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

CONTENTS OF OTHER VOLUMES

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

CONTENTS OF OTHER VOLUMES

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

CONTENTS OF OTHER VOLUMES

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

This Page Intentionally Left Blank

The Enzymes VOLUME IV

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

This Page Intentionally Left Blank

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 I<. McGceney, Irish J . Med. Sci. 491, 502 (1966).

4

F. J . REITHEL

bein (9) with respect to the use of Sephadex.] The procedure of Lynn (12) also employed molecular sieves (17 ) . The jack beans grown in Japan, Canavalia ensiformis var. gladiata, present special problems in the isolation procedure. Hanabusa (18) has proposed a modified procedure for extraction without the use of acetone.

ENZYMIC ACTIVITY MEASUREMENT Sumner (1) defined the unit of activity as the amount of enzyme that liberates 1 mg of ammonia nitrogen from urea a t 20" in 5 min. Despite the fact that the phosphate buffer (used to maintain a pH near 7) is slightly inhibitory, as is ammonia, the assay has been widely used with reasonable satisfaction. Chin and Gorin (19) have studied this assay in detail and have shown that the liberation of ammonia is not linear and that the p H changes appreciably during the 5-min incubation. They reported that 1 Sumner unit = 14.28U;O where the latter is defined in accordance with IUB recommendations: a unit catalyzing the decomposition of 1 microequivalent of the bond involved in 1 min (20°,p H 7). Because of the shortcomings of the Sumner assay, Gorin and Chin (20) have proposed a more precise assay employing a trisEDTA buffer a t pH 9 (25"). This unit is symbolized as U r and 1 Sumner unit = 11.0 U:6. Comparing activities a t three different temperatures, Uz6 = 1.22 Uio and Ui6 = 0.79 Uzo. I n this assay the product is mostly ammonium carbamate, explicitly shown in the following equations: Ha-CO-NH2

+ 2H10

4

S

+ + +

+

NH4+ NHzC02H20 2NH4+ c0:NH4+ NH3 HCOI-

+

After a 2.0-min reaction time (two bonds broken per molecule urea), addition of excess HC1 stops the reaction and converts the carbamate to ammonium ion. Back titration with NaOH estimates the unreacted acid. A somewhat more elaborate assay has been proposed by Blakeley et al. (13) employing a recording p H stat. In this assay no buffer is used, but dithiothreitol ( D T T ) is present during the titration (that maintains the pH a t 7.0). The unit, IU, is defined as the amount that causes the decomposition of 1 pmole of urea per minute (38", pH 7.0, 0.05M urea, 2 pM DTT). One Sumner unit = 17.6 IU/mg = 11 UiK/mg.Specific 17. Personal correspondence with K. R. Lynn. Throughout this paper there are stated activities that are at least one order of magnitude too high. 18. K. Hanabusa, Nature 189, 551 (1961); 193, 1078 (1962). 19. C. C. Chin and G. Gorin, Anal. Biochem. 17, 60 (1966). 20. G. Gorin and C. C. Chin, Anal. Biochem. 17, 49 (1966).

1.

URWEB

5

activity is defined as I U per ml/A2,, where the denominator is 0.589. Other types of assay procedures proposed are based on diffusion (16, d l ) , potentiometry (22),and estimation with Nessler’s reagent (23). A brief review of procedures has recently appeared (24). The calculation of specific activity requires a reliable method for determining the weight of urease used. The most convenient method is measurement of the ultraviolet absorption but this is not highly accurate. 0.771 (6)for the absorption of 1 mg/ml Values of 0.589 (IS),0.754 (20), at 280 nm have been reported as well as 0.77 a t 277 nm (26).Urease isolated in the presence of P-mercaptoethanol yielded a value of 0.640 (20) and a t p H 2, 0.597 (6). Until far more data are a t hand, obtained in systematic studies to establish the relation of activity to concentrations of protein and the effects of ions, there seems little justification for attaching undue weight to specific activity determinations as an indication of purity. I n the author’s laboratory urease that appears homogeneous by electrophoretic and ultracentrifuge criteria has an activity of 160 Sumner units (at 20”) per milligram based on A,,, = 0.762 in turn based on dry weight measurements (26). The catalytic action of urease has a large temperature coefficient,and it would be highly desirable if a single standard temperature could be agreed upon, particularly that recommended by the IUB, 30”. It is convenient to have a method for the detection and rough estimation of urease activity in gels. A simple staining procedure using cresol red has been described (27),but a more elaborate procedure (28) employing p-nitro blue tetrazolium results in sharper definition.

