,721 Tamura, G., 213 Tanaka, A.,617 Tanaka, H.,602 Tanaka, J., 219 Tanaka, K.,124,138(9), 245(136), 249 Tanaka, S.,772 Tandler, B.,345 Tanford, C.,504, 505(22), 509, 651, 710, 711, 715(274), 716, 725, 726, 733, 734 Tanis, R.J., 6,9(34), 14(47), 30, 31(55), 33(55) Taniuchi, H., 154(21, 22, 23), 155(5), 156(22), 160(22), 163(36, 37, 381, 175(25), 179, 180(20), 181(25, 26, 271,

182(23, 26, 271, 183(40). 184(47), 185(20, 25), 187(40), 195(48), 196 (47, 48, 531, 197(47, 48, 53, SO), 198 (47, 80, 81), 199(80, Sl), 200(48, 80, 811,203(81, 86) Taponeco, G., 70,71(154a) Tappel, A. L.,484, 489,490, 540, 750, 751 (415) Tarentino, A., 651 Tarien, E.,303 Tarr, H.L.A.,348,351 (75) Tashima, K.,618 Tashima, Y.,624 Tata, J. R.,549 Tatarskaya, R. I., 50, 245(137, 138), 249, 321 Taylor, I. E. P.,11 Teller, D.C.,503,504(16), 509(16) Teller, J. D.,117 Tench, A. L.,651,669(34),670(34) Tener, G.M.,316, 321(41), 325 Teplova, N. M.,333 Terao, T., 221,333,334(37), 758 Terner, J. Y.,609(354), 610 Testa, E.,612 Tewari, C. P.,49 Tewari, H. B., 347, 605(281, 282, 2881, 606 Tewari, K.K., 51,76(60),77(60, 189) Thach, R.T., 222(78), 223,233(78) Thain, D.E.,8, lO(42) Tham, R.,126 Thannhauser, S.J., 443,459 Thiele, G.,536,540(23) Thierfelder, H.,132 Thiery, J. P.,277,284(19),332 Thomas, C.A., Jr., 279,304 Thomas, M.,605(273), 606 Thomas, R.,254 Thompson, E.0.P., 129 Thompson, J. E.,548, 609(60) Thorbecke, G.J.,489 Thorne, C.B.,94 Thorp, R. H., 63 Thorpe, W.V., 125 Thurman, R.G.,644 Tice, S.V.,101, 120(1), 146, 147(127) Tikhonenko, T. I., 316,321 Timasheff, S. N.,275 Ting, C.C.,75

847

AUTHOR INDEX

Ting, S.-M., 600(212),601 Tinoco, I., 218 Titani, K.,128(68), 129(68), 130 Todd, A. R.,751 Toh-e, A.,230,247(101) Tolbert, G.,241, 244(126) Toljedal, I. B.,348 Tolman, R. I,., 51, 52(65), 53(65). 59 (651,60(65),61(65),63(65) Tornita, K., 536,540(24) Tomlinson, N.,338 Tornlinson, R.V.,325 Tomoyeda, M.,244(142), 246(160, 161, 1621, 247(171), 249, 250 Tomozawa. Y.,55, 56(92). 65(92), 70 (92),73(92), 74(92) Tono, H., 539 Tonornura, Y.,72 Tonutti, E.,606(328), 608 Tooney, N.,505 Torella, M., 605(318), 607 Toribara, T. Y.,432 Torriani, A.. 374, 375, 377, 378(32, 33). 384,385(20, 691,392(3), 393(3) Torti, G., 284 Tosa, T., 149 Tota, G., 605(301), 607 Totsu, J., 341,350(21) Tournaire, D.,603 Touster, O.,343,351(31),352(31) Tower, D.B.,103, 106(11), 107 Townend, R.,273,276(13) Traniello, S., 614, 615, 616(22), 617, 618 (15), 621(15, 16), 622(15, 44), 623, 624(15, 30, 45), 627(22), 831(30), 638 Trebst, A. V.,643 Trentharn, D.,392, 393(98), 394, 395 (107), 404(98), 408(98), 410(98), 411 (107), 442, 445(177) Trautrnan, M.L.,291,299(20) Tridgell, E.J., 28(36), 30, 31(36). 32(36), 43, 44(98) Trilling, D.M.,258 Trim, A. R.,49 Tristrarn. G. R.,274(14), 275 Tristram, H.,390 Tritsch, G.L.,132 Tronche, P.,606(337), 608 Trubowitz, S.,423 Trump, B.F.,soO(lS6), 601

Trupin, J., 230 Trupin, K.,651, 750, 751(402) Truupyl’d, A. Y.,605(290), 607 Tsair, L.,147 Tsernoglou, D.,163, 656, 657(62), 658 (62), 666(62), 667(62), 683, 724(62), 785 Ts’o, P. 0. P.,732 Tsong, T. Y.,740 Tsuboi, K. K., 454, 459(23), 460(40), 469(23), 481 Tsuda, Y.,261 Tsugita, A,, 244(141), 245(148), 249, 466 Tsuji, Y.,104,107 Tsukada, Y.,364,365 Tsushima, K.,343, 344, 345(32, 331, 348 (33) Tuchrnan, L. R., 496 Tully, E.,54(82), 55,56(82), 57(82) Tunis, M.,473 Tunski, W., 248(175), 250 Turchini, J. P.,800,605(294), 607 Turner, C.W., 49 Turner, D.H., 50, 66(48), 68(48), 69(48), 70(48), 609 Turner, J. F.,50, 66(48), 68(48), 69(48), 70(48), 609 Tutas, D. J., 315, 318(35), 319(35), 320 (35) Tuttle, L. C.,125

U Uchida, K., 246(158), 249 Uchida, T., 208(5, 6, 71, 209, 210(5, 71, 211(5, 7, 8), 212(5), 213(5), 215(34, 35), 216(11, 27), 218, 220(11), 221 (ll), 223(5), 224(81, 831, 225(81). 226(27, 85), 227(30, 32, 35, 851, 228 (30, 8 4 ) . 230(6), 231(6, 81, 232(6, 8, 104), 233(103, 1041, 234(7, 301, 235 (7,30), 236(30, 35), 237(30, 106, 1 W , 238(30, 108), 239(30), 241, 247(6, 8, 102, 105, 123, 1241, 248(7, 30, 108, 173), 250 Uchiyama, K., 94 Ueda, S., 609 Ueda, T., 213, 216(18) Uehara, K.,16, 17(90) Uemura, T., 149 Ui, M., 432

848

AUTHOR INDEX

Ukita, C., 756, 758 Ukita, T., 218, 221, 222(76), 223, 229, 240, 244(121), 246(97), 333, 334(37), 672, 752, 757, 759, 770 Uliana, J. A., 148 Ullman, A., 387, 389(77) Ulrich, D. V., 705,706 Umiastowski, J., 50 Unemoto, T., 356, 357(6, 71, 359(6, 7), 360(6), 361 Unkeless, J. C., 345, 351(48) Uno, I., 230, 247(101) Uozumi, T., 213 Urnes, P., 722 Usher, D. A., 783 Utter, M. F., 48, 72,516,594 Urawa, S., 451

Uziel, M., 698

V Vaidyanathan, C. S., 101, 117(30) Valaguasa, L., 605(274), 608 Valentine, R. C., 506 Valga, S. L., 196 Vallee, B., 378, 379(36, 41, 51), 380(41), 384(41), 387(41), 389(41), 391, 392, 393(96), 394(96), 395(96), 401(36), 402(36, 50, 51), 403, 404(96), 405 (1341, 406(50), 4@3(91, 96), 409(91, 96),410(96), 411(96) Vallee, B. L,173, 517, 532, 619, 723 Vallee, G., 49, 55(33) Valmikinathan, K., 14 Van Belle, H., 432 VanBruggen, J. T., 572 Vandendreische, L., 750, 751(400) Vandendreisache, L., 758 Van den Hende, C., 605(298), 607 Vanderdriesche, L., 750, 751(405) Van Der Werf, P., 150, 151(154) Van Duijn, P., 489 Van Dyck, J. M., 336 Vanecko, S., 284, 290, 299, 303(6), 310 (6, 41) Van Holde, K. E., 504, 706, 707, 708, 709, 727,729 Vanhoucke, A., 772 Van Montagu, M., 750, 751(422) Van Orden, H. O., 181 Vansebw, A. P., 81

Varbanets, L. D., 609 Varga, S. L., 694, 698 Varner, J. E., 2, 8, 13(3), 102 Vasarhely, F., 615, 630(20) Vasilenko, S. K., 313,314(3) Vazquer, J., 433 Veber, D. F., 196,694, 698 Venecko, S., 326 Venetianer, P., 695 Venkina, T. G., 34(76), 39 Venkov, P. V., 320 Venkstern, T. V., 245(137), 249, 321 Ventura, J. M., 247(168, 169, 170), 250 Venugopal, B., 540 Vereb, G., 568, 598(117) Verghese, N., 14 Verity, M. A., 491 Verne, J., 604 Veros, A. J., 113, 114 Vescia, A., 41 Vianna, A. L., 561, 580(104), 587(104), 591(104), 594(104), 595(104)

Vickery, H. B., 125, 132 Vigh-Teichmann, I., 609(355), 610 Vilar, O., 605(287), 606 Villar-Palasi, V., 247(168, 169, 170), 250 Vince, R., 59 Vinograd, J., 260 Viiiuela, E., 593(164, 165), 594, 596, 632, 633(71), 634(71)

Visek, W.J., 13 Vishniac, W., 640(97), 641 Visser, J. P., 699, 700(233), 701(233) Vitali, D. A., 698 Vitali, R. A., 196, 694 Vithayathil, J. J., 675, 676(97), 704(97) Vithayathil, P. J., 196, 669, 670(73), 671 (73), 683, 688, 699, 703(234), 704, 731

Vladescu, C., 605(277), 606 Vladimirova, G. E., 398 Vogel, D., 59 Voigt, B., 71 Vokin, E. J., 321 von Cramm, E., 132 von der Muehll, E., 433 von Ehrenstein, G., 178 von Euler, H., 418 von Heinecke, H., 750, 751(412) yon Hippel, P. H., 154, 185, 735 von Hofsten, B., 493

849

AUTHOR INDEX

Von Kaschnitz, R., 469 von Stedingk, L.-V., 539 yon Tigerstrom, R.G., 328 Vovk, I. N.,605(275), 606

W Wachstein, M., 421, 602 Wacker, W. E. C., 619 Wadal, M. J., 128(66), 129 Wade, H. E., 104, 116(38), 119(38, 84) Wadstrom, T.,183 Waelsch, H.,80, 94, 96, 102, 125, 133, 142(22) Wagle, S. R.,600(194), 601 Wagner, J., 103 Wagner, O., 108, 109(65), 111(65), 112 (651, 113(65), 114(65), 115(65), 116 (651, 118(65) Wagner, R. P.,49, 548 Waitzman, M.B., 368 Wajzer, J., 72 Wakabayasi, Y.,49 Waku, K., 752,759 Walczak, Z., 245(151, 152), 249 Waldschmidt-Leitz, E.,451 Waley, S.G.,125,142 Walker, D. G.,600(191, 1921, 801, 603 (191,192),W(191, 192) Walker, P. G., 392, 393(99), 394, 404(99, 110), 408, 410, 429, 431(117), 433, 435(143), 436(143), 437(117), 438, 439(165), 441(170), 442(117, 1701, 447 Wall, M. C., 19(109), 20 Wallel, H.D., 605(279), 606 Wallenfe’s. K., 605(307), 607 Walli, A. K.,431 Walsh, K.A.,90,91(25) Walter, R.W., 639 Walters, T. L., 363 Walton, E.,6% Walton. G. M., 613 Walz, F.G., 684,759,766 Wang, D.Y.,348 Wang, J. H.,782 Wang. M. H.,69 Wang, S. F.,9 Wang, T. P., 74 Wannamaker, L. W., 261 Wanner, H., 609

Waravdekar, V. S., soO(l86s),801 Ward, D.C.,757 Ward, K. A,, 550, 556 Ward, R.,403 Warner, E.C.,619 Warner, H.,432 Warner, H. R.,268 Warner, R. C., 216 Warnock, L. G.,634 Warren, R.J.,269 Warringer, M., 380 Washko, M. E., 553,557(72),580(72), 674(72), 575(72), 578(71), 590(72), 598(72), 599(71) Wasyl, Z., 476 Watanabe, A., 617, 628 Watanabe, K., 422 Waters, M. D.,422,433 Watson, D.,20 Watters, J. I., 523, 535 Wattiaux, R., 336 Waugh, D., 602 Weakly, D. R., 553, 800(74) Weaver, J. M., 40 Webb, E. C.,3, 4(13), 5(13), 6(13), 10 (13), 12, 15(59), 16(59), 17(59), 19 (13),591 Weber, G., 71, 54.4, 548, 568, 580, 581(7, 153), 582, 596, 597, 598(7), 600(211), 801,602,606(332), 608, 645 Weber, R., 72 Weber, R. E.,734 Webster, H.L.,65, 66(127), 67(127) Webster, R. E.,241, 242(129), 245(128,

129) Wechter, W. J.,315,318(35), 319(34, 35), 320(30, 35) Weetall, H.H., 291, 391 Wegmann, R.,604,605(283,293),608,607 Weidel, W., 374 Weil, A., 132 Weil, J.,254 Weil, L.,685 Weil-Malherbe, H., 432 Weimberg, R.,497 Weimer, H.E.,103 Weinbaum, G.,55, 56(84), 57(84) Weinhouse, S., 593(163), 594, 596, 597 Weiss, B.; 304,305, 306,367

850

AUTHOR INDEX

Weissmann, C., 216 Weitzman, P. D. J., 691 Wellner, D., 677, 678(112) Wellner, V. P., 137 Wenger, B. S., 497 Werner, G., 581(152), 582, 594(152) Werner, S., 581(152), 582, 594(152) West, E. S., 572 West, G. R., 603 Westall, R. G., 132 Westheimer, F. H., 454, 534 Westort, C., 740 Wetlaufer, D. B.,' 505 Wetzel, B. K., 362, 375 Wharton, C . W., 746 Whelan, H. A,, 107, 110, 111(61), 112 (61), 113(61), 114, 115(61), 118(61)

Whitaker, J. R., 55, 90, 91(24) Whitby, L. G., 419(32), 420, 429, 437 (116)

White, F. H., Jr., 648, 681, 691, 693(169a), 694,695 Whitfield, P. R., 215, 216, 217, 239, 244 (113), 331, 363, 746, 747

Whittaker, C. K., 104 Wiberg, J. S., 268 Widnell, C. C., 345, 351(48), 549 Wieczorek, V., 50, 70, 73(155) Wieker, H. J., 775, 790 Wieland, O., 367, 368(45), 626 Wigler, P. W., 318, 777 Wikler, M., 128(68), 129(68), 130 Wilchek, M., 173, 174, 179, 186, 187(61), 188(61), 189(61), 190(61), 192(61), 193(68), 195, 675(102), 677, 678(114), 681(114) Wilcher, M., 333, 334(31) Wiley, C. E., 248(174), 250 Wilkins, M. H. F., 158 Wilkinson, J. M., 128(67), 129(67), 130 Willenbrink, J., 643 Willett, R., 535 William, F. R., 243, 245(133) Williams, A., 404, 410(137) Williams, D. H., 111,44 Williams, E. J., 185, 307, 316, 319(40) Williams, H., 391 Williams, H. E., 603 Williams, M. W., 154, 155(18) Williams, R. W., 59

Williams, W. F., 49 Williams, W. J., 94 Williamson, D. H., 87, 95(16) Williamson, J. R., 347,644 Willighagen, R. G. J., 489 Wilson, A. T., 641 Wilson, H., 125, 131 Wilson, I. B., 377, 392(30), 393(30), 394 (301, 395(105), 397, 398(30, 105), 402, 403, 405(31), 406, 407(121, 1231, 408 (30, 1231, 409(118, 123), 410(30, 123, 413 Wilson, R. J., 43, 46 Wilson, W. E., 134, 634 Winegard, A. I., 614 Wingert, L.,154, 185 Winshell, E. B., 31 Winstead, J. A., 129, 778 Winzler, R. G., 273 Wishnia, A., 723 Wislocki, G. B., 605(316), 607 Witkop, B., 180, 671, 682 Witzel, H., 215, 216, 217, 220, 221(55), 239, 244(113), 712, 724, 775, 781, 790 Wojnar, R. J., 770 Wolberg, H., 450,451,453(3), 455 Wold, F., 129, 683, 696, 699(139), 703 (139), 720(212), 778 Wolf, H. P., 612 Wolf, P. L., 327, 433 Wolfe, H. J., 604

Wolfe, P. B., 366 Wolfenden. R., 51, 52(64, 67, 69), 53(67), 55, 56(92), 59(67), 60(67), 61(113). 62(67, 69), 63(67), 65(92), 70(92). 73, 74 Wolff, J.. 684, 689 Wolna, E., 430, 442(120) Wong, K. Y., 735 Wong, S. C. K., 502, 503(13), 501(13), 505(13), 506(13), 509(13, 30), 510(13, 30), 511(30) Woo, M . V., 313 Wood, W. A., 48, 614, 615, 616(22), 618 (15), 621(15, 161, 622(15), 624(15), 627 (22) Wood, W. F., 77 Woodard, H. Q., 450,496 Woodford, M., 632 633(72), 634(72) Woodhouse, B. A., 656

851

AUTHOR INDEX

Y

Yoshihara, H., 269, 270(75) Yoshino, T., 247(171), 250 Younathan, E.S., 540 Young, B. G.,758 Young, D.A., 594 Young, D. M.,671, 694, 699(89), 703 (89),710 Young, E.T., 11, 279,291 Young, G.T.,126,133(25) Young, W. J., 420 Yount, R. G.,535 Yourke, A.,548 Yphantis, D. A., 388, 503, 504(17), 509 (17), 696, 709 Yu,M.T.,639 Yuan, R.,263,264 Yutani, A., 694 Yutani, K.,694

Yabuta, A,, 536, 540(24) Yagi, T.,75 Yamada, K.,604 Yamada, Y., 227, 230(89) Yamagata, S.,219, 220 Yumagata. T.,216 Yamagishi, S., 29(44), 30, 31(44), 34(44), 38(44), 39(44), 40(44), 41(44), 42 (44),43(44) Yamamoto, M.. 366, 368(38), 369(38) Yamamoto, Y., 219 Yamanaka, M., 261 Yamasaki, M.,240, 244(119, 120, 121) Yamasaki, T.,244(141),249 Yanagida, M.,248(173),250 Yanari, S., 716 Yang, C.-C., 313 Yang. J. T., 505,722 Yang, K.-U., 438, 445(167), 446(167) Yang, S.-T., 739 Yung, Y., 532, 533(13), 535(13) Yankwich, P. E.,18 Yasuda, S.,269, 270(75) Yaks, M.G.,75 Yellin, T. O., 106, 107(56), 118(56) Yonath, A.. 156, 157(42) Yoneda, M.,51, 52(63), 53(63). 77(63). 245(153), 249,307 York. J. L.,59, 60(112), 61(112) Yoshida, K..231,247(105) Toshida, M.,240,244(121) Yoshida, N.. 222(76),223

Zarhau, H. G., 218,223(48) Zak, B.,567 Zakim, D., 544,596(11) Zamenhof, S.,291,304(24), 757 Zamfirescu-Gheorghiu, M.,605(277), 606 Zamir, A.,48, 126, 215, 222(29), 325 Zamoyska, A. M.,654 Zanetti, G.,699,700(230) Zbarsky, S. H.,301 Zderir, J. A.,454 Zerner, B.. 3, 4(13), 5(13), 6(13), lO(13). 12, 15(59), 16(59), 17(59), 19(13). 90, 91(23) Zerr. C.,580(146). 581(146), 582, 602 (146) Zervas. L.,133 Zetterqvist, O.,396, 423 Zielinski, J., 605(300),607 Zielke, C. L.. 65, 66(133), 69(133), 70 (133),71(133), 73(133) Zimmerman, 1,. N., 76 Zimmerman, S. B..267, 292, 722, 750, 751 (421) Zina. G.,605(311), 607 Zinder. N. D..241, 242(129), 245(128 129), 501 Zito R.. 273,274(12) Zittle. C.A,, 102, 436, 442,750,751(397a) Zorzoli, A., 600,602 Zottu. S.,552,600(202),601

Woods, W. D., 368 Woodward, G.E.,124, 142 Woody, R.W., 684,690(144) Wrathall, D. P.,740 Wretlind, A., 129 Wrigglesworth, J. M.,603 Wright, B. E.,640(93),641 Wriston, J. C.,Jr., 102, 104, 105, 106, 107 (56), 108(56, 57), 110, 111(61), 112 (61), 113(61), 114(77), 115(61), 116 (35),118(56,611, 119(35) Wrobel, K., 606(326), 608 Wyckoff, H. W., 163, 196, 656, 657(62), 658, 666(62), 667(62), 669, 683(61), 724(62), 763, 785, 800 Wynne, A. M., 435

Z

852 Zoukis, M., 137 Zucchelli, G., 59 Zucker, M. B., 457,495 Zupanska, B., 548, 605(51, 272, 300), 606, 607

Zuppinger, K., 548

AUTHOR INDEX

Zwaig, N., 380 Zydowo, M., 49, 64, 65(125), 66(125), 67(125), 68(125), 70(125)

Zyk, N., 34(75), 39, 40(75), 41(75), 43, 44(75), 45(75, 101, 1151, 46(74, 101, 115, 120)

Subject Index A Acetate, spleen acid deoxyribonuclease and, 281, 283 Acetazoleamide, 3’,5’-cyclic phosphate diesterase and, 369 Acetic acid, urease dissociation and, 9 Acetic anhydride, alkaline phosphatase and, 391, 427428 fructose diphosphatase and, 631 p-Acetoxymercurianiline, staphylococcal nuclease and, 156 N-Acetylglucosamine, deoxyribonuclease I and, 293, 297 N-Acetyl homocysteine thiolactone, ribonuclease and, 681 Acetylimidazole, fructose-1, 6diphosphatase and, 616, 619620 pyrophosphatase and, 515 ribonuclease TI, and, 221 S-Acetylmercaptosuccinic anhydride, ribonuclease and, 698 Acid phosphatase(s), assay problems, 454 distribution, 450, 451 electrophoretic behavior, 454455, 477, 480481, 485, 486 historical, 450 specificity, 450, 45M54 Acinetobacter, fructose diphosphatase of, 639-640 Acrocylindrium, ribonuclease of, 246 Acrylonitrile, ribonuclease and, 678, 679 Actinomyces aureoventicillatus, ribonuclease of, 245 Actinomycin D, spleen acid deoxyribonuclease and, 281-282 853

Active site, adenosine aminohydrolase, 5 8 5 9 alkaline phosphatase, 404406, 445,446 deoxyribonuclease I, 297-299 glucose-6-phosphatase, 586-587 pyrophosphatase, 526-527 ribonuclease A, 748 ribonuclease TI, 220-221,222 staphylococcal nuclease, size and specificity, 191-195 sterochemical probes, 195-196 urease, 20 Acyl carrier protein, fructose-1, 6diphosphatase and, 624, 626, 631,644 Adenine, adenine aminohydrolase and, 54 adenosine aminohydrolase and, 60 analog’s, absorbancy change by hydrolysis, 52-53 deamination, absorbancy change, 52 Adenine aminohydrolase, distribution of, 49 historical, 48 properties of, 51, 53-54 substrates for, 54, 74 Adenine nucleotide aminohydrolase, properties of, 75-76 Adenine xylofuranosyl 3’,5’-cyclic phosphate, cyclic phosphate diesterase and, 367 A4denosine, adenosine aminohydrolase and, 60, 62 analogs, adenosine aminohydrolase and, 60-61 vasodilation and, 63 deamination, absorbancy change, 52 derivatives, ribonuclease and, 760, 769, 785, 786, 788, 790, 797, 798, 799-800, 803

SUBJECT INDEX

Adenosine (cont.) heart and, 347,348 ribonucleotide 2’,3’-cyclic phosphate diesterase and, 360-361 Adenosine aminohydrolase, catalytic properties, mechanism, 59-63 nature of active site, 58-58 reaction parameters, 5 6 5 7 distribution of, 49-50 kinetic constants, 60-61, 74 molecular properties, chemical and physical properties, 55-56 purification, 54-55 nonspecific, properties, 73-74 physiological function, 63-64 Adenosine 2’,3’-cyclic phosphate, ribonuclease Tzand, 225, 228, 229 ribonuclease U, and, 237 Adenosine 3’,5’-cyclic monophosphate, fructose diphosphatase and, 631 glucose-6-phosphatase and, 598 Adenosine 3’,5’-cyclic phosphorothioate, cyclic phosphate diesterase and, 367 Adenosine diphosphate, fructose-1, 6diphosphatase assay and, 615 pyrophosphatase and, 525,526,538, 540

Adenosine 3’-monophosphate, 5’-nucleotidase and, 342 ribonucleotide 2’,3’-cyclic phosphate diesterase and, 360-361 Adenosine monophosphate, alkaline phosphatase and, 428,430 analogs, 5’-adenylic acid aminohydrolase and, 66 fructose diphosphatase and, 614, 630631, 633, 634, 639, 640, 642, 643, 644, 646 binding sites, 628 conformation and, 629, 636637, 638 papain effect, 619 pH and, 618-619 pyridoxal phosphate and, 620 tyrosine residues and, 619-620 nitrated staphylococcal nuclease and, 173

phosphofructokinase and, 613, 644 phosphoryl alkaline phosphatase and, 405, 413 Adenosine triphosphate, 5’-adenylic acid aminohydrolase and, 65, 68-69, 70 alkaline phosphatase and, 405, 413, 429, 430, 431 deoxyribonuclease and, 259, 261-262, 263, 264 fructose diphosphatase and, 631,639 glucose-6-phosphatase and, 561,573, 576, 594, 595, 596, 599 5’-nucleotidase and, 338-339, 341, 349, 352 phosphofructokinase and, 613 pyrrolidone carboxylate, formation, 136, 137, 139 utilization, 150-151 Adenosine triphosphate-pyrophosphatase, venom exonuclease and, 314 S-Adenosylmethionine, restriction endonuclease and, 263, 264 Adenylate-N-oxide phosphodiester bonds, ribonuclease T I and, 227 5’-Adenylic acid aminohydrolase, catalytic properties, kinetics, 66-70 mechanism, 70-71 physiological function, 71-73 specificity, 66 distribution of, 50 historical, 48 molecular characteristics, chemical and physical properties, 65-66 purification, 64-65 Adenylyl-(3’,5’)-nucleoside, synthesis, ribonuclease U, and, 2&239 Adrenalectomy, glucose-6-phosphate and, 598 Adrenocorticotropic hormone, glucose-6phosphatase and, 598 Aerobacter aerogenes, see also Enterobacter fructose diphosphatase of, 639 5’-nucleotidase of, 340 Aerobacter cloacae, p-lactamase,

SUBJECT INDEX

dissociation constants, 38 inhibition of, 44 Agaricaceae, y-glutamyltransferase of, 95-96 Agaritine, glutamyltransferase and, 95 Agouti, serum, asparaginase in, 103, 107 Alanine residues, fructose diphosphatase, 627 ribonuclease, 658,665, 672-673 Alcohols, acid phosphatase, erythrocyte, 481482 prostatic, 474475 p-lactamases and, 44 phosphotransferase and, 570, 571, 579 ribonuclease and, 779, 791 Aldolase, sedoheptulose 1,7diphosphate hydrolysis and, 615 Alkali, p-lactamases and, 45 Alkaline phosphatase, catalytic properties, competitive inhibitors, 394-396 kinetic studies, 409-415 number of active sites, 404406 phosphoryleneyme, 396-401 role of zinc, 401404 specificity, 392-394 transphosphorylation, 406-409 distribution of, 374-376 function of, 376377 historical background, 373-374 mammalian, assay techniques, 432-434 chemical modification, 427-428 distribution, 420-421 function, 421-422 general survey, 417-420 kinetic studies, 434443 mechanism, 4 4 H 4 7 physical properties, 423-427 purification procedures, 422-423 reaction catalyzed, 430-432 substrate specificity, 428-430 molecular properties, chemical modification, 389-392 composition, 378-380 crystal structure, 389

855 isoeymes, 384-387 physical properties, 387-389 purification, 377-378 subunits, 380-384 staphylococcal nuclease and, 179 Alkyl sulfates, p-lactamases and, 44 Alloxan, glucose-6-phosphatase and, 545, 568, 578, 581 w-Amidase, asparaginase activity, 101 Amides, hydrolysis, asparaginase and, 107 Amide groups, deoxyribonuclease I and, 293 Amidines, adenosine aminohydrolase and, 59 Amines, glutaminases and, 94 Amino acid(s), activation, inorganic pyrophosphatase and, 501 alkaline phosphatase, composition, 378-380, 424-425 inhibition, 442 sequence, 380, 381 asparaginase, composition, 111-113, 115 partial sequence, 116 bovine liver acid phosphatase composition, 492 deoxyribonuclease I, composition of peak A, 293,296 terminal, 297 fructose diphosphatases, 645 fungal ribonucleases, composition, 209 glucose-6-phosphatase and, 579 y-glutamyl transpeptidase and, 96-97 p-lactamases, composition, 31, 33 sequences, 35, 3&37 terminal, 33, 35, 42 5’-nucleotidase activation by, 342 pancreatic ribonuclease, composition, 653 sequence, 655 pyrophosphatase, 531 composition, 512, 513 terminal, 512-514 spleen acid deoxyribonuclease composition, 274

SUBJECT INDEX

Amino acid(s) (cont.) staphylococcal nuclease, sequence, 162, 180 urease composition, 11 a-Aminoadipate, glutamine synthetase and. 137 a-Aminobutyrate, y-glutamylcysteine synthetase and, 137 7-Aminocephalosporanic acid, p-lactamases and, 25, 40 2-Amino-6-chloropurine, adenine aminohydrolase and, 54 Amino groups, alkaline phosphatase, 427 ribonuclease A, 711 modification of, 677-682, 698 ribonuclease TI,219, 221 6-Aminopenicillanic acid, p-lactamases and, 25, 34, 38,40,42 7-Aminothiaaolo [5,4 d l pyrimidine, adenine aminohydrolase and, 54 a-Aminotricarballylate, n-glutamate cyclotransferase and, 135 Aminotyrosine residues, staphylococcal nuclease and, 173 Ammonia, bacterial asparaginase and, 110 glutaminase assay and, 81 glutamyl transferase and, 94,95 liberation, urease and, 4 Ammonium hydroxide, glucose-6phosphatase and, 559-560 Ammonium ions, 5’-adenylic acid aminohydrolase and, 67-69 Amphibia, adenosine aminohydrolase of, 57 Ampicillin, p-lactamases and, 34,a Anions, acid phosphatases and, 466, 477-480 adenine nucleotide arninohydrolase and, 75, 76 5’-adenylic acid aminohydrolase and, 69 ribonuclease urea transition and, 733 Antibodies, 5’-adenylic acid aminohydrolase and, 65 alkaline phosphatase subunits and, 382,390

fructose diphosphatase and, 629-630, 633, 641, 645 /3-lactamase activity and, 45 modified ribonuclease and, 692 prostatic acid phosphatase and, 467, 468 Aorta, phosphatase and, 422 Arabinosides, venom exonuclease and, 320 Arginine residues, alkaline phosphatase, 382-383 replacement of, 390 asparaginase, 116 bovine liver acid phosphatase, 491 fructose diphosphatase, 633 ribonuclease A, 665, 669, 676, 711, 725, 739, 770, 785 modification of, 689-690,704 and, 221, 222 ribonuclease TI, spleen acid deoxyribonuclease, 276 staphylococcal nuclease and, 167, 171, 197-198, 203 Arsenate, acid phosphatases and, 477,480,497 alkaline phosphatase and, 394, 395, 396-397, 405, 442 glucose-6-phosphatase and, 581 ribonuclease and, 785 Asparaginase(s), bacterial, amino acid composition, 111-113, 116-119 general properties, 109-110 isolation, 107-108 physiological properties, 117-121 structure, 113-116 substrate specificity and inhibitor effects, 11e111 guinea pig serum, 102 amino acids, 113 isolation, 105-106 properties, 106-107, 118-119 lymphosarcoma and, 102 occurrence, 102-105 Asparagine, glutaminase and, 94, 97 staphylococcal nuclease aspartate residues and, 203 Asparagine residues, asparaginase, 115

857

SUBJECT INDEX

ribonuclease, 651, 665,699, 787-788, 794 spleen acid deoxyribonuclease, 275 Asparagine synthetase, asparaginase sensitivity and, 118 Aspartate residues, deoxyribonuclease I, 293, 297 ribonuclease, 665,676, 677, 702, 7 1 6 717, 739, 763, 788, 794,800,803 staphylococcal nuclease, 167, 171, 173, 203 p-Aspartylhydroxamate, asparaginase and, 107 Aspergillus nigh-, asparaginase, 103 ribonuclease of, 246 Aspergillus oryzae, nonspecific adenosine aminohydrolase, 73-74 ribonucleases, main properties, 208, 246 Aspergillus satoi, ribonuclease of, 246 Auriculoventricular block, adenosine and, 63 Autoretardation, deoxyribonuclease I and, 303-304, 307 Axial ratio, ribonuclease, 704,710 8-Azaadenosine, adenosine aminohydrolase and, 60,63 8-Azaguanine, adenosine aminohydrolase and, 58 deamination, absorbancy change, 53 guanine aminohydrolase and, 76, 77 Azaserine, glutaminase and, 98 Azatryptophan, alkaline phosphatase and, 390 Azotobacter agilis, glutaminase of, 85, 94, 97-98 ribonuclease of, 240, 244 Azotobacter vinelandii, adenine aminohydrolase of, 51, 53, 54 urease of, 14

B Bacillus amyloliquefaciens, ribonuclease of, 244 Bacillus cereus, adenosine aminohydrolase of, 55 p-lactamase, 26 amino acid composition, 33

conformation, 45 dissociation constants, 38 molecular properties, 31, 32 purification, 28, 30 substrates, 34, 40, 41 temperature and, 43, 44 thiol reagents and, 4 3 4 4 ribonuclease of, 244 Bacillus coagulans, asparaginase of, 104, 116, 117, 119 Bacillus licheniformis, p-lactamase, amino acid composition, 33, 42 amino acid sequence, 35, 36-37 catalytic constants, 41 dissociation constants, 38 molecular properties, 32 mutations and, 42 purification, 28 substrates, 34 Bacillus megaterium, pyrophosphatase of, 539 Bacillus pasteurii, urease of, 14, 19 Bacillus pumilus, ribonuclease of, 244 Bacillus subtilis, deoxyribonuclease, 253 bacteriophage SP-3-induced, 258-259 glutamyl transferase of, 94 3’-nucleotidase of, 354 5’-nucleotidase of, 340, 349, 350 pyrophosphatase of, 539 pyrrolidone carboxylyl peptidase of, 148, 149 ribonuclease, extracellular, 239-240 intracellular, 240 ribonucleoside 2’,3’-cyclic phosphate diesterase, 356, 357 kinetic constants, 360 physical and chemical properties, 358, 359 substrate specificity, 357, 358 Bacteria. asparaginase in, 103-104 fructose diphosphatases of, 639-640 guanine aminohydrolase in, 50 intestinal, D-ghtamate and, 136 5’-nucleotidase of, 338-340 Bacteriophage k, deoxyribonuclease induced by, 253-254

SUBJECT INDEX

Bacteriophage h (cont.) deoxyribonucleic acid, endonucleases and, 263, 267, 268, 280, 304 Bacteriophage R-17, ribonucleic acid, terminal identification, 326-327 Bacteriophage SP-3, deoxyribonuclease induced by, 258-259 Bacteriophage T2, deoxyribonucleic acid, venom exonuclease and, 316 Bacteriophage T2 and T4, deoxyribonuclease induced by, 255 Bacteriophage T4, deoxyribonucleic acid, spleen exonuclease and, 333 endonucleases induced by, 266-269 Bacteriophage T5, deoxyribonuclease induced by, 261 Bacteriophage T7, endonuclease induced by, 265-266 Bacteriophage(s) T-even, 3’-deoxynucleotidase induced by, 354 Barium, staphylococcal nuclease and, 157, 163 Base(s), ribonuclease and, 754-758, 760, 781, 782 Bentonite, urease and, 12 Benzylcephalosporin, p-lactamases and, 34 Benzylpenicillin, p-lactamases and, 38, 4243, 4&46 Beryllium chloride, ribonuclease and, 772 Beryllium ions, alkaline phosphatase and, 440-441 Bicarbonate, glucose-6-phosphatase and, 580,581, 587, 594, 596 Bile acids, alkaline phosphatases and, 419 glucose-6-phosphatase and, 581 Birds, adenosine aminohydrolase of, 57 serum, asparaginase in, 103 Blood, glucose-6-phosphatase in, 548 Blood vessels, 5’-nucleotidase in, 347-348 Blowfly, adenosine aminohydrolase in, 49 Blue-green algae, urease in, 14 Bone, acid phosphatase of, 450,49f397

alkaline phosphatase of, 418,419,420, 421, 423 stability, 426 substrates, 428 kinetic studies, 438 Borate, acid phosphatase and, 497 glucose-6-phosphatase and, 581 Borohydride, ribonuclease and, 691 yeast pyrophosphatase and, 531 Bothrops atrox, 5’-nucleotidase of, 342, 350, 352 venom exonuclease, 318 Bovine, tissues, glucose-6-phosphatase in, 600, 604, 605, 606 Bovine liver, acid phosphatase of, 491493 Brain, adenosine aminohydrolase of, 55 adenosine monophosphate aminohydrolase in, 50 5’-adenylic acid aminohydrolase of, 65-66 activation, 67, 68 specificity, 67 alkaline phosphatase of, 423 3’,5‘-cyclic phosphate diesterase of, 366-368, 370 glucose-6-phosphatase in, 596 y-glutamyl cyclotransferase of, 143, 144-146 guanine aminohydrolase in, 51, 76-77 9-(Bromoacetamidobenzyl) adenine(s), adenosine aminohydrolase and, 58 Bromoacetamidophenyl derivatives, staphylococcal nuclease, 195 Bromoacetate, ribonuclease A, histidine residues, 686-687, 688 lysine residues, 682 methionine residues, 683 ribonuclease T, and, 229 a-Bromo-n-butyrate, ribonuclease and, 687 a-Bromocaproate, ribonuclease and, 687 Bromocresol green, glutaminases and, 87 a-Bromopropionates, ribonuclease and, 687