111. Molecular Properties

Physicochemical studies on jack bean urease during the past 10 years undoubtedly were stimulated by the 1960 paper of Creeth and Nichol 21. J. A. Blain and M . Caskie, Chem. & Znd. (London) p. 17 (1965). 22. S. A . Katz. Anal. Chem. 36, 2500 (1964). 23. E. Bernt and H. U. Bergmeyer, in “Methods of Enzymatic Analysis” (H. U. Bergmcyer, ed.), p. 401. Academic Press, New York, 1963. 24. A . Kaplan, Methods Biochem. Anal. 17, 320 (1969). 25. C. J. Bailey and D. Boulter, BJ 113, 669 (1969). 26. R. Goodrich and F. J. Reithel, Anal. Biochem. 34, 538 (1970). 27. D. P. Blattler, C. C. Contaxis, and F. J. Reithel, Nature 216, 274 (1967) ; C. C. Contaxis and F. J. Reithel, JBC (in press). 28. W. N. Fishbein, Proc. Int. Symp. Chromatog. Electrophoresis, 6th, Brussels, 1968, p. 238. Presses Acad. Eur., 1969.

6

F. J. REITHEL

(29). I n this, the evidence strongly suggested that the persistent “impurities” with sedimentation constants greater than 20 were oligomers of the main component of urease preparations. The main component had an extrapolated s;,~ value of 20 k 0.5, the most prevalent “impurities,” 28 S and 36 S, suggesting a In, 2n, 3n relationship. It was noted that a 4-6 S, and occasionally a 12 S, peak could also be discerned. I n the presence of sulfite such preparations yielded only 19-20s values during ultracentrifugation and showed an increase in activity. The latter observation has been cited repeatedly as evidence that the 28 S and 36 S forms possessed lower specific activities than the 20 S form, but this has not been demonstrated with isolated oligomers. The action of sulfite on the 2 0 s form, isolated as such, has not been demonstrated. The ultracentrifuge evidence strongly indicated that the above forms were not in rapid equilibrium. This conclusion was strengthened by the experimental results of Siegel and Monty (SO) who chromatographed a commercial preparation of jack bean urease (of unspecified age) on Sephadex G-200. They obtained an elution pattern with four peaks, and from the elution volumes they calculated Stokes radii of 61, 79, 92, and 122A. If the first three components were assumed to have S values of 19, 28, and 36, then the molecular weights could be calculated. They were found to be in the ratio 1.0, 1.9, and 2.9. It was noted that only 10% of the sample used was soluble. Hill and co-workers (31,32) chromatographed freshly prepared urease on carboxymethyl cellulose in acetate-phosphate buffer, with and without sulfite. They succeeded in obtaining fractionation with respect to activity, but the fractions were heterogeneous with respect to the sedimentation values obtained. Sedimentation coefficients of 3, 3.9, 19, 28, and 32 were noted and there was no correlation that permitted an assignment of specific activity to each component. More recently (IS), it was found t h a t the activity of urease could be maximal even when 3Q% of the urease had sedimentation coefficients greater than 1 8 s . I n another laboratory (9) it was found that specific activity did not correlate with monodispersity. Several investigators have presented evidence for low molecular weight forms exhibiting urease activity. Hand (33), in 1939, obtained diffusion data indicating particles of 17,000 daltons or less that retained enzymic activity. More recently, sucrose density gradient ultracentrifugation (34) 29. J. M. Creeth and L. W. Nichol, BJ 77, 230 (1960). 30. L. M. Siegel and K . J. Monty, BBRC 19, 494 (1965). 31. M. Shadaksharaswamy and R. M. Hill, ABB 97, 607 (1962). 32. R. M. Hill and A. Elliot, ABB 113, 405 (1966). 33. D. B. Hand, JACS 61, 3180 (1939). 34. P. P. Sehgal, R. J. Tanis, and A. W. Naylor, BBRC 20, 550 (1965).