SUBJECT INDEX

8-Bromopropionate, ribonuclease and, 687

p-Bromopyruvate, ribonuclease and, 687 N-Bromosuccinimide, alkaline phosphatase and, 391 5’-nucleotidase and, 341 prostatic acid phosphatase and, 471 ribonuclease TI, and, 221 spleen acid deoxyribonuclease and, 281 a-Bromovalerate, ribonuclease and, 687 Buffers, alkaline phosphatase and, 436,437438 urease activity and, 19-20 Bull seminal plasma, 5’-nucleotidase of, 342-343, 351

1,4-Butanediol, prostatic acid phosphatase and, 472473,474 Butanolamine, alkaline phosphatase and, 406

C Cadmium ions, alkaline phosphatase and, 402,405 yeast pyrophosphatase and, 535 Caffeine, 3’,5’cyclic phosphate diesterase and, 368, 369 Calcium ions, acid phosphatase and, erythrocyte, 479 prostatic, 466, 479 deoxyribonucleases and, 290, 297,302303

5’-nucleotidase and, 352 pyrophosphatase and, 532, 535 ribonuclease and, 770 staphylococcal nuclease, 154, 155, 157, 163, 186187, 190, 191, 192

binding of, 163-171, 174, 203 Canavalia ensijormis, urease, isolation and purification, 2-5 molecular properties, 5-13 Canavanine, alkaline phosphatase and, 390

Candida utilis, adenine aminohydrolase in, 49,53-54 fructose diphosphatase, inhibition by adenosine monophosphate, 636-637

purification and properties, 635-636 relation to sedoheptulose diphosphatase, 638 structure, 637-638 Caprylohydroxamate, urease and, 16 Carbamate, urease and, 15-16 Carbamyl phosphate, glucose-6phosphatase and, 547,559,567,568, 569-570, 573, 576, 590,591, 592,595, 596, 599

Carbohydrate, deoxyribonuclease I and, 293 ingestion, plasma phosphatase and, 422

ribonuclease Tzand, 224-225 N-Carboxyanhydrides, ribonuclease, 678, 680-681

Carboxyl groups, ribonuclease A, 711 modification of, 675-677 ribonuclease TI and, 219-220 Carboxylic acids, 5’-adenylic acid aminohydrolase and, 70 Carboxymethyl cellulose, urease and, 6 N-Carboxymethylisatoic anhydride, 5’+ 3’ exonuclease and, 258 Carboxypep tidase, comparison to glutaminase, 90,91 pepsin inactivated ribonuclease and, 673

pyrophosphatase and, 514 ribonuclease-S and, 671 urease and, 12 Carboxypeptidase A, asparaginase and, 115 ribonuclease TI and, 222 staphylococcal nuclease and, 181 0-Carboxyphenyl phosphate, alkaline phosphatase and, 393, 394, 442 Carrot, phosphatase of, 473 Castor beans, fructose diphosphatases of, 641-642, 643 Cat, tissues, glucose-6-phosphatase in, 600, 604

Cations, adenine nucleotide aminohydrolase and, 75 divalent, glucose-6-phosphatase and, 580, 587, 590, 591,592

SUBJECT INDEX

Cavioidea, serum, asparaginase in, 103 Cell membrane, p-lactamases and, 45 Cephalosporic acid, unstable, 27 Cephaloridine, p-lactamase and, 34, 38 Cephalosporin(s), #J-lactamases and, 24, 25, 27 Cephalosporin C, p-lactamases and, 34, 38,42, 43 Cetavlon, glucose-6-phosphatase and, 557 Cetyldimethylbenzylammoniumchloride, glucose-6-phosphatase and, 557 Cetyltrimethylammonium bromide, glucose-6-phosphatase and, 557, 558559, 561, 563, 580, 595 Chalaropsis, ribonuclease of, 246 Chaos chaos, acid phosphatase of, 498 Chelators, fructose diphosphatase and, 644,646 glucose-6-phosphatase and, 580,587 yeast pyrophosphataae and, 532-534 Chicken, 5’-nucleotidase of, 343-344, 352 tissues, glucose-6-phosphatase in, 600605 Chloramphenical, urease and, 17 Chloride ions, ribonuclease and, 769, 778,801 ribonucleotide 2’,3’-cyclic phosphate diesterase and, 358, 359 Chlorine-starch-iodide, pyrrolidone carboxylate detection and, 126 Chloroacetate, ribonuclease and, 687 2-Chloroadenosine, heart block and, 63 6-Chloroadenosine, adenosine aminohydrolase and, 60,62 2-Chloroethanol, ribonuclease and, 734 p-Chloromercuribenzenesulfonate, staphylococcal nuclease and, 156 p-Chloromercuribenzoate, p-lactamases and, 44 urease derivative and, 12 pChloromercuripheny1 sulfonate, glucose-6-phosphatase and, 587 Chloromerodrin, u r e a and, 17 Chloroplasts, fructose diphosphatase in, 641,642,643 pyrophosphatase of, 540

Chlorpromazine, 3’,5’-cyclic phosphate diesterase and, 368, 369 6-Chloropurine, adenine aminohydrolase and, 54 Chlorthalidone, 3’, 5’-cyclic phosphate diesterase and, 369 Cholate, glucose-6-phosphatase and, 557, 580 Cholesterol, glucose-6-phosphatase and, 554 Chromatium, fructose diphosphatase in, 642 Chromatography, pyrrolidone carboxylate detection and, 126 Chromium ions, acid phosphatases and, 479 Chymotrypsin, comparison to glutaminase, 90, 91 ribonuclease A and, 674 ribonuclease T, and, 222 staphylococcal nuclease and, 181 Chymotrypsinogen B, deoxyribonuclease I and, 292 Circular dichroism, pyrophosphatase, 505 ribonuclease, 722 Citrate, fructose diphosphatase and, 639 glucose-6-phosphatase and, 568, 580, 581, 586, 594 phosphofructokinase and, 613 Citrobacter, cyclic diesterase of, 362 Citrulline, staphylococcal nuclease arginine residues and, 203 Citrus fruits, acid phosphatase of, 451 Clearance rate, asparaginase antitumor effectiveness and, 120 Clostridia, glutaminase of, 87 Clostridium acetobutylicum, ribonuclease of, 244 Clostridium welchii, glutaminase of, 95 Cloxacillin, p-lactamases and, 34, 38 Cobalt ions, acid phosphatases and, 479 alkaline phosphatase and, 391, 401-402, 403,404, 405,408, 410-411,427, 440, 444 deoxyribonuclease I and, 303 5’-nucleotidase and, 339, 341, 342, 344, 345, 350-351, 352

SUBJECT INDEX

pyrophosphatase and, bacterial, 51%519, 520, 540-541 yeast, 535 ribonucleotide 2’,3’-cyclic phosphate diesterases and, 358, 359 Cobalt sulfide, alkaline phosphatase localization and, 433 Coenzyme A, fructose-l,6-diphosphatase and, 624 625626, 631, 644 synthesis of, 230 Conformation, glutaminase mechanism of action and, 92-93 ribonuclease, 726726 added electrolytes, 735740 organic solvents, 733-735 thermal and acid transitions, 726-731 thermodynamics and, 740-744 urea transition, 731-733 Conformational change, fructose diphosphatase, 629 Copolyglutamate: tyrosine, ribonuclease and, 759 Copper ions, acid phosphatase and, Gaucher’s disease, 496 spleen, 494 alkaline phosphatase and, 402, 403 fructose-1,Cdiphosphatase activation and, 624 ribonuclease and, 770-772 Cortisone, glucose-6-phosphatase and, 556, 560 Corynebacterium renale, urease of, 14, 19 Crayfish, adenosine aminohydrolase in, 49 glucose-6-phosphatase in, 606 Cresol red, urease detection and, 5 p-Cresyl phosphate, alkaline phosphatase and, 398 Crotalus adamanteus, 5’-nucleotidase of, 342 venom exonuclease, 318 Crystal structure, alkaline phosphatase, 389 Cyanate, alkaline phosphatase and, 428 pyrophosphatase and, 516-517 -. ribonuclease and, 679

Cyanide, alkaline phosphatase and, 401, 426 fructose diphosphatase and, 635 p-Cyanoalanine, asparaginase and, 111 Cyanogen bromide, pyrophosphatase and, 514 ribonuclease and, 683 staphylococcal nuclease and, 180-181, 182, 198 Cyanosulfonium salt, ribonuclease and, 682 Cyanuric fluoride, ribonuclease and, 685 Cyclic phosphates, fungal ribonucleases and, 206, 208 ribonuclease and, 746 Cyclic phosphodiesterase, substrates of, 333, 334 N-Cyclohexyl-N’-p-( 4-methylmorpholinium) ethylcarbodiimide p-toluenesulfonate, derivatives, ribonuclease U2and, 237, 238 uridine derivative, venom exonuclease and, 321 water-soluble ribonuclease adducts, 675, 676-677 Cystamine, fructose diphosphatase and, 622-623, 634 occurrence of, 623 Cysteamine S-phosphate, acid phosphatase and, 452, 453 alkaline phosphatase and, 407, 413, 430, 452, 453 Cysteine, acid phosphatase and, prostatic, 469 spleen, 494 alkaline phosphatase and, 427,442 glucose-6-phosphatase and, 568, 579 spleen acid deoxyribonuclease and, 281 Cysteine residues, adenosine aminohydrolase, 74 5’-adenylic acid aminohydrolase, 65 fructose-l,6diphosphatase, 621-622 p-lactamase, 31, 43, 44 pyrophosphatase, bacterial, 508-509, 511, 512, 515 yeast, 531, 534 ribonuclease, 665

SUBJECT INDEX

Cystine residues, alkaline phosphatase, replacement of, 390-391 asparaginase, 111, 116 fungal ribonncleases, 209-210 ribonuclease, 658 urease, 11-12 Cytidine, adenine aminohydrolase and, 54 derivatives, ribonuclease and, 760, 762, 764, 765, 767, 771, 774-777, 785, 786, 787, 788, 790, 792, 794, 797800, 801-806 Cytidine-?-bensyl phosphate, ribonuclease and, 747, 758 Cytidine-Y,3’-cyclic phosphate, ribonuclease and, 749, 758 Cytidine diphosphate, glucose-6phosphatase and, 573,574-575, 578 Cytidine 2’-monophosphate, ribonuclease and, 673, 761, 762 Cytochrome c, spleen acid deoxyribonuclease and, 281, 283 Cytosine, adenine aminohydrolase and, 54 ribonuclease and, 760

D 7-Deazaadenylyl (3’,5’) uridine, ribonuclease T2and, 227 ribonuclease U, and, 236 Deoxyadenosine, adenosine aminohydrolase and, 57,60, 63 Deoxyadenylate-thymidylate polymer, deoxyribonuclease I and, 309 Deoxychola te, glucose-6-phosphatase and, 553, 554, 556, 557-558, 560, 580 5’-nucleotidase and, 345, 347, 348 Deoxycytidylate, T4 endonuclease I V and, 268 2-Deoxy-o-glucose, glucose-6phosphatase and, 570, 571,579 Deoxyribonuclease(s), acid, species and, 285 substrates of, 333, 334,335 adenosine triphosphatedependent, 259, 261-262, 263, 264

bacteriophage induced, 253-254 A, 253-254 T2 and T4, 255 T5, 261 T7, 265-266 classification of, 251-252 endonucleolytic, nonspecific, 259-262 specific, 26Z270 Escherichia coli, 253, 254-255 lysosomes and, 271, 286 3’-nucleotidase and, 353 specificity of, 291 spleen, components, 275 dimeric structure, 275-276 distribution, localization and role, 285-287 general catalytic properties, 280-283 general features of degradation, 276-278 isolation, 27Z273 mechanism, 27g280 methods of investigation, 278 physical and chemical properties, 273-275 specificity, 28S285 Deoxyribonuclease I , active center, 297-299 chemical nature, 292-297 historical, 291 inhibitor, naturally occurring, 299-302 ions and, 302-303 kinetics of, 303-308 methodology and, 289-290 physical and chemical characteristics, 298 physiological role, 310 specificity of, 302, 307, 308-310 Deoxyribonuclease 11, bacteriophage T4-induced, 267-268 Deoxyribonuclease IV, bacteriophage T4-induced, 268-269 Deoxyribonucleic acid, degradation by spleen acid deoxyribonuclease, general features, 276-278 mechanism of initial reaction, 278-280 foreign, degradation of, 287 modified staphylococcal nuclease and, 173

SUBJECT INDEX

native and denatured, staphylococcal nuclease and, 185 native, spleen acid exonuclease and, 332 repair synthesis, 256, 310 sequences, staphylococcal nuclease and, 178 structure, 158 synthesis, adenosine aminohydrolase and, 64 inorganic pyrophosphatase and, 501 venom exonuclease and, 319 Deoxyribonucleic acid ligase, deoxyribonuclease I and, 290, 304, 306 terminal identification and, 327 T4 endonuclease I1 ansay and, 267 Deoxyribonucleic acid polymerase, associated exonucleases, 255-256 3’ 5’ exonuclease, 256 5’-3’ exonuclease, 256-258 2’-Deoxyribonucleotides, ribonuclease and, 752 Desulfovibrio desuljuricans, adenine nucleotide aminohydrolase, 74, 75-76 Detergents, glucose-6-phosphatase and, 556-560, 566, 578, 580, 581, 599 inhibitor action, 560-562, 595 prostatic acid phosphatase and, 459, 460 Deuterium, ribonuclease and, 657 Deuterium oxide, glutaminase and, 90, 93 Development, glucose-6-phosphatase and, 568 Diabetes, fructose diphosphatase and, 645 glucose-6-phosphatase in, 559, 568, 596-597, 598, 599 2,6-Diaminopurine, adenine aminohydrolase and, 54, 58, 59 Diazoacetoglycinamide, ribonuclease and, 675, 676 Diazobenzene sulfonate, frutose-1,6-diphosphatase and, 620 prostatic acid phosphatase and, 471 p-Diazobenzoate, ribonuclease and, 698, Diazomethane, formation, glutaminase and, 86-87 Diazonium derivatives, staphylococcal nuclease and, 195

-

863 Diazonium salt, alkaline phosphatase localization and, 433, 434 5-Diazonium-1H-tetrasole, alkaline phosphatase and, 391 pyrophosphatase and, 515, 517-518 ribonuclease and, 689 ribonuclease T, and, 221 6-Diazo-5-oxonorleucine, glutaminase and, 82,85-87,93,98 5-Diazo-4-oxo-~-norvaline, asparaginase and, 110-111 Diazoxide, 3’, 5’-cyclic phosphate diesterase and, 369 Diborane, pyrrolidone carboxylate reduction and, 127 NB-2’-O-Dibutyrylcyclic adenosine monophosphate, cyclic phosphate diesterase and, 367 Dictyostelium discodeurn, 3’, 5’-cyclic phosphate diesterase in, 366, 368 Di-p-dinitrophenyl phosphate, venom exonuclease and, 319 Diesterases, comparison to staphylococcal nuclease, 188-189 Diet, fructose diphosphatase and, 645 glucose-6-phosphatase and, 598 Diethylaminoethyl cellulose, urease and, 3 Diffusion coefficient, pyrophosphatase, 504 subunits, 509 ribonuclease, 704, 708 Diffusion constant, urease, 10, 12 1,5-Difluoro-2,4-dinitrobenzene, ribonuclease and, 696 Digitonin, glucose-6-phosphatase and, 553, 554, 580 Dihydrouracil, ribonuclease and, 755 5,6 Dihydrouridine-2’( 3’) -phosphate ribonuclease and, 763 5,6-Dihydrouridylate, ribonuclease TZ and, 227, 230 Dihydroxyurea, urease and, 16-18 Diisocyanate, ribonuclease and, 697 Diisopropylfluorophosphate, alkaline phosphatase and, 443 prostatic acid phosphatase and, 471-472 pyrophosphatase and, 515

864 Diketene, ribonuclease and, 680 Dimerization, alkaline phosphatase, 375, 382 Dimethoxybenzoylpenicillin, p-lactamase catalytic constants and, 41 Dimethyl adipimidate, ribonuclease and, 6%697 Dimethylaminonaphthalene-5sulfonylchloride, bovine liver acid phosphatase and, 491 ribonuclease and, 695, 698 Dimethylformamide, ribonuclease and, 733 Dimethylguanylate, ribonuclease T2and, 228 N2-Dimethylguanylyl-(3’, 5’)-cytidine3’ phosphate, venom exonuclease and, 321 Dimethyl sulfoxide, asparaginase and, 111 ribonuclease and, 733 urease and, 17 1,3-Dimethyl urea, adenosine aminohydrolase and, 58-59 Dinitrofluorobenzene, see Fluorodinitrobenzene 2,4-Dinitrophenol, fructose diphosphatase and, 631 Dioxane, alkaline phosphatase and, 438439 liver acid phosphatase and, 488 ribonuclease, 779, 780 acid transition, 734 fluorescence, 718-719 Dipeptides, p-lactamase and, 43 1,3-Diphosphoimidazole, ribonuclease and, 698 Diplococcus yneitmoniae, deoxyribonuclease of, 253, 261, 279 Diplotomic mechanism, spleen acid deoxyribonuclease and, 278-279 Dissociation constants, staphylococcal nuclease, 188, 190-191 Disulfide bonds, deoxyribonuclease I and, 297 ribonuclease A, 659, 690 fluorescence and, 718 oxidation, 691, 693 reduction and protection, 691-693

SUBJECT INDEX

reoxidation and refolding, 693-696 ribonuclease TIand, 218-219 spleen acid deoxyribonuclease, 274 urease, 12 venom exonuclease and, 318 Disulfide interchange, enzymic, 695 fructose-l,6-diphosphatase, 634 coenzyme A and acyl carrier protein, 623-624 cystamine, 622-623 homocystine, 624-625 physiological regulation and, 625-626 5,5’-Dithio-bis(2-nitrobenzoate), fructose-1,6-diphosphatase and, 623, 633 pyrophosphatase and, 512 urease and, 11 Dithioerythritol, 5’-adenylic acid aminohydrolase and, 70 Dithiothreitol, fructose-1,6-diphosphataseand, 625 glucose-6-phosphatase and, 568 glutaminase and, 87 pyrophosphatase and, 510-511,531 urease assay and, 4 Dog, glucose-6-phosphatase in, 600-604 Drosophila melanogaster, acid phosphatase of, 498 adenosine aminohydrolase in, 49 Duck, tissues, glucose-6-phosphatase in, 600,604 Duodenum, adenosine aminohydrolase, 54-56 active site, 58 kinetic properties, 57, 60-61 mechanism, 59, 61, 62

E Edman degradation, pyrophosphatase, 512-514 staphylococcal nuclease and, 181 Ehrlich ascites tumor cells, 5’-nucleotidase of, 348-349, 351 Elastase, ribonuclease and, 670, 672673 Electrolytes, ribonuclease conformation and, 735-737

SUBJECT INDEX

Electron density maps, validity of, 157-158

Electron microscopy, alkaline phosphatase, 389 pyrophosphatase, 506-508 Electron paramagnetic resonance, ribonuclease, 725 Electrophoresis, acid phosphatases, 454-455, 468-469, 477, 480-481, 485, 486

asparaginase, 114 Endomyces, ribonuclease of, 246 Endonuclease ( s ) , exonuclease activity of, 290 snake venom, 314, 328 Endoplasmic reticulum, glucose-6phosphatase in, 562, 586 Enterobacter, see also Aerobacter cyclic diesterase of, 362 Enterobacter cloacae, 8-lactamase, 31 amino acid composition, 33 purification of, 29 substrates, 34 substrate modifications, 40 Enterobacteriaceae, ribonucleotide 2’,3’-cyclic phosphate diesterase, 355, 358, 359, 362

Epinephrine, fructose diphosphatase and, 631 Erwinia aroideae, asparaginase of, 105, 116, 119

Erwinia carotovora, asparaginaae of, 116117, 119

Erythrocytes, acid phosphatase, 450, 451, 457, 469 general properties, 477 purification and separation of genetic types, 477-484 5’-adenylic acid aminohydrolase, activation, 67-69 Escherichia coli, acid phosphatases of, 498 adenine aminohydrolase in, 49 adenosine aminohydrolase of, 55 adenosine triphosphate dependent deoxyribonuclease, 259 alkaline phosphatase, 453 chemical modification, 389-392 competitive inhibitors, 394-396

865 composition, 378380 crystal structure, 389 distribution, 374-376 function, 376-377 historical background, 373-374 isozymes, 384-387 kinetic studies, 409-415 number of active sites, 401406 phosphoryl enzyme, 396-401 physical properties, 387-389 purification, 377-378 role of zinc, 401-404 specificity, 392-394 subunits, 380-384 transphosphorylation, 406-409 asparaginase, 102, 104 isloation, 107-108 properties, 109-116, 118-119 3’, 5’-cyclic phosphate diesterase of, 366,368, 369

3’-deoxynucleotidase in, 354 deoxyribonuclease IV, 254-255 deoxyribonucleic acid ligaae, 267 deoxyribonucleic acid polymerase, exonucleases associated with, 255-258

endonuclease I of, 259-260,279 endonuclease I1 of, 264-265 exonucleases I and 111, 253 fructose diphosphatase, 638-639 mutants and, 614, 639 glutaminase, 80-81 acyl transfer reactions, 84-85 aasay, 81 deuterium oxide and, 90 6diazo-5-oxonorleucine and, 85-87 mechanism of action, 90, 92-93 occurrence, 81 other inhibitors and, 87 p H effects on kinetic parameters, 88, 89

purification, 82 relationship to other acylases, 90, 91 specificity, 82-84 temperature and, 88 hydrolases, localization of, 287 inorganic pyrophosphatase, 500 chemical composition, 512-514 chemical modification, 514-518

SUBJECT INDEX

Escherichia coli, inorganic pyrophosphatase (cont.) effects of ions and inhibitors, 518-519 electron microscopy, 506-508 interaction with inhibitors, 525-526 homogeneity, 502-503 nature of substrate binding, 522-525 pH effects, 518 physical properties, 504-506 purification, 501-502 reconstitution, 510-512 reversal of reaction, 519-520 size, 504 substrate specificity and stoichiometry, 520-522 subunits in guanidine hydrochloride, 508-510 5‘-nucleotidase of, 338,349,350 penicillinase, 23-24 amino acid composition, 33 dissociation constants, 38 molecular properties, 31, 32 purification, 29 substrates, 34, 40, 41 restriction endonucleases of, 263-264 ribonucleases of, 241-243, 244-245 ribonucleoside 2’, 3’-cyclic phosphate diesterase, 356, 357 cellular localization, 361362 kinetic constants, 360 metals and, 362-363 physical and chemical properties, 358, 359 substrate specificity, 357 Ethacrynic acid, 3’,5’-cyclic phosphate diesterase and, 369 1,2-Ethanediol, urease and, 8 Ethanol, alkaline phosphatase and, 438-439 fructose diphosphatase and, 631 prostatic acid phosphatase and, 472 ribonuclease and, 744 Ethanolamine, alkaline phosphatase and, 398,407, 408, 410, 411 Ethidium bromide, endonuclease I and, 260

Ethoxyformic anhydride, ribonuclease and, 698

Ethyl acetate, pyrrolidone carboxylic acid extraction and, 125 l-Ethyl-3-( 3dimethylaminopropyl) carbodiimide, ribonuclease and, 677 Ethyl disulfide, fructose diphosphatase and, 634 Ethylenediamine platinum I1 dichloride, ribonuclease and, 683 Ethy lenediaminetetraacetate, acid phosphatase and, erythrocyte, 482 spleen, 494 adenosine aminohydrolase and, 56 alkaline phosphatase, bacterial, 374-375, 378, 385,401 mammalian, 426427, 443 deoxyribonuclease I and, 297 fructose-l,6-diphosphataseand, 624, 630, 632, 636,640, 646 glutarninase and, 87, 98 y-glutamyl transpeptidase and, 96 p-lactamases and, 44 5’-nucleotidase and, 339-340, 341, 342, 343, 345, 346,348, 350-351, 352 ribonucleotide 2’, %-cyclic phosphate diesterase and, 359, 360, 361, 362 spleen acid deoxyribonuclease and, 281,283 spleen exonuclease and, 335 urease and, 3, 4, 9 yeast pyrophosphatase and, 532-534 Ethylene glycol, ribonuclease conformation and, 733-734, 740 Ethyleneimine, ribonuclease and, 691 y-Ethyl glutamate, cyclization, ammonolysis and, 133 N-Ethylmaleimide, adenosine aminohydrolase and, 58 alkaline phosphatase and, 428 fructose-1,6-diphosphatase and, 622 pyrophosphatase and, bacterial, 509, 510, 515 yeast, 531 urease and, 10, 11 Ethyl phosphate, alkaline phosphatases and, 428 Ethyl thioltrifluoroacetate, ribonuclease and, 679 Euglena aracilis. fructose diphosphatase of, 641, 642

867

SUBJECT INDEX

ribonuclease of, 248 Exonuclease(s), endonuclease activity, 315, 321 snake venom, 314

F Fasciola hepatica, 3’,5’-cyclic phosphate diesterase in, 366 Fast blue RR (or BB), alkaline phosphatase localization and, 434 Fasting, glucose-6-phosphatase and, 568, 598, 599 Fat, ingestion, plasma phosphatase and, 422 Fatty acid(s), activation, inorganic pyrophosphatase and, 501 fructose diphosphatase and, 631,643 glucose-6-phosphatase and, 579 Feces, alkaline phosphatase in, 423 Ferrodoxin, fructose diphosphatase and, 643 Fibrinogen, pyrrolidone carboxylate in, 128, 130, 149 Fibroblasts, alkaline phosphatase of, 422 skin, acid phosphatase of, 454 Fish, adenosine aminohydrolase of, 57 5’-adenylic acid aminohydrolase, 64, 65 activation of, 68 kinetic constants, 67 eggs, adenosine monophosphate aminohydrolase in, 50 glucose-6-phosphatase in, 600 Flavin adenine dinucleotide, 5’-nucleotidase and, 341 Fluorescence, acid phosphatase assay and, 454, 457 alkaline phosphatase assay and, 433 pyrophosphatase, 506, 533 ribonuclease, 710, 718-719 thermal transition and, 727 urea transition and, 733 urease, 10-11 Fluoride, acid phosphatase and, erythrocyte, 482

fungal, 497 liver, 487 plant, 497 prostatic, 459-462, 477, 480 spleen, 494 glucose-6-phosphatase and, 545, 581 pyrophosphatase and, 519 ribonuclease and, 778 l-Fluoro-2,4-dinitrobenzene, adenosine aminohydrolase and, 55 fructose-l,6diphosphatase and, 621622, 633,637,638 liver acid phosphatase and, 48E-489, 490 ribonuclease A and, 678,680 ribonuclease TIand, 221 staphylococcal nuclease fragments and, 181 Fluorophenylalanine, alkaline phosphatase and, 389-390, 443 Fluorophosphate, alkaline phosphatase and, 429 Fluorotryptophan, alkaline phosphatase and, 390 Formaldehyde, acid phosphatases and, 477, 482, 495, 496 alkaline phosphatase and, 427 Formanidase, 1 Formamide, asparaginase and, 114 ribonuclease and, 733, 779 Formic acid, ribonuclease and, 696 Formycin, ribonuclease and, 757, 789, 797, 799 Formyltetrahydrofolate deformylase, 1 French press, glucose-6-phosphatase and, 560 Frictional ratio, ribonuclease, 704 Frog, 5’-nucleotidase of, 344 tissues, glucose-6-phosphatase in, 600, 602 Fructose, glucose-6-phosphatase and, 546, 583, 598 Fructose-l,6-diphosphatase, activity of, 594 assay and mechanism of action, 615-616

868

SUBJECT INDEX

Fructose-l,6diphosphatase (cont.)

Candida utilis, 646 inhibition by adenosine monophosphate, 636-637 purification and properties, 635636 relation to sedoheptulose diphosphatase, 638 structure, 637-638 comparative properties, 645846 historical, 61!?-613 kidney, 645 purification and properties, 629-630 regulation, 630-631 liver, 645, 646 disulfide exchange, 622-626 molecular structure, 626-629 proteolysis and, 618 purification and properties, 61-18 regulation of, 618-620 sulfhydryl groups, 621422 liver acid phosphatase and, 487 microorganisms, 638-640 muscle, 645 evidence for, 632 physiological role, 634-635 purification and properties, 632-633 structure and relation to other diphosphatases, 633-634 physiological role of, 644-645 plants, 642-643, 645 regulation and physiological function, 613-615

slime mold, 640,646 Fructose-l,6diphosphate, binding sites, 627-628, 634

Gaucher’s disease, serum acid phosphatase, 496 Gelatin, p-lactamase and, 44 Gel electrophoresis, urease and, 7, 9 Geneb), p-lactamases and, 26,42 Genetic recombination, deoxyribonucleases and, 254, 259 Gills, adenosine monophosphate aminohydrolase in, 50 Glucagon, glucose-6-phosphatase and, 598 5’-nucleotidase and, 348 Glucocorticoids, fructose diphosphatase and, 645 glucose-6-phosphatase and, 560,568, 578, 598, 599

Gluconeogenesis, fructose diphosphatase and, 612-614, 626,634,644 Glucosamine, alkaline phosphatase and, 406 deoxyribonuclease I and, 293,296 ribonucleases and, 651 spleen acid deoxyribonuclease and, 273-274, 275

Glucose, alkaline phosphatase and, 406,408 binding, glucose-6-phosphatase and, 563, 583, 586

phosphorylation by various enzymes, 593

transferase activity and, 440, 446 transphosphorylation, acid phosphatase and, 461-462 transport, glucose-6-phosphatase and, 562-564

Fructose-6-phosphate, glucose-6phosphatase and, 567,569,590 Fungi, acid phosphatase of, 451,497 asparaginases of, 103, 104, 105 Fusarium tricinctum,asparaginase of, 116, 117, 119

G p-Galactosidase, urease and, 13 Galactose, glucose-6-phosphatase and, 570, 571

Gastric mucosa, urease in, 14-15

Glucose oxidase, glucose-6-phosphatase assay and, 566 Glucose-6-phosphatase, catalytic properties, 565-566 assay, 566-567 control, 592-595 kinetics and mechanism, 572-592 reactions catalyzed, 567-571 thermodynamic considerations, 571572

distribution, intracellular, 548-551 tissue and phylogenetic, 547-548, 600-611

SUBJECT INDEX

historical, phosphohydrolase activity, 545-546 phosphotransferase activity, 546-547 metabolic roles and regulation in G V O , 596-599 molecular properties, detergents and, 556-562 membranous nature, 562-564 phospholipids and, 554-556 solubilization and attempted purification, 553-554 reactions catalyzed, 544-545 multifunctional nature, 567568 substrate specificity, 588-571 relation to other enzymes, 552 Glucose phosphate(s), acid phosphatases and, liver, 487, 490,493 plant, 497 spleen, 494 Glucose 1-phosphate, alkaline phosphatase and, 408,430 Glucose 6-phosphate, alkaline phosphatase, 396-397 bacterial, 374 mammalian, 419, 439 Glucoseb-phosphate dehydrogenase, fructose-l,6diphosphatase assay and, 615 phosphotransferase assay and, 568 Glutamate, conversion to pyrrolidone carboxylate, enzymic, 133-139 nonenzymic, 130-132 derivatives, n-glutamate cyclotransferase and, 135 glutaminase and, 82-84 y-esters, cyclization of, 132 formation, pyrrolidone carboxylate and, 150-151 glutaminase and, 83,84,89, 92, 98 D-Glutamate cyclotransferase, assay, 126 purification and properties, 134-136 pyrrolidone carboxylate formation by, 133-136 L-Glutamate cyclotransferase, pyrrolidone carboxylate formation and, 138

869 Glutamate dehydrogenase, urease assay and, 13 Glutamate residues, ribonuclease A, 685, 788 ribonuclease TIand, 220,221,222 staphvlococcal nuclease, 167, 173, 182, 201,203 Glutamic acid y-N-methylaide, transamination product, 146-147 Glutaminase(s), 1 acyltransfer reactions, 84-85 assay of, 81 6diazo-5-oxonorleucine and, 82, 85-87, 93, 98 kinetic parameters, p H and, 88, 89, 93 98, mechanism of action, 90, 9?!9-3, 99-100 occurrence of, 81 other acylases and, 90,91 other inhibitors of, 87 purification of, 82 relationships among, 98-100 specificity, 82-84, 99 substrate kinetic parameters, 83 survey of, 93-95 temperature effects, 88,89,93 Glutaminase I, acceptors for, 93 Glutamine, asparaginases and, 107, 110, 115 cyclization, enzymic, 139-141, 143 nonenzymic, 132, 133 glutaminase and, 83,&3,89,90 staphylococcal nuclease glutamate residues and, 203 transamination of, 146 Glutamine cyclotransferase, purification and properties, 140-141 pyrrolidone carboxylate formation and, 139-141 Glutamine residues, asparaginase, 115 ribonuclease, 665,704,787,788,794 StaPhYlococcal nuclease, N-terminal, conversion to pyrrdidoae carboxylate, 129 Glutamine synthetase, pyrrolidone carboxylate and, 124,136-137

SUBJECT INDEX

L-Glutaminyl-L-asparagine, glutamine cyclotransferase and, 140, 141 Glutaminyl peptides, cyclization of, 139 Glutaminyl-transfer ribonucleic acid, conversion to pyrrolidone carboxylyl-transfer ribonucleic acid, 130, 141 y-Glutamyl cycle, function of, 151 y-Glutamyl cyclotransferase, assay of, 127 purification of, 144 pyrrolidone carboxylate formation and, 141, 142-146 reaction catalyzed, 94, 124 y-Glutamylcysteine synthetase, pyrrolidone carboxylate formation and, 137 y-Glutamylglycine, pyrrolidone carboxylate formation and, 142, 144 y-Glutamylhydroxamate, cyclization of, 132 Glutamyl methylamide, glutaminase and, 83,88,89,90 y-Glutamyl peptides, cyclization of, 132, 133 y-glutamyl cyclotransferase specificity and, 145-146 y-Glutamyl phosphate, pyrrolidone carboxylate formation and, 137 Glutamyltransferase, mushroom, 94, 95-96 totally nonhydrolytic, 94 y-Glutamyl transpeptidase, y-glutamyl cyclotransferase activity and, 145-146 kidney, 94, 96-97 pyrrolidone carboxylate formation and, 141, 142 Glutathione, glutamyl transferase and, 94, 96, 97 pyrrolidone carboxylate formation and, 125, 133,142,143, 144 reduced, fructose-l,6diphosphatase and, 623, 624 ribonuclease and, 692-693 Glyceraldehyde, phosphotransferase and, 570 Glyceraldehyde-3-phosphate dehydrogenase, activity of, 594

sedoheptulose, 1,irdiphosphate hydrolysis and, 615 n-Glycerate, acid phosphatase and, 463, 464 Glycerol, glucose-6-phosphatase and, 553, 554, 568, 570, 573 ribonuclease and, 779 transferase activity and, 440, 446 urease and, 8, 18 Glycerophosphates, acid phosphatase and, 450 erythrocytes, 477, 478 fungal, 497 Gaucher’s disease, 496 liver, 484, 487,488,492, 493 plasma, 495 prostatic, 456, 457-459, 473 spleen, 491 alkaline phosphatases and, 428,429, 430, 434-435, 438, 440 a-Glycerophosphate, gluconeogenesis and, 634-635 Glycine, alkaline phosphatase and, 427, 440 deficiency, asparaginase and, 119-120 fructose-l,6-diphosphatase and, 617 staphylococcal nuclease histidine residues and, 202 Glycine residues, fructose diphosphatase, 627 ribonuclease, 665 staphylococcal nuclease, 167 Glyciridiu maculata, urease of, 14 Glycolytic intermediates, glucose-6phosphatase and, 579 Glycoproteins, ribonuclease and, 651 Glycylglycine, glutamyl transferase and, 94, 97 Glyoxal, ribonuclease TIand, 221 Glyoxal guanylate phosphodiester bonds, ribonuclease T,and, 228 Glyoxalguanylyl-(%, 5’)-uridine, ribonuclease U, and, 236,238 Glyoxylate, glycine synthesis from, 120 Grass, pyrrolidone carboxylate in, 128 Ground squirrel, glucose-6-phosphatase in, 600 Guanidine, erythrocyte acid phosphatase and, 481

87 1

SUBJECT INDEX

spleen acid deoxyribonuclease and, 276 Guanidine hydrochloride, adenosine aminohydrolase and, 56, 74 alkaline phosphatase and, 382, 385, 389 asparaginase and, 114, 115 j3-lactamases and, 45 pyrophosphatase and, 508-510, 512, 519 ribonuclease and, 709, 710,726, 729 staphylococcal nuclease and, 184 urease and, 9, 11 Guanidinium salts, ribonuclease and, 735, 736 Guanine, adenine aminohydrolase and, 54 analogs, guanine aminohydrolase and, 76 deamination, absorbancy change, 53 ribonucleaae T,and, 216 Guanine aminohydrolase, distribution of, 50-51, 76 historical, 48 properties of, 76-77 Guanosine, adenine aminohydrolase and, 54 deamination, absorbancy change, 53 derivatives, ribonuclease and, 760 Guanosine aminohydrolase, properties of, 77-78 Guanosine 2’, 3’-cyclic phosphate, ribonuclease N, and, 232, 233 ribonuclease TI, and, 210, 213, 215, 216, 218, 221 Guanosine 3’,5’-cyclic phosphate, ribonuclease T,and, 218 Guanosine 2’-monophosphate, and, 220 ribonuclease T1, Guanosine triphosphate, 5’-adenylic acid aminohydrolase and, 65, 69, 70 venom exonuclease and, 327 3’-Guanylyl phosphodiester bond, splitting, neighboring nucleoside and, 216-217 Guanylylnucleosides, synthesis of, 222 Guanylyl-(3’, 5’)-uridine, synthesis of, 232-233 Guinea pig, tissues, glucose-6-phosphatase in, 549, 567, 6OO-606 Guinea pig serum,

asparaginase, 102 amino acids, 113 isolation, 105-106 properties, 106-107, 118-119