1. UREASES

7

has been used to demonstrate an active 1 2 s form of urease in a commercial preparation, in jack bean meal, and in extracts of soaked jack bean seeds. Less than 5% of the total activity was in this form. Urease preparations extracted with water and precipitated with NaCl and polyethylene glycol-4000 yielded (35) a form of urease with a s&, = 8.5. Urease exposed to 0.1 M acetate buffer, p H 3.5, has been reported (36) to be converted to a metastable, transiently enzymically active, species with szo,,, = 9.8. Using quantitative gel electrophoresis (37) and a simple activity stain (27), Blattler examined urease preparations showing evidences of “aging” changes (38,38,). When urease, initially electrophoretically homogeneous, was allowed to age in buffer or 50% diol, enzymically active species appeared that had electrophoretic mobilities between the usual bands. These intermediate bands corresponded to a variety of species that differed by approximately 60,000 molecular weight between 240,000 and 480,000, and between 480,000 and 960,000. Fishbein and co-workers (9,39) have examined a variety of urease preparations by gel electrophoresis. The use of an improved staining technique (98) allowed the visualization of activity in bands corresponding to n, 2n,3n, 4n, 5n, 6n (where n = 480,000 daltons) . These bands were designated as polymeric isozymes. Other bands of activity corresponding to molecular weights of 240,000, 600,000, and 800,000 were observed and termed epiisozymes assumed to consist of multiple active assemblies of a single active subunit. Filtration of preparations (containing only n species) through columns of Sephadex G-150 or G-200 resulted in the appearance of 2n-6n species and a very minor amount of a form estimated to have a molecular weight of 17,000. Finally, the existence of conformeric isozymes was postulated on the basis of variations in electrophoretic mobility observed. The mechanism involved in the production of urease oligomers on gel columns has not been clarified and seems anomalous because all of the species were excluded from the gel. Moreover, as Creeth and Nichol observed (as), if an equilibrium exists, the kinetic constants are small. The demonstration of so many enzymically active forms of urease 35. K. K. Stewart and L. C. Craig, Federation Proc. 25, 590 (1966). 36. G. Gorin and C. C. Chin, Federation Proc. 26, 605 (1967). 37. D. P. Blattler and F. J. Reithel, J . Chromatog. 46, 286 (1970). 38. D. P. Blattler, Ph.D. Thesis, University of Oregon, 1967; Dissertation Abstr. B29, No. 1, 38 (1968). 38a. D. P. Blattler and F. J. Reithel, Enzymologia 39, 193 (1970). 39. W. N. Fishbein, C. L. Spears, and W. Scurai, Nature 223, 191 (1969).

8

F. J. REITHEL

with such disparate mobilities point to a complex association-dissociation process. However, until these several forms have been isolated and characterized, particularly with respect to specific activity, unequivocal interpretations of the data are not possible. Moreover, specific activity values per se are not measurements of precision unless they are related to a mass of a characterized conformer. And, finally, it must be shown that the enzyme retains the same character during the enzyme assay that it has in isolation conditions.

A. MOLECULAR WEIGHTDETERMINATIONS The molecular weight of recrystallized urease was carefully determined in Svedberg's laboratory by combining sedimentation velocity and diffusion data (40). The value reported ( 4 1 ) , 483,000, is substantially the same as that of a more recent study ( 6 ) reporting the value 489,000, obtained by sedimentation equilibrium. These values pertain to the most prevalent molecular species in freshly prepared urease. If the single polypeptide chain weight (n) in urease is 30,000, then Sumner's urease can be considered as (16n). This type of symbolism prevents misunderstanding in the discussion of polymorphic proteins with the proviso that (n) is defined. There is another active molecular species (8n) obtainable by treatment with propanediol, glycerol (27), or 1,2-ethanediol (4.Z))having a molecular weight of 236,000. Urease activity persists unaltered when the enzyme is dissolved in 8 M urea although the ultracentrifuge data indicate a molecular weight of about 90,000 ( 7 ) . This form of urease has not been sufficiently characterized but does indicate that neither enzymic activity nor specific activity is dependent upon very high molecular weight aggregates. Dissociation and Association The foregoing discussion provides numerous references to observations of the dissociation-association behavior of urease. It is common knowledge among those using this enzyme that association occurs but during storage. Disulfide bonding is not required for association (7,427) 40. J. B. Sumner, N. Gralen, and I.-B. Eriksson-Quensel, JBC 125, 37 (1938). 41. The value of 483,000 is in the original publication but was misquoted by both Sumner ( 1 ) and Varner (3) as 473,000. Moreover, the diffusion constant was determined in the presence of sodium sulfite but the sedimentation constant was not. The main component in phosphate buffer was found to have a sedimentation constant of 18.65 a t 20". 42. G. Gorin, D. P. Blattler, and D. E. Thain, BBRC 36, 1045 (1969).

1. UREASES

9

may occur if not prevented as suggested by the findings of Creeth and Nichol (29). Evidence of dissociation under various conditions has been noted ( 29, 3 2 4 6 ). The effects of “dissociating agents” have been described. In 6 M guanidine hydrochloride a value of 83,300 was found ( I d ) , but this value was probably the result of partial reassociation ( 4 3 ) . Urease dissociates even in 3 M guanidine hydrochloride to (2n) subunits of 60,000 daltons (71, but it reaggregates upon standing. Ultracentrifuge studies (44) as well as gel electrophoresis studies (46) of urease in sodium dodecyl sulfate (SDS) suggest dissociation to a molecular weight of 6O,OOO, with total loss of activity. Another gel electrophoresis study (26) in SDS and p-mercaptoethanol yielded a mobility corresponding to a molecular weight of 75,000. Dissociation of urease in 0.1% SDS in 45% thioglycerol, followed by dialysis against tris-EDTA, p H 9.2, yielded a 120,000 molecular weight species (4n) as judged from ultracentrifuge data ( 7 ) . Urease is remarkably resistant to unfolding in urea. No change in optical rotatory dispersion or specific activity of the enzyme occurs in concentrations of urea up to 8 M ( 7 ) . Between 8 and 9 M , unfolding occurs and the molecular weight becomes 60,000, but some activity persists. Solution in 0.1 M acetate buffer, p H 3.5, resulted in an enzymically active species (8n) of 240,000 daltons (46). Tanis and Naylor (4’7) have reported that a t low concentration of protein the 18s form predominated above pH 5.3 and the 1 2 s form below pH 4.8. Between these p H values a rapid equilibrium of the 12 S and 18 S species was observed The dissociation behavior of urease a t low pH depends on the buffer used. I n 0.1 M potassium phosphate buffer, adjusted to p H 2.0 with HC1, a heterogeneous mixture of dissociated forms was obtained ( 6 ) with an Mw of about 150,000. I n acetate buffer a t pH 3.5 dissociation into a 120,000 molecular weight species (4n) was observed (48). I n 34% acetic acid a t p H 2.2 there is effected a dissociation to subunits (n) of 30,000 daltons (7). This same value was obtained for urertse ultracentrifuged in 8 M urea 0.5 M thiol and in performic acid oxidized urease (48).

+

A. Andrews and F. J. Reithel, ABB 141, 538 (1970). G. Gorin, G. Mamiya, and C. C. Chin, Experieiitiu 23, 443 (1967). D. P. Blattler and G. Gorin, Can. J . Biochem. 47, 989 (1969). G. Gorin, C. C. Chin, and S. F. Wang, E x p e r i e n t k 24, 685 (1968). R. J. Tanis and A. W. Naylor, BJ 108, 771 (1968). F. J. Reithel and J. E. Robbins, Proc. 7th Intern. Congr. Biochem., Tokyo, 1967 Abstracts IV, p. 585. Sci. Council Japan, Tokyo, 1968. 43. 44. 45. 46. 47. 48.

10

F. J. REITHEL

At this time there is no definitive evidence concerning either the identity of the subunits or the groups involved in their interaction. Since it is possible to observe the dissociation of urease in four discrete steps to (8n), (4a), (2n), and ( a ) subunits, it seems likely that a t least four types of binding arrangements are present. Since the dissociations are symmetrical it seems likely that the subunits are identical a t least down t o 60,000 daltons. At least two groups (1S,2.5) have evidence for a subunit weight of 75,000-80,000 daltons. This is supported by other evidence (14,445). It is likely that these data are artifactual and do not relate to a structural subunit weight.