H Hamster, glucose-6-phosphatase in, 604 Haplotomic mechanism, spleen acid deoxyribonuclease, 279 Heart, acid phosphatase, electrophoresis of, 454, 455 adenosine aminohydrolase, 55, 56, 6364 kinetic properties, 57 3, 5’-cyclic phosphate diesterase of, 367, 368 5‘-nucleotidase of, 347-348,351 Heavy metals, 5’-adenylic acid aminohydrolase and, 70 glutaminase and, 87 5’-nucleotidase and, 341 spleen exonuclease and, 335 staphylococcal nuclease and, 187 urease and, 19 Hela cells, alkaline phosphatase of, 422, 426, 442 or-Helix, pyrophosphatase, 505 ribonuclease A, 666-667, 721-722, 736 ribonuclease T1and, 219 spleen acid deoxyribonuclease, 275 staphylococcal nuclease, 161, 163,183 Hemachatus haemachates, 5’-nucleotides of, 342 venom exonuclease, 318 Hematoporphyrin, ribonuclease and, 772 Hemophilus influenzae, deoxyribonucleic acid, spleen acid deoxyribonuclease and, 280 restriction endonuclease in, 263 Heparin, ribonuclease and, 758 Heptoseb), phosphotransferase and, 570, 571 Heptuloses, phosphotransferase and, 570 Heteroduplexes, restriction endonuclease and, 264 Hexanucleotides, staphylococcal nuclease and, 185-186

872 Hexitol diphosphate, fructose-1,6diphosphataae and, 616 Hexosamine, alkaline phosphatase and, 424 Hexose, alkaline phosphataae and, 424 phosphotransferase and, 570, 571 Hexose phosphates, alkaline phosphatase and, 428 glucose-6-phosphatase and, 568-569, 590 Histidine, alkaline phosphatase and, 442 fructose-l,6-diphosphataseand, 617, 624, 635, 644 5’-nucieotidase and, 343 pyrophosphatase and, 541 Histidine residues, alkaline phosphatase, 383, 391 replacement of, 390 deoxyribonuclease I and, 297-299 fructose diphosphataee, 633 fungal ribonucleases, 210 glucose-6-phosphatase, 574, 586, 587, 591, 592 prostatic acid phosphatase, 471 pyrophosphatase, 515, 517-518 ribonuclease A, 657, 659, 665, 667, 669, 676, 682, 711-712, 724-725, 736, 756, 759,770, 771, 779, 788,802 catalytic mechanism and, 780, 782, 783, 784, 785, 787, 790, 792-793, 794-795, 796, 797,798,799 denaturation and, 739 dimers and, 745, 746 modification of, 685-689, 690, 698, 702, 761 substrate binding and, 763-765, 769, 800, 801, 803-805 ribonuclease TIand, 220,222 spleen acid deoxyribonuclease, 276, 281 staphylococcal nuclease, 182, 195, 202 urease, 20 Histochemistry, alkaline phosphatase, 433 Homocystine, fructose-l,6diphosphatase and, 624-625, 626,644 Homoglutamine, cyclisation of, 133 Horse, glucose-6-phosphatase in, 604

SUBJECT INDEX

Human, 6‘-nucleotidase of, 345 tiasues, glucose-6-phosphatase in, 551, 567, 6o0-606 Hydration, ribonuclease, 704, 709-710 Hydrasine, glutamyl transferase and, 94 Hydrasinolysis, asparaginase, 115 6-Hydrazinopurine, adenine aminohydrolase and, 54 6-Hydrazinopurine riboside, adenosine aminohydrolase and, 62 Hydrochloric acid, alkaline phosphatase release and, 378 Hydrocortisone, alkaline phosphataae and, 422 Hydrogen bonding, ribonucleaae, 666-667, 714, 717, 718, 787-788 substrate and, 754-755 Hydrogen exchange, ribonuclease, 712-714 Hydrogen ion equilibria, ribonuclease, 711-712 Hydrogen peroxide, ribonuclease and, 682 spleen acid deoxyribonuclease and, 281 Hydrophobic groups, ribonuclease, 659 Hydroxamate, p-lactamaae assay and, 39 urease and, 14,16, 17,20 p-Hydroxyaniline, glutamyl transferase and, 95 a-Hydroxycarboxylic acids, acid phosphatase and, 462-465 p-Hydroxyglutamate, cyclisation of, 133 5-Hydroxy4-keto-~-norvaline, asparaginase and, 111 Hydroxylamine, alkaline phosphatase and, 408 dihydroxyurea hydrolysis and, 16 glutaminase and, 80, 81, 84-85, 92, 93, 94, 97-98, 99 glutamyl transferases and, 95, 97 pyrrolidone carboxylate determination and, 126-127 urease and, 17 p-Hydroxymercuribenzoate, alkaline phosphatase and, 428 pyrophosphatase and, 515, 519

873

SUBJECT INDEX

Hydroxymethyl cytosine, glucosylated, venom exonucleaae and, 316, 321

9-(l-Hydroxymethyldecanyl) adenine, adenosine aminohydrolase and, 59 Hydroxymethylene diphosphonate, yeaat pyrophosphatase and, 535 7-Hydroxy-y-methylglutamate, cyclization of, 133 5-Hydroxy-N-methylpryr yrrolidone carboxylate, formation of, 147 2-Hydroxy-5-nitrobenzyl bromide, prostatic acid phosphatase and, 471 ribonuclease Ti, and 221 2-Hydroxy-5-nitrobenzylphosphonate, alkaline phosphatme and, 393,411 S-Hydroxyquinoline-5-sulfonate, alkaline phosphatase and, 401,402 Hydroxyurea, urease and, 16, 17, 18 Hyperchromic shift, deoxyribonuclease I and, 307 spleen acid deoxyribonuclease action and, 276,277,278 Hypoglycemic agents, glucose-6phosphatase and, 681 Hypophysectomy, glucose-6-phosphatase and, 598

Inosine monophosphate, ineffectivenem of, 47-48 Inosinyl-(3’, 5’)-nucleoside, synthesis of, 233

Insulin, cyclic 3’,5’-adenosine monophosphate and, 369-370 glucose-6-phosphatase and, 568, 597, 598

glutamine residues, conversion to pyrrolidone carboxylate, 129 Intestine, acid phosphatase, electrophoresis of, 454, 455

adenosine aminohydrolase in, 50 alkaline phosphatase, 418, 419, 420, 421, 422, 453

chelating agents and, 426-427 chemical modification, 427-428 composition, 424, 425 kinetic studies, 434-438, 440,442 purification, 423 stability, 425426 substrates, 429, 430 transferase action, 431 5’-nucleotidase of, 345,351,352 glucose-6-phosphatase in, 551, 565, 567, 571, 578, 596,603

I Imidazole, 3’,5’-cyclic phosphate diesterase and, 369, 370 Imidazole groups, ribonuclease Ti and, 219, 220

Immunoglobulin, pyrrolidone carboxylate in, 128, 130 Immunology, fungal ribonucleases, 210-211 p-lactamases, 46 staphylococcal nuclease, 182, 201 urease, 13 Indigogenesis, alkaline phosphatase localization and, 433 Inhibitors, alkaline phosphatase, 394-396 Inorganic pyrophosphate, see Pyrophosphate Inosine, adenosine aminohydrolase and, 59, 61, 62

occurrence of, 48

Intrinsic viscosity, ribonuclease, 704, 710 thermal transition and, 727, 729 Invertebrates, adenosine monophosphate aminohydrolase in, 50 alkaline phosphatase in, 420 nerves, cyclic phosphate diesterase and, 365 Iodide, glucose-6-phosphatase and, 581 Iodine, fructose diphosphatase and, 637,638 ribonuclease and, 684, 689, 702 staphylococcal nuclease and, 157,163 Iodine monochloride, prostatic acid phosphatase and, 469-471 Iodoacetamide, adenosine aminohydrolase and, 58 alkaline phosphatase and, 391, 428, 442 fructose-l,6-diphosphatase and, 622, 623

874 Iodoacetamide ( c o d ribonuclease and, 686,688 Iodoacetate, 5’-adenylic acid aminohydrolase and, 70 deoxyribonuclease and, 298-299 fructose-1 ,g-diphosphatase and, 622 ribonuclease A, cysteine residues, 691 dimers and, 745 histidine residues, 686687, 688, 780, 804 lysine residues, 682, 686, 688 methionine residues, 683,686 ribonuclease TIand, 210, 211, 220 spleen acid deoxyribonuclease and, 281 yeast pyrophosphatase and, 531 5-Iododeoxyuridine-3’, 5’diphosphate, staphylococcal nuclease and, 157 6-Iodopurine, adenine aminohydrolase and, 54 Iodosobenzoate, alkaline phosphatase and, 428, 442 5-Iodouridylate, ribonuclease and, 755, 799 Ions, deoxyribonuclease I and, 30Z303 ribonuclease and, 769-772 Ionic strength, alkaline phosphatase and, 435, 437-438 ribonuclease, acid transition, 729-730 inhibition and, 758 kinetics and, 777-778, 802 staphylococcal nuclease and, 190 Isionic point, ribonuclease, 701, 710-711 Isoadenosine, adenosine aminohydrolase and, 60, 62 3-Isoadenosine monophosphate, 5’-adenylic acid aminohydrolase and, 65, 69 Isoelectric point, asparaginases, 114-115, 116 urease, 10 Isoeneymes, alkaline phosphatase, 384-387 asparaginase, 110, 115 erythrocyte acid phosphatase, 480-483 Isoleucine residues, asparsginase, 115

SUBJECT INDEX

ribonuclease, 665 NO-Isopentenyladenosine, ribonuclease Tzand, 226

J Jack beans, availability of, 3

K Ketene, alkaline phosphatase and, 427 a-Ketoglutaramate, formation of, 146 a-Ketoglutarate, reaction with methylamine, 147 a-Ketoq-methylglutaramate,formation of, 147 a-Ketosuccinamate, hydrolysis of, 101 Kidney, acid phosphatase, 450,451 electrophoresis, 454, 455 adenosine monophosphate aminohydrolase in, 50 alkaline phosphatase, 417,418, 419, 420, 421,446 chelating agents and, 426, 427 chemical modification, 427, 428 composition, 424, 425 purification, 423 fructose-l,6diphosphatase, 645 purification and properties, 629-630 regulation, 630-631 glucose-6-phosphatase in, 549, 558, 565, 567,576,596, 602 glutaminases of, 87, 95 glutamyl transferase, glutathione and. 94 y-glutamyl transpeptidase of, 94, 96-97 5’-nucleotidase of, 348 Kinetic constants, alkaline phosphatase, 436-439 ribonuclease TI, 217 ribonuclease Tz, 228, 229 ribonucleoside 2’, 3’-cyclic phosphate diesterase, 358, 360 Kinetic studies, alkaline phosphatase, 409-415 factors affecting activity, 434-436 inhibition and, 442443 kinetic constants, 436,439

875

SUBJECT INDEX

metal ions and, 440-442 phosphorylenzyme and, 439 transferase activity and, 439440 glucose-6-phosphatase, 568, 572-574 activators and inhibitors, 578-582 mechanism, 582-592 pH, 574-576 substrate concentration, 576-577 temperature, 577-578 prostatic acid phosphatase, fluoride and, 459462 a-hydroxycarboxylic acids and, 462-465

ions and, 466 pH and substrate effects, 457459 surface inactivation, 459 ribonuclease, ionic strength and, 777-778 Michaelis constants and turnover numbers, 772-777 organic solvents and, 779-780 yeast pyrophosphatase, 535-538 Klebsielln aerogenes, 8-lactamase, catalytic constants, 41 inhibition of, 44

L 8-Lactamase(s), background, 23-25 catalytic properties, assay methods, 35, 39 enzyme structural modifications, 41-42

factors affecting activity, 4 2 4 4 kinetics and substrate specificity, 3940

substrate structural modifications, 40-41

catalytic reaction, 27 conformation and function, nonspecific conformational transitions, 44-45 specific transitions : conformative response, 4 M 6 definitions and specificity, 2 b 2 6 function of, 24 immunological studies, 45 inhibitors of, 43-44

molecular properties, 31, 32 composition and sequence analysis, 31-35

purification and physical properties, 27-31

occurrence of, 26 substrates, 34 Lactate dehydrogenase, activity of, 594 fructose-l,6diphosphatase assay and, 615

Lactate dehydrogenase elevating virus, asparaginase effectiveness and, 120-121

Lactobacillus acidophilus, exonuclease of, 326 Lactobacillus casei, ribonuclease of, 245 Lactobacillus plantarum, ribonuclease of, 245 Lamprey, glucose-6-phosphatase in, 600 Laurylamine, glucose-6-phosphatase and, 557

Lecithinase, glucose-6-phosphatase and, 554, 555

Lenzites tenuis, ribonuclease of, 247 Leucine, glucose-6-phosphatase and, 679 Leucine aminopeptidase, ribonuclease T,,and, 222 staphylococcal nuclease and, 181, 182 Leucine residues, alkaline phosphatase, replacement of, 390

asparaginase, 115, 116 deoxyribonuclease I and, 297 ribonuclease, 665, 669 staphylococcal nuclease, 167, 182, 202 Leukocytes, alkaline phosphatme, 423 Lipoprotein, alkaline phosphatase and, 423

Lithium ions, 5’-adenylic acid aminohydrolase and, 65, 67-69 Liver, acid exonuclease in, 336 acid phosphatase of, 450, 451 bovine, 491493 mouse, 489-491 rat, 484-489 adenosine aminohydrolase of, 55-58, 57 asparaginase of, 101-102

876

SUBJECT INDEX

Liver (cont .) 3’, 5’-cyclic phosphate diesterase of, 367, 368 fructose-l,6diphosphatase, 645, 646 disulfide exchange and, 622-626 molecular structure, 626-629 proteolysis and, 618 purification and properties, 616-618 regulation of, 618-620 sulfhydryl groups, 621-622 glucose-6-phosphatase in, 549, 550, 551, 554, 559, 565, 567, 570, 574, 576, 578,596, 597,600

nuclei, pyrrolidone carboxylate formation and, 138-139 5’-nucleotidase of, 343-345, 349, 351, 352

phosphatases, 418, 420, 421,446 chemical modification, 427 composition, 424 purification, 423 stability, 425-426 pyrrolidone carboxylyl peptidase of, 149

spleen acid deoxyribonuclease inhibitor in, 282 Lobster, adenosine aminohydrolase in, 49 guanine aminohydrolase in, 50 Lubrol W, glucose-6-phosphatase and, 553,557 Lumbricus terrestris, 3’, 5’-cyclic phosphate diesterase in, 366

Lung, adenosine aminohydrolase of, 55, 56 alkaline phosphatase of, 420 Lupine, acid phosphatase of, 497 Lymphosarcoma, asparagine synthetase in, 118 bacterial asparaginases and, 102, 104-105, 116

guinea pig serum and, 102,106 Lysine residues, adenosine aminohydrolase, 58 alkaline phosphatase, 382383, 391 asparaginase, 116 fructose-l,6diphosphatase, 620, 631 pyrophosphatase, 514, 515-516, 518, 526

ribonuclease, 665, 711, 717, 725, 727, 770, 779, 780, 782, 783, 784, 785, 787, 788, 793, 794, 795, 803, 805 denaturation and, 739 modification of, 678482,696897, 698,801 spleen acid deoxyribonuclease, 276 staphylococcal nuclease, 167, 171, 173, 195, 197, 201

Lysolecithin, glucose-6-phosphatase and, 561, 578, 579

Lysosomes, acid deoxyribonuclease in, 271,286 acid exonuclease in, 336 acid phosphatase in, 484, 485, 489, 491, 498

5’-nucleotidase of, 343,349, 351, 352 9-(a+Lyxofuranosyl) derivatives, ribonuclease T, and, 228 9-(a-L-Lyxofuranosyl)-hypoxanthine 2’, 3’-cyclic phosphate, ribonuclease TIand, 218 CY-L-L~XO thymidine cyclic phosphate, ribonuclease and, 754 fl-n-Lyxo-uridine-2’,3’-cycbcphosphate ribonuclease A and, 752, 760

M Macrophages, acid phosphatase of, 496 Magnesium ions, acid phosphatase and, 492 erythrocyte, 478, 482 liver, 492 platelet, 495 spleen, 494 alkaline phosphatase and, 418, 4#1, 427,429,431,435,440442 3, %-cyclic phosphate diesterase and, 368 deoxyribonuclease I and, 302-303, 307, 308, 309 fructose diphosphatase and, 612, 617, 622, 623, 624, 625, 628, 632, 633, 636, 639, 641, 642, 643, 646 microbial ribonucleases and, 241,242 5’-nucleotidase and, 342, 343, 344, 345, 346, 348, 350-351, 352 pyrophosphatase and,

SUBJECT INDEX

animal, 540 bacterial, 518519,520, 521-525, 526-527 plant, 540 yeast, 530,532-534, 535-538 ribonuclease and, 770

spleen acid deoxyribonuclease and, 281,283

Maize, pyrophosphataae of, 540 Malachite green, phosphomolybdate and, 432 Maleic anhydride, fructose diphosphatase and, 627 Malonaldehyde, ribonuclease and, 689 Maltose, dietary, glucose-6-phosphatase and, 598

Mammals, adenosine aminohydrolase in, 50,57 adenosine monophosphate aminohydrolase in, 50 5’-adenylic acid aminohydrolase, activation of, 68 alkaline phosphatase, assay techniques, 432434 chemical modification, 427-428 distribution, 420-421 function, 421422 general survey, 417-420 kinetic studies, 434-443 mechanism, 443-447 physical properties, 423-427 purification procedures, 422423 reaction catalyzed, 430-432 substrate specificity, 428-430 3’, 5’-cyclic phosphate diesterase in, 366

prostatic acid phosphatase in, 455 serum, asparaginase in, 10%103, 105 tissues, y-glutamyl cyclotransferase in, 143

Mammary gland, alkaline phosphatase of, 421 5’-nucleotidase of, 348 Manganese ions, acid phosphatases and, 479 alkaline phosphatase and, 402,440 3, 5’-cyclic phosphate diesterase and, 368

deoxyribonuclease I and, 302403, 308, 309 fructose-1,6diphosphatase and, 617, 622, 623, 624, 625, 627, 632, 633, 636,639,641,642,644,646 binding sites, 628 5’-nucleotidase and, 339, 344, 345, 346, 348,350-351,352 pyrophosphatase and, bacterial, 518-519,520,539 yeast, 532,534,535,537 Mannose, deoxyribonuclease I and, 293,296,297 glucose-6-phosphatase and, 570, 571, 579 ribonucleases and, 651 spleen acid deoxyribonuclease and, 273-274, 275 Mannose-6-phosphate, glucose-6phosphatase and, 547, 563, 567, 568, 569, 575, 576, 583, 584, 586, 590, 591, 592 Marmot, glucose-6-phosphataee in, 605 Mechanism of catalysis, ribonuclease, discussion, 784-786 Mathias, Rabin et al., 780-781 Roberts et al., 784 Usher, 783-784 Wang, 782-783 Wetzel, 781-782 Membrane(s), glucose-6-phosphataee and, 562-564 Mercaptoethanol, 5’-adenylic acid aminohydrolase and, 65 alkaline phosphatase and, 385 deoxyribonuclease I and, 297 pyrophosphatase reconstitution and, 510-511 ribonuclease and, 691 spleen acid deoxyribonuclease and, 276 urease and, 3,9 6-Mercaptopurine, resistance, 5’-nucleotidase and, 349 p-Mercuribenzoate, adenosine aminohydrolase and, 68, 74 5’-adenylic acid aminohydrolase and, 65

SUBJECT INDEX

p-Mercuribenzoate (cont.) glutaminase and, 87, 98 fructose-l,6diphosphatase and, 622, 623, 633, 637 prostatic acid phosphatase and, 469 ribonuclease and, 691 Mercuric ions, @-lactamases and, 44 Mercury, alkaline phosphatase and, 401,402 ribonuclease and, 692 Metalloenzyme, yeast pyrophosphatase, 532-534 Methanol, glutaminase and, 85, 94 polyadenylate digestion and, 226 ribonuclease and, 747 esterification, 675-676 Methicillin, @-lactamasesand, 34,38,42 Methionine residues, alkaline phosphatase, replacement of, 3W391 fructose diphosphatase, 633 fungal ribonucleases, 210 ribonuclease, 658, 659, 665-666, 669, 702, 735 modification of, 682-683,690,691 spleen acid deoxyribonuclease and, 281 staphylococcal nuclease, 180, 202 urease, 12 L-Methionine sulfoxime, pyrrolidone carboxylate utilization and, 150-151 9-(4-Methoxyphenyl) guanine, guanine aminohydrolase and, 77 6-Methowpurine ribonucleoside, adenosine aminohydrolase and, 59, 60,62 Methyl acetimidate, ribonuclease and, 679 N1-Methyladenosine, adenosine aminohydrolase and, 60, 62 Ne-Methyladenosine, ribonuclease T1 and, 230 1-Methyladenylate, ribonuclease T, and, 227 I-MethyIadenylyl-(3’,5‘)-uridine, ribonuclease U, and, 236 Methylamine, reaction with a-ketoglutarate, 147 Methylene-bis-phosphonate, pyrophosphatase and, 525, 535

y-Methyleneglutamine, cyclization of, 133 3-0-Methylfluorescein phosphate, alkaline phosphatase assay and, 433 Methyl n-fructofuranoside-l,6-diphosphate, fructose-l,6diphosphatase and, 616 Methylglutamate, glutaminase and, 83, 88, 89, 90 a-Methylglutamine, deamidation of, 133 y-Methylglutamine, deamidation of, 133 transamination of, 147 Methyl green, phosphomolybdate and, 432 Methyl groups, endonucleases and, 262, 263, 265 7-Methylguanine, ribonuclease T, and, 227 1-Methylguanylate, ribonuclease T, and, 228 2’-0-Methyl guanylate, ribonuclease T, and, 228 7-Methylguanylyl residues, ribonuclease U, and, 236 2-Methylhistidine, alkaline phosphatase and, 390 0-Methylhydroxylamine, glutaminase and, 81 Methyl iodide, ribonuclease and, 682 0-Methylisourea, ribonuclease and, 678, 679 Methyl methane sulfonate, endonuclease I1 and, 264-265 2-Methyl-2,4-pentanediol, ribonuclease and, 657 staphylococcal nuclease crystallization and, 156 N-Methylpseudouridylate, ribonuclease and, 799 2’-0-Methyl ribose, derivatives, venom exonuclease and, 320 2-Methylthio-Ne- (A*-isopentenyl) adenosine, ribonuclease T,and, 226-227 4-Methylumbelliferyl phosphate, alkaline phosphatase assay and, 433, 436, 438

879

SUBJECT INDEX

Methyl urea, adenosine aminohydrolase and, 58-59 Micrococcal nuclease, sequence determination and, 326 substrates of, 334 Micrococcus luteus, adenosine triphosphate-dependent deoxyribonuclease of, 261-262 ultraviolet repair enzymes of, 269-270 Microorganisms, acid phosphatase of, 451,497498 adenosine monophosphate aminohydrolase of, 50 glucose-6-phosphatase in, 609 3’-nucleotidase in, 354 urease in, 13 Microsomes, glucose-6-phosphatase and, 546-547, 548-551 5‘-nucleotidase and, 344 Microsporum audouini, adenine nucleotide aminohydrolase, 74, 75 Milk, alkaline phosphatase of, 423, 424, 426, 431, 436, 437, 439 Mitochondria, pyrophosphatases of, 540 Molybdate, acid phosphatase and, 494,497 glucose-6-phosphatase and, 568, 580, 581 Monascus pilotus, ribonuclease of, 247 Monkeys, serum, asparaginase in, 103 tissues, glucose-6-phosphatase in, 600603 Mononucleotides, deoxyribonuclease I and, 310 Monophosphatases, 328 removal from venom exonuclease, 317-318 Mouse, liver acid phosphatase, 489-491 pyrophosphatase of, 536, 540 tissues, glucose-6-phosphatase in, 600604 Mucor genevemnsis, ribonuclease of, 247 Mud puppy, glucose-6-phosphatase in, 602 Mungbean, nuclease I, 315

3’-nucleotidase of, 353 Muscle, acid exonuclease in, 336 acid phosphatase, electrophoresis, 454, 455 adenosine monophosphate aminohydrolase in, 50 5’-adenylic acid aminohydrolase, 64-65 activation, 67-69 dystrophy and, 71 inhibition, 70 kinetic constants, 67 physiological function, 72 specificity, 66 fructose-1,6-diphosphatase, 645 evidence for, 632 physiological role, 634-635 purification and properties, 632-633 structure and relation t o other diphosphatases, 633-634 5’-nucleotidase of, 348, 351 Mushroom, glutamyltransferase of, 94, 95-96 Mutations, p-lactamase, 26, 42 Mycobacterium avium, ribonuclease of, 245

Mycobacterium tuberculosis, adenosine aminohydrolase of, 55 asparaginase of, 117 Myelin, cyclic phosphate diesterase in, 364-365

N Naja naja atra, 5’-nucleotidase of, 342, 351, 352 venom exonuclease, 318 Naphthyl phosphate(s), acid phosphatase and, erythrocyte, 477 liver, 484 a-Naphthyl phosphate, acid phosphataae, prostatic, 456,457 p-Naphthyl phosphate, acid phosphatase, prostatic, 456 alkaline phosphatase and, 393, 398, 433, 434, 435 Neoplasias, 5’-adenylic acid aminohydrolase and, 71

SUBJECT INDEX

Nervous tissue, 5’-nucleotidase of, 346-347, 351, 352 ribonucleoside 2’,3’-cyclic phosphate diesterase, intracellulal localization, 364 physiological role, 365 properties and substrate specificity, 364 Neurospora crassa, acid phosphatase of, 497 ribonuclease, 230-234 main properties, 208,247 Newt, glucose-6-phosphatase in, 602 Nickel ions, acid phosphatases and, 479 alkaline phosphatase and, 402 5’-nucleotidase and, 341,342,352 yeast pyrophosphatase and, 535 Nicotinamide adenine dinucleotide phosphate, alkaline phosphatase and, 394,432 fructose-l,6diphosphatase and, 615, 623 N i t h t e ions, ribonuclease and, 778,804 p-Nitro blue tetrazolium, urease detection and, 5 Nitrocellulose membrane filters, deoxyribonucleic acid retained by, 266 p-Nitrophenacyl bromide, ribonuclease and, 698 p-Nitrophenyl esters, spleen exonuclease and, 333, 334,335 p-Nitrophenyl phosphate, acid phosphatases and, 452, 453 erythrocyte, 481, 482 liver, 487, 490, 491493 prostatic, 456, 457-459, 462463, 465, 473 spleen, 494, 495 serum, 495 alkaline phosphatase and, 377, 386, 390, 392, 396, 405, 406, 408, 412, 413, 433, 435, 436, 437, 438, 439, 452, 453 ribonudeotide 2’,3‘-cyclic phosphate diesterase and, 357, 358 bis(p-Nitrophenyl) phosphate, 5’-nucleotidase and, 338-339 ribonucleotide 2’,3’-cyclic phosphate diesterase and, 357, 358,360,361

spleen acid deoxyribonuclease and, 273, 276, 283 spleen exonuclease and, 333, 334,335 0-p-Nitrophenyl phosphorothioate, alkaline phosphatase and, 395,396 4-Nitrophenylsulfeny1 chloride, ribonuclease and, 691 0-p-Nitrophenyl thiophosphate, acid phosphatase and, 452, 453 p-Nitrophenyl thymidine-3‘-phosphatee, spleen acid exonuclease and, 335,336 Nitrosoguanidine, ultraviolet repair mutants and, 270 Nitrous acid, alkaline phosphatase and, 427 asparaginase and, 116 Norleucine, alkaline phosphatase and, 390 staphylococcal nuclease and, 202 Nuclear magnetic resonance, ribonuclease, 723-725,763-765, 790, 794795, 801,802,804 Nuclei, glucose-6-phosphatase in, 549, 550, 551 5‘-nucleotidase in, 349, 351 Nucleoside(s), 5‘-nucleotidase and, 344, 346 ribonuclease and, 759, 760 Nucleoside 3’,5’-cyclic phosphate diesterase, 365-366 distribution in nature, 366 inhibitors and activators, 368-370 intracellular localization, 367-368 metal ions, pH and substrate d n i t y , 368 physiological function, 370-371 possibility of other diesterases, 370 substrate specificity, 366-367 Nucleoside diphosphate sugars, 5’nucleotidaae and, 339,340 Nucleoside phosphoacyl hydrolase, substrates of, 333, 334 Nucleoside polyphosphatase, substrates of, 333, 334, 335 Nucleoside triphosphates, 3’,5’-cyclic phosphate diesterase and, 368,369 5’-nucleotidase and, 346, 347 3’-Nucleotidase, microorganisms, 354

SUBJECT INDEX

mung bean, 353 ribonucleotide 2’,3’-cyclic phosphate diesterase and, 360-361 rye grass, 353 wheat seedlings, 353-354 5‘-Nucleotidase, bacterial, 338-340,349, 350 bull seminal plasma, 342-343,351 cardiac tissue, 347-348, 351 comparison of, 349-352 distribution of, 337 Ehrlich ascites tumor cells, 348-349, 351 intestine, 345, 351, 352 liver, 343-345,351, 352 nervous tissue, 346347, 351,352 other vertebrate tissues, 348 physiological function, 352 pituitary gland, 346, 351,352 potatoes, 349 protein inhibitor of, 340 snake venom, 314, 317-318, 320, 328, 342, 350351, 352 terminal identification and, 327 yeast, 341342, 350 Nucleotides, acid phosphatases and, Gaucher’s disease, 496 liver, 487, 490, 493 plant, 497 prostatic, 476 spleen, 494 alkaline phosphatase and, 393, 430 binding by ribonuclease, 759-769 glucose-6-phosphatase and, 547, 561, 567, 569, 580, 582, 583, 584, 586, 590, 591-592, 594, 595 pyrophosphatase and, 521 spleen acid deoxyribonuclease and, 284-285 synthetic, staphylococcal nuclease and, 188 Nucleotide pyrophosphatase, 5’aucleotidase and, 341

0 Oligoguanylate, synthesis of, 222 Oligomers, urease, 6 Oligonucleotides,

formation, spleen acid deoxyribonuclease and, 276, 277,278 glucosylated, spleen exonuclease and, 333 prostatic acid phosphatase and, 476 staphylococcal nuclease binding constants, 192 synthesis, 240 ribonuclease A and, 747 ribonuclease N1 and, 232-234 ribonuclease U, and, 238-239 Ophthalmic acid, pyrrolidone carboxylate formation and, 142 Optical rotatory dispersion, pyrophosphatase, 505 subunits, 510 ribonuclease, 719-723 detergents and, 736 thermal transition and, 727, 728 Organic solvents, ribonuclease and, 732, 733-735, 779-780 Osmotic shock, 5’-nucleotidase and, 338, 340 alkaline phosphatase and, 374, 377, 378 ribonucleotide 2’,3’-cyclic phosphate diesterase and, 361-362 Ossification, alkaline phosphatase and, 421 Ovalbumin, prostatic acid phosphatase and, 476 Oxacillin, j3-lactamases and, 34, 38 Oxalate, acid phosphatases and, 477,480, 482, 488 glucose-6-phosphatase and, 568, 580 Oxygen, exchange, yeast pyrophosphatase and, 538 Oxytetracycline, urease and, 17

P Palmityl coenzyme A, glucose-6phosphatase and, 559,561, 563, 578, 579, 595 Pancreas, acid phosphatase, electrophoresis of, 454, 455 glucose-6-phosphatase in, 604 5’-nucleotidase in, 348

SUBJECT INDEX

Pantethine, fructose-1,6-diphosphatase and, 624 Papain, comparison to glutaminase, 90,91 fructose-1,6-diphosphatase and, 618, 619

glucose-6-phosphatase and, 553 protein hydrolysis, ammonia formation and, 140 Papaya, L-glutamine cyclotransferase of, 139-141

Paramecium aurelia, ribonucleases of, 248

Partial specific volume, ribonuclease, 704, 705-707

Pea, seeds, adenosine monophosphate aminohydrolase in, 50 Penicillin (s) j3-lactamases and, 25 ribonuclease and, 772 side chain, j3-lactamase catalytic constants and, 44, 45-46 Penicillinase, see p-Lactamase Penicilloic acid, p-lactamase and, 25, 27 Pentose(s), phosphotransferase and, 570, 571

Pepsin, comparison to glutaminase, 90, 91 ribonuclease A and, 670, 673,685 ribonuclease T,and, 222 Pep tides, p-lactamase induction and, 26 pyrrolidone carboxylate-containing, 127, 128, 130

Peptide bonds, ultraviolet absorption of, 127

Performic acid, adenosine aminohydrolase and, 56 ribonuclease and, 691 Periodate. alkaline phosphatase and, 382, 387 ribonuclease and, 698 Periplasmic space, alkaline phosphatase and, 374-375 asparaginase and, 109 cyclic diesterase in, 362 Perphthalate, ribonucleic acid and, 227 PH, acid phosphatase, 454

erythrocyte, 482 liver, 492 prostatic, 457458, 465, 473 spleen, 495 alkaline phosphatase and, 380, 382, 425-426, 434-435, 436438, 441, 443, 444445 bacterial asparaginase and, 109, 113114, 115

3’,5’-cyclic phosphate diesterase and, 368

fructose diphosphatase and, 612, 613, 614, 624, 633, 642,

617-619, 620, 621, 622, 623, 625, 627-628, 629, 630, 632, 635-637, 638, 639, 640, 641, 643, 644, 645, 646 glucose-6-phosphatase and, 556, 557558, 559, 560, 574-576, 578, 581582, 590, 591492, 594, 595, 599 glutaminase kinetics and, 88, 89, 93 p-lactamases and, 42 microbial ribonucleases and, 208, 216, 244-248 5’-nucleotidases and, 342443, 350351

phosphoryl alkaline phosphatase and, 397, 405

pyrophosphatase, bacterial, 518, 519, 520 yeast, 535 pyrrolidone carboxylate formation and, 131 ribonuclease A, 790 conformation and, 729-731, 742 inhibition and, 758, 759, 761 kinetics and, 772-777, 779 thermal transition and, 727-728 variation of rates and equilibria with, 801-806 ribonuclease N1and, 231-232, 233 ribonuc1ease-S and, 672, 738 ribonuclease T, and, 224 ribonucleotide 2’,3’-cyclic phosphate diesterase and, 359 spleen, acid deoxyribonuclease, 280-281 exonuclease, 334-335 staphylococcal nuclease and, 18S184, 186, 190-191

urease activity and, 9, 19-20

SUBJECT INDEX

venom exonuclease specificity and, 323-324 o-Phenanthroline, 5’-adenylic acid aminohydrolase and, 70 alkaline phosphatase and, 401,402,427 glutaminase and, 87 Phenol, ribonuclease and, 734 Phenolic groups, ribonuclease T%and, 219 Phenolphthalein phosphate(s), alkaline phosphatase assay and, 433 erythrocyte acid phosphatase and, 477, 480 prostatic acid phosphatase and, 456, 472 Phenoxymethylpenicillin, p-lactamases and, 34 Phenylacetylpenicillin, p-lactamase catalytic constants and, 41 Phenylalanine, alkaline phosphatase and, 419, 429, 430,441,442,445 staphylococcal nuclease tyrosine or tryptophan residues and, 202,204 n-Phenylalanine, urease and, 17 a-Phenyl-a-alanine, alkaline phosphatase and, 443 Phenylalanine residues, alkaline phosphatase, 383 replacement of, 389-390 asparaginase, 115 ribonuclease, 658, 659, 666, 672, 673, 704-705, 787, 788 acid transition and, 738 substrate binding and, 763, 797 urea transition and, 732 Phenylglyclypenicillin, p-lactamase catalytic constants and, 41 Phenylglyoxal, ribonuclease A and, 689-690 ribonuclease TIand, 221 Phenylhydrazine, glutamyltransferase and, 95 Phenylisocyanate, alkaline phosphatase and, 427 Phenylmercuriacetate, adenosine aminohydrolase and, 58 Phenyl phosphate, alkaline phosphatase and, 418, 422,

428,429, 430, 433, 434435,436 erythrocyte acid phosphatase and, 477, 478, 481 Gaucher’s disease, 496 prostatic acid phosphatase and, 456, 457458, 461,473,474-475 Phenylurea, urease and, 17 Phlorisin, glucos+6-phosphatase and, 545, 561,563, 568,580,595 Phosphatase(s), see also Acid, Alkaline glucose-6-phosphatase assay and, 567 snake venom, 314, 317-318 Phosphatase inactivating system, glucose-6-phosphatase and, 581 Phosphate, acid phosphatase and, 487, 495, 497 alkaline phosphatase and, 394, 395, 396-397, 398-400, 401, 402, 405, 410, 418, 420,422, 442, 447 exchange, pyrophosphatase and, 519520 glucose-6-phosphatase and, 561, 580, 582, 587, 590, 591, 594, 595, 596 glutamine cyclization and, 132 pyrophosphatase and, 525-526 ribonuclease A crystals and, 657 ribonuclease TIand, 222 spleen acid deoxyribonuclease and, 281, 283 transport of, 377 Phosphate group, ribonuclease A and, 758, 781-782, 791-792, 796 Phosphocellulose, 5’-admylic acid aminohydrolase and, 64 alkaline phosphatase and, 394 p-lactamase I and, 30 staphylococcal nuclease and, 179 Phosphocholine, alkaline phosphatase and, 394 Phosphocreatine, alkaline phosphatase and, 429,430,442 Phosphodiesterase, see also Venom exonuclease activity of spleen acid deoxyribonuclease, 283 pancreatic, substrates of, 334 prostatic acid phosphatase and, 467 Phosphoenol pyruvate, alkaline phosphatase and, 393, 394, 398

SUBJECT INDEX

Phosphoenol pyruvate (cont.) fructose-l,6-diphosphataseassay and, 615 gluconeogenesis and, 644 glucose-6-phosphatase and, 547,567, 579 pyrophosphatase and, 540 Phosphoenolpyruvate carboxykinase, activity of, 594 Phosphofructokinase, activation of, 644 inhibition of, 613 fructose-l,6diphosphatase assay and, 615 Phosphoglycerate kinase, activity of, 594 Phosphohexose isomerase, fructose-l,6diphosphatase assay and, 615 Phospholipase C , glucose-6-phosphatase and, 555-556 Phospholipids, glucose-6-phosphatase and, 551,554556, 579 5’-nucleotidase and, 345 Phosphomandelate, alkaline phosphatase and, 394, 446 Phosphomolybdate, alkaline phosphatme assay and, 432 Phosphonates, alkaline phosphatase and, 395, 430, 442 Phosphoramidate, glucose-6-phosphatase and, 547,567,568,570,573,586,590, 592 Phosphorothioates, acid phosphatase and, 452453 alkaline phosphatase and, 395 ribonuclease and, 691,692,784,791 Phosphorylenayme, alkaline phosphataae, 439,443-444,445 glucose-6-phosphatase and, 583,685, 586, 587, 590-591 yeast pyrophosphatase and, 538-539 Phosphoserine, alkaline phosphatase, bacterial, 394 mammalian, 419 Phosphothreonine, alkaline phosphatase and, 394 Phosphotransferase, glucose-6-phosphataee,546-547, 549551, 555, 557, 559, 561, 563, 565,