B. OTHER PHYSICAL PROPERTIES The diffusion constant a t 20" determined many years ago (400) in cm2 sec-l remains unchallenged. The partial sodium sulfite as 3.46 X specific volume value of 0.73 reported (40) has been verified by recent measurements using other techniques (6). At p H 7 and 25", 0.733 was found; a t pH 2 and 25", 0.734; a t p H 7 and 5",0.722. The wavelength of maximum absorbance is 278.5 nm (13,20).The absorbance values reported show wide divergences: A2,8.G(1mg/ml) = 0.601, A280 0.589 (IS) ; A 2 7 8 = 0.771, A 2 7 8 (PH 2) = 0.597, A 2 7 8 (6 M GCl) = 0.671 (6, 14) ; At78 (PH 7) = 0.754 (20); A 2 7 8 = 0.64 (4.2, 45) ; A 2 7 8 = 0.80 (49). Since protein concentration is often estimated by measuring the absorbance, it is evident that the spectrum of activities to be found in the literature very likely reflects the spectrum of values given above. The AZ80/&60 ratio has been reported as 1.8-1.9 ( 1 3 ) . The isoelectric point, determined by Sumner and Hand (50) to have a value of 5.0-5.1, has been redetermined by the electrofocusing technique (43). The value obtained was 4.8. The solubility is extremely small a t this pH, but the urease can be located by its enzymic action. It is a point of interest that the solubility of the isoelectric species increases spectacularly if the sulfhydryl groups are substituted with N-ethylmaleimide (NEM) (4.9). Evidence for conformers of urease has been presented by two groups of investigators. Chernitskii et a2. (51) noted fluorescence maxima a t pH 3 4 , 5.5-6.5, 7.5-9, and >10.5 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<,,, and V,,,:,,for several ureases. The activation energy of urea hydrolysis catalyzed hy ureasc h a s been reported as 8.0 kcal/mole (103) and 9.8-12.2 kcal/mole (101). Ti1 one report no temperature anomalies were evident (103) although, again, it is difficult to know what ureaw species was being invcstigated sincc. glycerol was being used (27). In contrast, another invrstigatioii (104) revealed anomalous temperature tlrpendencc of hydrolysis rates as did an earlier one ( 7 7 ) . A prcparation of soybean urcase in sulfitc buffer was used. Activation energies similar to those ahovc were reported. 101. I<. R. 1,ynn and P. E. Ymkwiclr. BBA 56, 512 (1962). 102. K. R . Lynn :ind P. E. Ynnkwic.11. BBA 81, 533 (1964). 103. R . J. Millcr. C . Pinkllnm. .I. H . Orermnn, and S. W. Dumfortl BBA 167, 607 (1968). 104. G . Talsky and G . K1unkc.r. 2. Physiol. (‘hem. 348, 1372 (1967).

1.

19

UREAELEG

K,

AND

TABLE 111 V,,,,, VALUESFOR UREASEFROM VARIOUSSOURCES

T Buffer

Sources

pH

("C)

K , X 1W (mole/ liter)

V,.. (mole/ li ter/sec)

Ref.

~~

Phosphate Phosphate Phosphate Sulfite Thiosulfate Tris-HC1

S S C B S S

J J

7 7 7 5.7 6.7 7.7 7 7 7.4 7.0

25 25 37 20 20 20 25 25 25 38

20 19 30 100 130 40 4 76 66 4 3.28

15.4 X 10.5 X

37 17.2

x x

10-8

(Arbitrary units) Maleate

J

TrieHtSOI

J

J

5.0 7.0 7.0 9.0 8.0

21

20.8

4.0 4.9 5.1 2.7 4

2.5 x 6.7 x 8.4 x 1.7 x 15.5 x

10-3 10-3 10-3 10-3

a Abbreviations: S, soybean; C, Corynebacterium renale; B, Bacillus pasleurii; and J, jack bean.

The relative reaction rates of urease at low water activities have been measured (105). Mixtures of lyophilized urease and 14C-urea were exposed to various relative humidities at 20". The rates observed ranged from negligible activity below 60% huiiiidity to full activity at 100% relative humidity. It was estimated that hydrolysis could occur only after the water content exceeded onc molecule per polar group. There is an extensive older literature describing effects of various ions on urease activity. A recent study employing potentiometry (106) indicated inhibition by copper and zinc, questionable inhibition by nitrate, and slight activation by phosphate. Use of the same technique (107) has provided data showing 50% inhibition is produced by Hg". Cu'*, ZnL+,Cd"', Co2+,and Ni'+ arraiigeil in decreasing effectiveness (increasing coiiccntration required) . Tho data were held to provide support for the ol)inion that inhibition results from metal substitution of the sulfhydryl group. The pH of maximal catalytic activity in maleate buffer was found to be 6.5 compared to 7.5 in tris buffer [cited in Lynn ( 1 2 )I . The 105. J . J . Skujins :ind .4. L). McI,arrn, Science 158, 1569 (1967). 106. S. A . Kntz and J . A. Cowans, HBA 107, 605 (1965). 107 R . B. Hughes. S. A Katz, nnd S. E. Stuhbins. Enzymologia 36, 332 (1969).