566, 567, 568, 574-575, 583, 595, 596, 598 acceptors, 570-571,573 donors, 569-570, 573 Pho tooxidation, alkaline phosphatase, 391 ribonuclease A, histidine residues, 685-686,702 methionine residues, 683,685-686 ribonuclease TIand, 220 Photosynthesis, pyrophosphate and, 539540

pH stat, deoxyribonuclease I and, 290, 303 glutaminase assay and, 81 Phthalimidomethylesters, ribonuclease, 677 Physarum polycephalum, ribonucleases of, 241, 247 pig, 5’-nucleotidase of, 344 pancreatic ribonuclease, 655 tissues, glucose-6-phosphatase in, 600, 605 Pigeon, glucose-6-phosphatase in, 600 Pigeon crop gland, deoxyribonuclease inhibitor of, 299, 301 Pituitary, 5’-nucleotidase from, 346,351, 352 Placenta, acid phosphatase of, 454 alkaline phosphatase, 420,421,422 chelating agents and, 427 composition, 424-425 kinetic studies, 437,441,442 purification, 423 stability, 426 substrates, 429-430 Plants, acid phosphatases of, 465, 497-498 asparaginase in, 105 fructose diphosphatsses, 646 physiological role, 642843 purification and properties, 640-642 regulation, f343 glucose-6-phosphatase in, 609 pyrophosphatase of, 540 Plasma, acid phosphatase of, 450,451,457,480, 495

SUBJECT INDEX

alkaline phosphatase in, 418,422,426 Plasma membranes, 5’-nucleotidase and, 344-345 Platelets, acid phosphatase of, 457, 495 ,%Pleated sheet, pyrophosphatase, 505 ribonuclease A, 667, 721, 722, 736 ribonuclease TIand, 219 spleen acid deoxyribonuclease, 275 staphylococcal nuclease, 161 Pleaspora, ribonuclease of, 247 Polyacrylamide gel, urease and, 12 Poly (adenosine diphosphate), venom exonuclease and, 317 Polyadenylate, alkaline phosphatase and, 394 ribonuclease A and, 755-756, 790, 797798 ribonuclease TIand, 217 ribonuclease Tzand, 226 Polyanions, ribonuclease and, 75S-759 Polyaspartate, ribonuclease and, 759 Polycytidylate, ribonuclease TIand, 217 ribonuclease T, and, 226 Polydeoxyinosinate polydeoxycytidylate, deoxyribonuclease I and, 302 Polyethylene sulfonate, ribonuclease and, 759 Polyformycin, ribonuclease and, 757, 790, 797, 798, 799 Polyhedrosis virus, ribonucleic acid, ribonuclease TI and, 216 Polyinosinate, ribonuclease and, 755-756 Poly-3-isoadenylate, ribonuclease and, 757 Poly-7-methylguanylate, ribonuclease TI and, 217-218 Polynucleotide kinase, deoxyribonuclease I and, 290, 304,305 terminal identification and, 327 Polyornithine, ribonuclease and, 758 Polyphosphate(s), organic, yeast pyrophosphatase and, 534 yeast pyrophosphatase and, 534 Polyribonucleotides, spleen acid deoxyribonuclease and, 282-283

spleen acid exonuclease and, 332-333 venom exonuclease and, 319-320 Polyribose phosphate, ribonuclease and, 757 Polysphondelium pallidurn, fructose diphosphatase of, 640 Polyuridylate, derivatives, ribonuclease A and, 755 ribonuclease A and, 756 ribonuclease TIand, 217 ribonuclease Tzand, 225, 226 Porous glass, alkaline phosphatase binding to, 391-392 Porphyra crispata, adenine nucleotide aminohydrolase, 74, 75 Potassium ions, 5’-adenylic acid aminohydrolase and, 65, 66-69, 70 microbial ribonucleases and, 241, 242 Potato, acid phosphatase of, 452,472 5’-nucleotidase of, 349 Prednisolone, alkaline phosphatase and, 422 Presssure, ribonuclease and, 744 Proline residues, alkaline phosphatase, 379 deoxyribonuclease I and, 297 ribonuclease, 658, 659, 666, 669 staphylococcal nuclease, 163, 181-182 Pronase, deoxyribonuclease I and, 293 Propanediol, urease and, 8 Propanediol phosphate, prostatic acid phosphatase and, 473 n-Propyl alcohol, fructose diphosphatase and, 633 Prostate gland, acid phosphatase, 450, 451,452, 455457 assay, 457 electrophoresis, 468-469 functional groups, 469-472 kinetics, 457-466 physical properties, 476 preparation, 466-468 transphosphorylation, 472-473 use as a reagent, 473-476 Protamine, ribonuclease and, 758 Protein (s), catabolism, gluconeogenesis and, 626

SUBJECT INDEX

Protein(s) (cont.) pyrrolidone carboxylate in, 128, 129, 130 spleen acid deoxyribonuclease and, 281 synthesis, asparagine and, 119 Proteinases, deoxyribonuclease I and, 297 fructose-1,6-diphosphatase and, 618 pyrrolidone carboxylate formation and, 129 ribonuclease TIand, 222 Proteus mirabilis, ribonuclease of, 245 ribonucleotide 2’,3’-cyclic phosphate diesterase, 356, 357 kinetics and mechanism, 360-361 physical and chemical properties, 359 substrate specificity, 357 urease of, 14 Proteus rettgeri, urease of, 14 Proteus vulgaris, glutamyl transferase of, 94 5’-nucleotidase of, 340 ribonucleoside 2’,3’-cyclic phosphate diesterase, 356 kinetic constants, 360 substrate specificity, 357 urease of, 14 Protons, exchangeable, stapyhlococcal nuclease and, 155, 184 Proton transfer, ribonuclease and, 795796,801 Pseudomonas, glutaminase of, 94 Pseudomoms convexa, guanosine aminohydrolase of, 77-78 Pseudomonas cruciviue, L-glutamate cyclotransferase of, 138 Pseudomoms fluoreseem, pyrrolidone carboxylyl peptidase of, 147-149 Pseudomonas pyocyanea, 8-lactamase, dissociation constants, 38 substrate modifications and, 40 Pseudomonas saccharophila, fructose diphosphatase of, 639 Pseudouridine, ribonuclease and, 755,797 venom exonuclease and, 321 Pseudouridylate, ribonuclease T, and, 227

Purines, biosynthesis, asparaginase and, 120 Purine aminohydrolases, assay of, 51 Putrescine, ribonuclease and, 772 Pyridine nucleotides, 5’-nucleotidase and, 341, 349 Pyridoxal phosphate, alkaline phosphatase and, 394 fructose-1,6-diphosphatase and, 620, 630, 631, 645 Pyrimidine, ribonuclease and, 796-797 Pyrimidine dimers, 5’+3’ exonuclease and, 258 ultraviolet repair enzymes and, 269 Pyrophosphatase, bacterial, 500 chemical composition, 512-514 chemical modification, 514-518 effects of ions and inhibitors, 518519 electron microscopy, 506-508 interaction with inhibitors, 525-526 homogeneity, 502-503 nature of substrate binding, 522-525 pH effects, 518 physical properties, 504-506 purification, 501402 reconstitution, 510-512 reversal of reaction, 519-520 size, 504 substrate specificity and stoichiometry, 520-522 subunits in guanidine hydrochloride, 508-510 distribution of, 500, 539 glucose-6-phosphatase and, 558, 560, 565, 567-568, 569,570,573,578 yeast, assay, 534 divalent cation binding, 531-532 kinetics, 535-538 mechanism, 5 W 3 9 metalloenzyme, 532-534 physicochemical parameters, 530-531 purification, 530 specificity, 534-535 Pyrophosphate, alkaline phosphatase and, 408, 413, 428, 429, 431, 437, 443, 447 3’,5’-cyclic phosphate diesterase and,

368

SUBJECT INDEX

glucose-6-phosphatase and, 546-547, 567, 572, 574-575, 576, 582, 585, 590, 591-592, 594, 595, 596, 597 magnesium salts, pyrophosphatase and, 523-525 ossification and, 421 Pyrophosphoserine, alkaline phosphatase and, 394 2-Pyrrolidone, pyrrolidone carboxylyl peptidase and, 148, 149 Pyruvate, gluconeogenesis and, 634 Pyruvate carboxylase, activity of, 594 Pyrrolidone carboxylate, derivatives, enzymic formation of, 146-147 detection and determination, 125-127 formation of, enzymatic, 124-125 nonenzymatic, 130-133 D-glutamate cyclotransferase and, 133136 L-glutamate cyclotransferase and, 138 glutamine synthetase and, 124, 136-137 y-glutamylcysteine synthetase and, 137 y-glutamyl transpeptidase and yglutamyl cyclotransferase and, 141 historical, 124-125 metabolism of, 149-151 natural occurrence, 127-133 rat liver nuclear preparations and, 138-139 Pyrrolidone carboxylyl peptidase, assay of, 148 isolation and properties, 147-149

R Rabbit, tissues, glucose-6-phosphatase in, 549, 551,567, 571,578,600406 Radius of gyration, ribonuclease, 704, 707-708 Raia clavata, 5’-adenylic acid aminohydrolase of, 64,65 Random coil, ribonuclease TIand, 219 Rat, guanine aminohydrolase in, 51 5’-nucleotidases of, 343, 344, 345, 361, 352 liver acid phosphatase, cellular location, 484-485

isolation and purification, 485-489 pancreatic ribonuclease, 655, 669 pyrophosphatase of, 536,510 tissues, glucose-6-phosphatase in, 549, 550, 558, 559, 567, 570, 574, 578, 600-806 Rate constants, ribonuclease-nucleotide interaction, 766-768 Refractive index increment, ribonuclease, 704, 707 Reptiles, adenosine aminohydrolase of, 57 Rhizopua, ribonuclease of, 247 Rhodospirillum rubrum, pyrophosphatase of, 539 Rhodotorula glutinis, ribonucleases of, 246 N-9-(B-n-Ribofuranosyl) purin-6ylcarbamoyl threonine, ribonuclease Tzand, 227,230 ribonuclease Uzand, 236 Ribonuclease(s), Bacillus subtilis, extracellular, 239-240 intracellular, 240 bovine pancreatic, see Ribonuclease

A, B 2’,3’-cyclic phosphates and, 365 definition and claasification, 205-206 deoxyribonuclease I and, 292 Escherichia coli, 241-243,244 fragments, complementation of, 196 fungal, general survey, 208-211 microbial, list of,243-249 3’-nucleotidase and, 353 pancreatic, minor components, 650-653 ratio to cyclic phosphate diesteraae, 363 snake venom, 314 Ribonuclease A, acyl derivatives, refolding of, 695 aggregation of, 744-746 catalytic properties, assays, 747-760, 751 discussion of mechanism and specificity, 784-806 mechanism, 780-784 reaction catalyzed. 746-747 specificity, 750-758 stable complexes, 7W772 steady state kinetic data, 772-780

SUBJECT INDEX

Ribonuclease A (cont.) chemical synthesis, 697-705 denatured conformations, 737 derivatives, oligonucleotide synthesis by, 747 discussion of mechanism and specificity, 784 intermediate stabilization and, 794795 lysine 41 and, 801 opposite vs. adjacent attack, 791-794 proton transfer and rate-limiting step, 795-796 structure and, 785-788 substrate specificity and, 7!%-801 transphosphorylation or hydrolysis and, 788-791 variation of rates and equilibria with pH, 801-806 function of, 848649 inhibitors, macromolecular, 758-759 miscellaneous, 772 small molecule effectors, 759-772 isolation and chromatography, 649653 modification of covalent structure, chemical synthesis and S-peptide summary, 697-705 enzymic cleavage, 669-674 functional groups, 674-697 molecular properties, aggregation, 744-746 chain conformation and solventinduced changes, 725-744 physical parameters, 705-725 polypeptidyl, refolding of, 695 reoxidation and refolding of, 693-696 specificity of, 206 structure, mechanism and specificity and, 785-788 structure, 209 amino acid sequence, 653-654 three-dimensional, 654-669 transitions, 738 carboxymethyl, 739 dinitrophenyl, 739 polyvalyl, 740 Ribonuclease B, ratio to major component, 650

Ribonuclease NI, 230-231 amino acid composition, 209 applications, 232-234 inhibitors and activators of, 210,211 preparation of, 231 properties, 208, 231-232 specificity, 232 Ribonuclease PP,, properties of, 241 Ribonuclease-s, chemical synthesis of, 698899 formation of, 670-671 reoxidation and refolding of, 695-696 transitions in, 738-739 trypsin and, 674 X-ray diffraction of, 65fj-657,658 Ribonuclease TI, amino acids, composition, 209 sequence, 219 applications, 222-223 inhibitors and activators of, 210,211 main properties of, 208 preparation of, 212-213 properties of, 213 specificity and mode of action, ZOS, 208,214-218 structure and function, 218-222 Ribonuclease T,, amino acid composition, 209 applications, 229-230 inhibitors and activators of, 210,211 preparation of, 223-224 properties of, 208,224-225 specificity and mode of action, 22Ei229 Ribonuclease U1, amino acid composition, 209 inhibitors and activators of, 210,211 main properties of, 208 Ribonuclease U2, applications, 237-239 inhibitors and activators of, 210,211 preparation, 234-235 properties of, 208,235 specificity of, 206, 208,235-237 Ribonucleic acid, deoxyribonuclease inhibition by, 25% 260, 261 double-stranded, microbial ribonuclease and, 216,242

SUBJECT INDEX

sequence analysis, ribonuclease TIand, 222 ribonuclease T,and, 229-230 ribonuclease U2and, 237-238 staphylococcal nuclease, 1%-187 modified, 173 synthesis, inorganic pyrophosphatase and, 501 venom exonuclease and, 319-320 yeast, ribonuclease assay and, 749 Ribonucleoside 2’, 3’-cyclic phosphate diesterase, microorganismal with 3’-nucleotidase activity, 356-357 cellular localization, 361-362 kinetics and mechanism, 358-361 metals and, 362-363 physical and chemical properties, 358, 359 physiological function, 363 substrate specificity, 357-358 vertebrate nerve, 363-364 intracellular localization, 36&365 physiological role, 365 properties and substrate specificity, 364 Ribonucleoside 3’-phosphates, ribonucleoside 2’, 3’-cyclic phosphate diesterase and, 357-358 Ribooligonucleotides, sequence determination, venom exonuclease and, 324-326 Ribosomal ribonucleic acid, degradation of, 363 venom exonuclease and, 319,320 Ribosomes, ribonuclease of, 242 Ribulose diphosphate, fructose diphosphatase and, 641, 642 Rubidium ions, 5’-adenylic acid aminohydrolase and, 67-69 Rye grass, 3’-nucleotidase of, 353

S Saccharomyces cerevisiae, adenine aminohydrolase in, 49, 53 fructose diphosphatase of, 640 ribonuclease of, 246 Saccharomyces mellis, acid phosphatase of, 497

Saccharomyces oviformis, 5‘-nucleotidase of, 341-342, 349, 350 Salmonella heidelberg, 5’-nucleotidase of, 340 ribonucleoside 2’,3’-cyclic phosphate diesterase, 356 cellular localization of, 362 Salmonella typhimurium, heterogenote, alkaline phosphatase in, 376 p-lactamase, 31 5’-nucleotidase of, 340 ribonuclease of, 245 Sanger’s reagent, asparaginase and, 115 Sarcina ureae, urease of, 14 Sea urchin, eggs, adenosine aminohydrolase in, 49 glucose-6-phosphatase in, 606 Sedimentation coefficients, asparaginase, 113 pyrophosphatase, 504 subunits, 509 ribonuclease, 704, 709 thermal transition and, 727, 729 Sedoheptulose-1,7-diphosphatase, 646 relation to fructose diphosphatase, 638 Sedoheptulose 1,7-diphosphate, fructose 1,6diphosphatase and, 615,618, 632-633, 640, 641, 646 Seeds, acid phosphatase in, 451 Selenium, ribonuclease and, 692 Selenium, sulfur replacement in alkaline phosphatase, 390-391 Semen, acid phosphatase of, 469 Sephadex, adenosine aminohydrolase and, 55, 56 urease and, 3-4,6, 7 Serine residues, alkaline phosphatase, bacterial, 379, 380, 3 W 9 7 , 398 mammalian, 419, 424425, 439, 445, 446 prostatic acid phosphatase, 471472 pyrophosphatase, 513, 515 ribonuclease, 658, 666, 667, 671, 698699 modification of, 696 Serratia marcescens, asparaginase of, 116, 119

SUBJECT INDEX

Serratia marcescens (cont.) episomal transfer to, alkaline phosphatase and, 385 ribonucleoside Y,%-cyclic phosphate diesterase, 356 cellular localization of, 362 Serum, acid phosphatase of, 495-496 adenosine aminohydrolase, 55, 56 kinetic properties, 57 Sheep, glucose-6-phosphatase in, 600, 603 Shigella sonnei, 5'-nucleotidase of, 340, 349, 350 ribonucleoside 2', %-cyclic phosphate diesterase, 356 cellular localization, 362 kinetic constants, 360 substrate specificity, 357 Sialic acid, acid phosphatase and, 454 alkaline phosphatase and, 419, 424 deoxyribonuclease I and, 293 Silicate, glucose6phosphatase and, 581 Silicotungstate, pyrophosphatase electron microscopy and, 506-508 Skin, acid phosphatase, electrophoresis of, 454, 455 pyrrolidone carboxylate in, 143-144 Slime molds, fructose diphosphatase in, 640, 646 Snake venom, diesterase, 188-189 Sodium cetyl sulfonate, glucosebphosphatase and, 557 Sodium chloride, asparaginase sedimentation constants and, 113 venom exonuclease and, 319 Sodium dodecylbenzene sulfonate, glucose-6-phosphatase and, 557 Sodium dodecyl sulfate, fructose diphosphatase and, 627, 629, 638 glucose-6-phosphatase and, 557, 580 pyrophosphatases and, 512, 530531 ribonuclease and, 735-737 urease and, 9, 11 Sodium ions, 5'-adenylic acid aminohydrolase and, 67-68 deoxyribonuclease I and, 302, 303