20

F. J. REITHEL

rate of hydrolysis a t p H 6.9 has been reported higher in potassium phosphate buffers than in sodium phosphate buffer (108).

D. ACTIVESITESTUDIES Three types of evidence provide a basis for speculation concerning the nature of the catalytic site. First, kinetic studies have provided data for plots of R,,‘and V,,,;,,vs. pH (12). From such plots molecular ionization constants can be deduced. Lynn’s interpretations (12) suggest a histidine group with a pK,, valuc of 5.8, an a-ammonium group with a pK, value of 7, and a sulfhydryl group with a pK, value of 8.3. It was suggested that the ammonium and sulfhydryl groups are involved in the formation of the ES complex, the binding, and that the histidine group is involved in the reaction of the complex, the catalysis. Second, the correlation of change in enzymic activity with the titration of “essential” sulfhydryl groups has led to a postulation of eight active sites per 480,000 ( 5 6 ) . Unfortunately, the possibility of structural changes during such titrations makes interpretation of such data equivocal. However, the observation that urease retained its activity in 8 M urea, where the molecular weight has been reduced a t least to 90,OOO ( 7 ) , supports the conclusion above. Third, inhibitor binding studies have led to the conclusion that only two active sites are present in a (16n) structure (94). This conclusion is based on the characterization of a complex containing only 2 moles of hydroxamic acid per (16n) urease and the demonstration that this complex has no catalytic activity. Again, the possibility of structural changes cannot be excluded. Obviously the number of active centers postulated until now are fewer than the number of protomer?, but there is fair evidence for a t least eight active regions per molecule of crystalline enzyme or one catalytic region in a (2n) unit.

VI. Summary

Until our knoweldge of the iireases discloses biological significance the research on these enzymes will continue to be restricted. Inquiries into the mechanism of the urease catalysis are clearly appropriate 108. D. Watson, Clirt. C’hirn. Aclo 14, 571 (1966). 109. M. C. Wall and K. J. Laidler. A B B 43, 299 (1953).

1.

UREASES

21

since both specificity and efficiency are remarkable. I n the past such inquiries have suffered because the urease preparations used were not demonstrated to contain a single species. The seeming arbitrariness of the kinetic data reflect the lack of discrimination and control in securing a catalyst of known and reproducible properties. Similarly, studies of the molecular structure of urease have suffered because care in preparation did not match the concern for precision in physical measurements. It is now possible to prepare and characterize ureases in a suitable state of homogeneity. The literature may soon reflect this opportunity to carry out investigations, both chemical and physical, of more enduring value.

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Penicillinase and Other P-Lactamases NATHAN CITRI I . Introduction . . . . . . . . . A . Background . . . . . . . B. Definitions and Specificity . . . . C . Occurrence . . . . . . . . D . The Catalytic Reaction . . . . . I1. Molecular Properties . . . . . . . A . Purification and Physical Properties . . B. Composition and Sequence Analyses . . I11. Catalytic Properties . . . . . . . . A . Methods of Assay . . . . . . B . Kinetics and Substrate Specificity . . . C . Structural Modifications in Substrates . D . Structural Modifications in the Enzyme . E . Other Factors Affecting Activity . . . IV . Conformation and Function . . . . . . A . Nonspecific Conformational Transitions . B . Specific Transitions: Conformative Response V . Immunological Studies . . . . . . .

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23 23 25 26 27 27 27 31 35

35 39 40 41 42 44 44 45 46

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1 Introduction

A . BACKGROUND In 1940 Abraham and Chain ( 1 ) reported that an extract of Escherichia C O Z ~ destroys the antibacterial activity of benzylpenicillin . The authors presented evidence indicating that the extract (and possibly several 1. E . P. Abraham and E . Chain, Nature 146. 837 (1940).