Soil, urease in, 15 Solid phase peptide synthesis, staphylococcal nuclease analogs and, 199-200 Solvent perturbation spectroscopy, ribonuclease, 717 Soybean, urease of, 18, 19 Species, acid deoxyribonucleases and, 285 S-Peptide(s), synthetic, summary of, 699-703 Spermine, ribonuclease denaturation and, 735 Spheroplasts, alkaline phosphatase and, 374, 375-376, 377, 378 ribonucleoside 2', %-cyclic phosphate diesterase and, 361 Sphingomyelin, 5'-nucleotidase and, 345 Spinach, fructose diphosphatases of, 641, 642, 643 pyrophosphatase of, 540 Spleen, acid deoxyribonuclease, catalytic properties, 276-285 chemical and physical properties, 27Z276 distribution, localization and role, 285-287 acid phosphatase of, 451,493-495 adenosine aminohydrolase, 50 properties of, 55, 56, 57 deoxyribonuclease inhibitor of, 299, 301 diesterase, 188-189 phosphatases of, 418, 420 Spleen acid exonuclease, catalytic properties, artificial substrates, 33b334 natural substrates, 331-333 pH, activators and inhibitors, 334-335 distribution and localization, 336 isolation, purity and physical properties, 330-331 nomenclature, 329-330 Spleen exonuclease, point of attack, 315 terminal identification and, 326 Staphylococcal nuclease, active site, size and specificity, 191-195

SUBJECT INDEX

891

stereochemical probes, 195-1913 Subtilisin, amino acid sequence, 162,180 5‘ + 3’ exonuclease and, 258 behavior in solution, 183-184 ribonuclease and, 669-672 choice for X-ray crystallography, Subunits, 153-156 alkaline phosphatase, 380-384,424 covalent structure, 180-183 isoaymes and, 385-386 crystallographic studies, bacterial asparaginase, 113-114, correlation with studies in solution, 115-116 172-174 fructose-l,6diphosphatase, 627, introduction, 156-159 637-638,645-646 fragments, complementation of, pyrophosphatase, 508-509 196-199 optical properties, 510 historical, 177-178 reconstitution of native enzyme isolation, 178-179 from, 510-512 mechanism of, 174-175 sedimentation and diffusion peptide chain conformation, 159-163 coefficients, 509 polynucleotide substrates, size, 509 kinetic measurements, 186-187 Succinate, spleen acid deoxyribonuclease specificity, 185-186 and, 281, 283 refolding of, 184 Sucrose density gradients, ureme and, substrate specificity, 174 6-7 synthetic analogs of, 199-204 Sugars, synthetic substrates and inhibitors, glucose-6-phosphatase and, 579 kinetic measurements, 190-191 ribonuclease and, 752,754,782-783 specificity, 187-189 Sulfate ions, unit cell parameters, 156 ribonuclease crystals and, 657,785, Staphylococci, p-Iactamases of, 26 786, 788, 802,804 spleen acid deoxyribonuclease and, Staphylococcus aureus, 281,283 acid phosphatase of, 498 Sulfhydryl groups, p-lactamase, 26 adenosine aminohydrolase, 58 amino acid composition and 5’-adenylic acid aminohydrolase, 70 sequence, 33,35, 36-37 alkaline phosphatase and, 380, 428 catalytic constants, 41 asparaginase, 110 dissociation constants, 38 fructose diphosphatase and, 614 inhibitors, 43 fructose-l,6-diphosphatase, 629 molecular properties, 32 modification of, 621-622,633 purification, 29 glutamyl transferase, 95-96 substrates, 34 prostatic acid phosphatase, 469 methionine auxotroph, nuclease of, urease active site, 19,20 182-183 venom exonuclease, 318 Streptococcus pyogenes, deoxySulfhydryl inhibitors, glucose-6ribonuclease of, 2W261 phosphatase and, 581 Streptodornase, properties of, 260-261 Sulfite, Streptomyces albogriseolus, ribonuclease adenosine aminohydrolase and, 56 of, 245 ribonuclease and, 691 Streptomyces erythreus, ribonuclease of, urease and, 6,10,12 245 4-Sulfony1oxy-2-nitrofluorobenzene, Strontium, staphylococcal nuclease and, ribonuclease and, 680 187 N-2-p-Sulfophenylazoguanylylbond, Subsites, staphylococcal nuclease, 193 ribonuclease TIand, 216

SUBJECT INDEX

Sword bean, urease, inhibition of, 16 Synovial fluid, alkaline phosphatase of, 423,424,435,436,437

T Takadiastase, adenosine aminohydrolase of, 59, 61, 62 Tartrate, acid phosphataae and, erythrocyte, 482 fungal, 497 Gaucher’s disease, 496 liver, 487, 490 platelet, 495 prostatic, 454, 457, 462465, 470, 471, 473 spleen, 494 Temperature, acid phosphatase and, 454, 458-459, 481 alkaline phosphatases and, 426,438 bacterial asparaginase and, 109 glucose6-phosphatase and, 568, 577478 glutaminase and, 88,89, 93 p-lactamases and, 42-43,44 microbial ribonucleases and, 244-248 ribonuclease A, 051,653 conformation and, 726-729, 738 ribonucleaae NI and, 232,233 ribonucleaee TI and, 213, 218,223 ribonuclease T, and, 224 ribonucleoside 2’,3’-cyclic phosphate diesterase and, 358,359,360 staphylococeal nuclease and, 184 Testosterone, prostatic acid phosphatase and, 457 Tetraethylthiuram disulfide, urease and, 11 Tetrahymena pyrifomnis, ribonucleases of, 248 Tetramers, alkaline phosphatase, 384, 403, 406 Tetranitromethane, ribonucleaae A and, 698 ribonucleaae T, and, 221 staphylococcal nuclease tyrosine residues and, 172,195 Tetrapolyphosphate, pyrophosphatase and, 520422,525,526

Theophylline, 3, 5‘-cyclic phosphate diesterase and, 368,369,370 Thermodynamics, glucose-6-phosphatase and, 571-572 ribonuclease conformational transitions, 740-744 Thienylalanine, alkaline phosphatase, 389390 Thiobacillus thwparus, ribonucleases of, 245 Thioglycerol, urease and, 9 Thioglycollate, ribonuclease and, 691 Thioguanine, guanine aminohydrolase and, 77 Thioguanosine 2’,3’-cyclic phosphate, ribonuclease U, and, 236 6-Thioguaylycytidine, ribonuclease TI and, 216 ribonuclease T,and, 226 Thiol reagents, p-lactamases and, 4344 Thiophosphate, alkaline phosphatase and, 394, 395 4-Thiouracil, nucleotides, ribonuclease and, 763, 797, 798 Thiourea, urease and, 14, 17 4-Thiouridylate, ribonuclease Tzand, 226 Threonine residues, alkaline phosphatase, 379 deoxyribonuclease I, 297 ribonuclease A, 666, 669, 787, 788, 798 modification of, 696 ribonuclease TI, 222 Thymidine, 2,4dinitrophenyl esters, venom exonuclease and, 328 Thymidinea, Sdiphosphate, staphylococcal nuclease, 155, 156, 163, 184, 196 binding of, 163-171, 203 Thymidine triphosphate, pyrophosphate trapping and, 519-520 Thymine dimers, excision of, 269-270 venom exonuclease and, 321 Thymolphthalein monophosphate, alkaline phosphatase aasay and, 433 Thymus, deoxyribonuclease inhibitor of, 299301

SUBJECT INDEX

Trinitrophenylation, ribonuclease TI and, 221 Tripolyphosphate, 5‘-adenylic acid aminohydrolase and, 65 polyphosphatase and, 520-522, 525, 526 605-606 Tissue cultures, urease in, 14 Tris, alkaline phosphatase and, 398, 406, Tobacco, 407,408, 410,412 leaves, acid phosphatase of, 497 Triton X-100,glucoseb-phosphatase Tobacco mosaic virus, and, 557,570,580 Trypan blue, liver acid phosphatase prostatic acid phosphatase and, 476 terminal identification, 326, 327 and, 488 Trypsin, Tomato juice, pyrrolidone carboxylate in, 128 alkaline phosphatase, 382-383 Torula utilis, guanine aminohydrolase isozymes and, 386 in, 51 asparaginase and, 115-116 1 ,l-Tosylamide-2-phenethyl-chlorodeoxyribonuclease I and, 299 methyl ketone, spleen acid deoxy5’ + 3’exonuclease and, 268 ribonuclease and, 276 glucose8phosphatase and, 553 Transferase activity, alkaline phosinhibitor of, 301 phatase, 43!3-440 pyrophosphatase and, 514 Transfer ribonucleic acid, ribonuclease A, 670,6734374,691 deoxyribonuclease and, 259 transition temperature, 727 ribonuclease TI and, 218 ribonuclease TI and, 222 ribonuclease T, and, 226,227,230 spleen acid deoxyribonucleases and, ribonuclease U, and, 236,237-238 274, 275 staphylococcal nuclease and, 155, 163, spleen acid exonuclease and, 331-332, 333 181, 182,195,19&197,198 venom exonuclease and, 319, 321 Tryptazan, alkaline phosphatase and, Transphosphorylation, 390 alkaline phosphatase and, 406-409 Tryptophan , erythrocyte acid phosphatase and, fluorescence, staphylococcal nuclease, 481482 184, 200 prostatic acid phosphatase and, fructose diphosphatases and, 633 472-473, 474 Tryptophan residues, ribonuclease, hydrolysis and, 788-791, alkaline phosphatase, 383,391 799 replacement of, 390 Triazolealinine, alkaline phosphatase deoxyribonuclease I, 297 and, 390 fungal ribonucleases and, 210 Tricarboxylic acid cycle intermediates, prostatic acid phosphatase, 471 fructose diphosphatase and, 631 pyrophosphatase, 512 glucose6phosphatase and, 579 ribonuclease TI and, 219, 221 Trichodenna koningi, ribonucleases of, spleen acid deoxyribonuclease, 276, 281 247 staphylococcal nuclease and, 204 Triethylammonium phosphomolybdate, urease, 13 phosphate recovery and, 432-433 venom exonuclease, 318 Trifluoroacetylation, staphylococcal Tubercidin 3’, &-cyclic phosphate, cyclic nuclease, 197-198 phosphate diesterase and, 367 Triiodide ion, ribonuclease and, 684 Tumors, n-glutamate in, 136 2,4,6-Trinitrobenzene sulfonate, Tweens, glucose6-phosphatase and, pyrophosphatase and, 516-516 557, 580 Thyroid, adenosine aminohydrolase of, 55 Thyroxine, glucose-6-phosphatase and, 598 Tissue (a), glucose-6-phosphatase in,

a94

SUBJECT INDEX

Tyrosinase, prostatic acid phosphatase and, 471 Tyrosine, alkaline phosphatase and, 443 glucose-6-phosphatase and, 578,579 Tyrosine residues, alkaline phosphatase, 383,391,403 asparaginase, 115 fructose-l,6diphosphatase, 619-820, 629,631,633,637

fungal ribonucleases and, 210 prostatic acid phosphatase, 470471 pyrophosphatase, 515, 518 ribonuclease A, 658, 666, 676, 677, 678, 683, 692, 694, 699, 711, 721, 722723, 726

acid transition and, 730 fluorescence and, 718-719 modification of, 684-1335, 690,698, 719 normalization of, 737 organic solvents and, 7&734 sodium dodecyl sulfate and, 735 ultraviolet absorption and, 714-717 urea transition and, 732, 738 ribonuclease TIand, 219,221 staphylococcal nuclease, 155, 161, 167, 172-174, 195, 202

venom exonuclease, 318

U Ultraviolet absorption, fungal ribonucleases, 208, 210 pyrophosphatase, 505-506 subunits, 510 pyrrolidone carboxylate determination and, 127 ribonuclease, 704, 714-717 thermal transition and, 727, 728 Ultraviolet repair enzymes, properties of, 26%270 Uracil, derivatives, ribonuclease and, 760 Uranyl acetate, pyrophosphatase electron microscopy and, 506 Uranyl salts, ribonuclease assay and, 749, 757

Urea, adenosine aminohydrolase and, 56, 58-59, 74

alkaline phosphatase and, 380, 426, 442 asparaginase and, 114,115, 117 deoxyribonuclease I inhibitor and, 299 erythrocyte acid phosphatase and, 481 fructose diphosphatase and, 637 glucosed-phosphatase and, 580 8-lactamases and, 45 5’-nucleotidase and, 341 prostatic acid phosphatase and, 487 pyrophosphatase and, 509-531 ribonuclease A and, 685, 691, 717, 729, 731-733, 738,739, 740,744

ribonuclease TI and, 221 spleen acid deoxyribonuclease and, 282

staphylococcal nuclease and, 155, 184 urease unfolding and, 8, 9, 20 Urease, activity, measurement of, 4-5 carboxymethyl and aminoethyl derivatives, 12-13 catalytic properties, active site studies, 20 kinetic studies, 18-20 mechanism, 15-16 substrate specificity, 16-18 difficulties in studying, 2-3 inhibitors of, 17 jack bean, isolation and purification, 2-5

molecular properties, 5-8 chemical composition and behavior, 11-12

derivatives, 12-13 immunological behavior, 13 molecular weight determination, 8-10 other physical properties, 16-11 other sources, 13-15 related enzymes, 1 ultraviolet absorption, 5, 10 Uric acid, production, guanine aminohydrolase and, 77 Uridine, derivatives, ribonuclease and, 760, 762, 767, 774-777, 785, 798, 799-800, 802,

804 Uridine 3’, 5’-cyclic phosphate, diesterase and, 370 Uridine 2’,3’-0,O-cyclophosphorothioate. ribonuclease and, 758, 794

895

SUBJECT INDEX

Uridine diphosphate-sugars, 5’-nucleotidase and, 338-339 Uridineb-oxyacetate, ribonuclease T1and, 227,230 ribonuclease Uzand, 236 Urine, acid deoxyribonuclease inhibitor in, 281 acid phosphatase in, 450,455 pyrrolidone carboxylate in, 128, 134, 135-136 ustilago sphaerogena, ribonucleases, 234-239 main properties, 208,248 Ustilago zeae, ribonuclease of, 248

V Valine residues, deoxyribonuclease I and, 297 ribonuclease, 658,666,669,671 staphylococcal nuclease and, 167, 182 Venom, 3’,5’-cyclic phosphate diesterase and, 370 enzymes hydrolyzing phosphate esters, 313-314, 328 Venom endonuclease, products, 328 Venom exonuclease, application to structure determination, identification of terminals, 326-328 ribooligonucleotide sequences, 324-326 chemical nature, 317-319 point of attack, 315 structural characteristics of substrs.tea affecting susceptibility, bases and, 320 conformation and, 319-320 monophosphoryl group and, 322-324 sugars and, 320-321 substrates of, 315-316, 334 Venom 5’-nucleotidase, properties of, 342, 350-351 Vibrio alginolyticus, ribonucleoside 2’,3’-cyclic phosphate diesterase, 356 kinetic constants and mechanism, 380,361

physical and chemical properties, 358,359 substrate specificity, 357 Vipera kbetinu, venom exonuclease, 318 Vipera russelli, 5’-nucleotidase of, 342 Viruses, ribonucleic acid, replication of, 242 Vitamin L, ribonuclease and, 772 Von Gierke’s disease, glucose-6phosphatase and, 568

W Water, acid phosphatase and, 462 alkaline phosphatase and, 446 fructose-l,6diphosphatase and, 615 glucose-6-phosphatase and, 583, 588, 587,590,591,594,596 glutaminase and, 84, 93-94 ribonuclease and, 779, 780, 785, 787, 789,796,797 urease activity and, 19 yeast pyrophosphatase and, 538,539 Wheat germ, acid phosphatase of, 452, 472 Wheat gluten, crude papain and, 140 Wheat seedlings, a’-nucleotidase of, 353-354

X Xanthosine 2’, 3’-cyclic phosphate, ribonuclease TIand, 216 Xanthydrol, ribonuclease and, 698 Xanthylate, ribonuclease Uz and, 237 X-ray diffraction, pancreatic ribonuclease, 654-669, 683, 692, 694, 697, 704-710 X-ray studies, alkaline phosphatase, 389, 406

Y Yeasts, asparaginase of, 103, 117 inorganic pyrophosphatase, assay, 534 divalent cation binding, 531-532 kinetics, 535-538

SUBJECT INDEX

Yeasts, inorganic pyrophosphatase (cont.) mechanism, 538-539 metalloenzyme, 532534 physicochemical parameters, 530-531 purification, 530 specificity, 534-535 B’-nucleotidase of, 341-342, 349,350 Yoshida ascites cells, adenosine aminohydrolase of, 55

Z Zinc ions, acid phosphatases and, 479 5’-adenylic acid aminohydrolase and, 70, 71

alkaline phosphatase, bacterial, 376, 379, 380,382, 384, 386, 391, 394, 401-404, 405, 410-411 mammalian, 419, 424, 426427, 439, 440-442, 443, 444-445 3’, 5’-cyclic phosphate diesterase and, 368 glucose-6-phosphatase and, 545 p-lactamase I1 and, 43, 44 3’-nucleotidase and, 353 5’-nucleotidase and, 339-340, 342, 346, 348, 349, 350-351 pyrophosphatase and, 540 bacterial, 618-519, 620 yeast, 532, 534, 535 ribonuclease A and, 770-771, 772 ribonucleoside 2’,3’-cyclic phosphate diesterase and, 359, 362-363

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Multiple Bonds between Metal Atoms, Third Edition

Multiple Bonds between Metal Atoms, Third Edition

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Volume IV

EHV Transmission, Volume Volume K, Third Edition

System Operation, Volume Volume L, Third Edition

Index, Volume Volume M, Third Edition

Enzymes

The Enzymes, Volume 24: Protein Methyltransferases (The Enzymes)

Energy Coupling and Molecular Motors, Volume 23, Third Edition (The Enzymes)

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Station Operation and Maintenance, Volume Volume G, Third Edition

Osteoporosis, Two-Volume Set, Volume 1-2, Third Edition

Protein Phosphate Protocols

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Station Planning and Design, Volume Volume A, Third Edition

Chemistry and Metallurgy, Volume Volume E, Third Edition

Boilers and Ancillary Plant, Volume Volume B, Third Edition

Handbook of Optics, Third Edition Volume IV: Optical Properties of Materials, Nonlinear Optics, Quantum Optics (set)

Engineering Mechanics: Statics (SI Edition), Third Edition (Volume 1)

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Enzymes, Volume IV: Hydrolysis: Other C-N Bonds. Phosphate Esters. Third Edition

The Enzymes, Vol V: Hydrolysis (Sulfate Esters, Carboxyl Esters, Glycosides), Hydration, 3rd Edition

Multiple Bonds between Metal Atoms, Third Edition

Multiple Bonds between Metal Atoms, Third Edition

Metal-Carbon Bonds in Enzymes and Cofactors

Volume IV

EHV Transmission, Volume Volume K, Third Edition

System Operation, Volume Volume L, Third Edition

Index, Volume Volume M, Third Edition

Enzymes

The Enzymes, Volume 24: Protein Methyltransferases (The Enzymes)

Energy Coupling and Molecular Motors, Volume 23, Third Edition (The Enzymes)

Engineering Mechanics: Statics, Third Edition (Volume 1)

Mucosal Immunology, Two-Volume Set, Third Edition

Corrosion 2 Volume Set, Third Edition

The Hydrolysis of Cations

Nuclear Power Generation, Volume Volume J, Third Edition

Station Operation and Maintenance, Volume Volume G, Third Edition

Osteoporosis, Two-Volume Set, Volume 1-2, Third Edition

Protein Phosphate Protocols

Control and Instrumentation, Volume Volume F, Third Edition

Station Planning and Design, Volume Volume A, Third Edition

Chemistry and Metallurgy, Volume Volume E, Third Edition

Boilers and Ancillary Plant, Volume Volume B, Third Edition

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