23

24

NATHAN CITRI

other bacterial preparations) contained a penicillin-inactivating enzyme which they named penicillinase. Subsequently, many other bacteria were found to contain or secrete similar enzymes for which the inore general term, p-lactnmnse, Iins I ) c c i i suggesttxl. Tlicrc is 110 autliciiticntcd evidence that organisms other than bacteria produce p-lactamase ( 2 ) . The apparently exclusive and wide distribution of p-lactamascs anioiig diverse bacterial species suggests that this family of enzymes may have cvolvctl in connection with a uniquely bacterial function such as bacterial crll wall formation ( 2 6 ). The role of 1wnicillinasc and other ,8-luctnmascs i n conferring resistance to penicillins a i d the closcly relatctl cephalosporiiis has been amply demonstrated (6-8),niitl rcpcatcdly coiifirmctl in tlic numerous reports tlealing with the clinical aspects of p-lactmiase activity (reviewed in references 6, 7 , 9, 10).At a more fundaincntal lcvc~l,penicillinase has been studied most c.stcnsivcly from the point of view of enzyme induction ( 2 , 11, 1 2 ) , enzyme secretion (13-16), and transfer of genetic elements (17, 18). Earlier investigations a t the niolecular level, which have beeii reviewed in previous editions of this book (.19,20) , have been recently cbspanded to cover additional p-lactamase preparations. The picture which emerges indicates an unexpected diversity of size, structure, and molecular properties. Even more surprising is the variety of catalytic activities, which became increasingly evident as more p-lactamase preparations were iso2. N. Citri and M . R . Pollock. A ~ U U IEnzymol. I. 28, 237 (1966). 3. M. R. Pollock. Brit. M e d . J. IV, 71 (1967). 4. H. G. Boman. K. G. Eriksson-Grennberg, J . Foldes, and E. B. Lindstrom, in “Regulation of Nil[-lric A4cid and Protein Biosynthcsis” (V. V. Koningaherper and I,. 13osch. eds.). Vol. 10. 11. 366. B. B. A . Library, Elsevier Publ. Co., Amstcrdam. 1967. 5. I,. G. Burman, K. Nordstrom. and H. G . Bomnn, J. Bacteriol. 96, 438 (1968). 6. M. Barber, in “The Scientific Basis of Medicine,” Annual Review, 1964, p. 169. Univ. of London, (Athlone), London, 1964. 7. W. M. M. Kirby :ind R. J. Bulger. Ann. 12ee. M e d . 15, 393 (1964). 8. H. C. Neu, BBRC 32, 258 (1968). ‘3. E. Rauenbusch, Antibiot. Cliemolhercipia 14, 95 (1968). 10. J . T. Smith, J . M . T. Hamilton-Miller, nnd It. Knox, J . Phurm. Phurmtrcol. 21, 337 (1969). 11. M. R. Pollock, “The Enzyrncs,” 2nd ed., Vol. 1, p. 619. 1959. 12. M. H. Richmond. Essays Biocliem. 4 , 105 (1968). 13. J . 0. Lxnipen. J . Gen. Microbiol. 48, 249 and 261 (1967). 14. M. R. Pollock. J . Gcn. Microbiol. 26, 239 :ind 267 (1961). 15. D. J . Kushner and M. R . Pollock, J. Gett. Micl-obiol. 26, 255 (1961). 16. D. J . Kushner, J. Gen. Microbiol. 23, 381 (1960). 17. R. P. Novick, Bucteriol. Rev. 33, 210 (1969). 18. M. H. Richmond. A d v n n . Microbiol. Fhysiol. 2, 43 (1968). 19. E. P . Abraham. “The Enzymes,” Vol. 1, Pnrt 2, 1’. 1170, 1951. 20. M . R. Pollock, “The Enzymes,” Vol. 4, p. 269, 1959.

2.

PENICILLINASE AND OTHER

P-LACTAMASES

25

lated and studied against the larger number of scmisyiithetic penicillins and cephalosporins which became available in recent years.

B. DEFINITIONS A N D SPECIFICITY The term p-lactaubase denotes an enzymc which catalyzed thc hydrolysis of the amide bond in the p-lactam ring of 6-amino-penicillanic acid (6-APA) or 7-amino-cephalosporanic acid (7-ACA) and of their iV-acyl derivatives ( 2 ) .Such derivatives arc commonly referred to as penicillins [Fig. 1 (I) 1 and cephalosporins IFig. I (III), (V), and ( V I I ) ] , respectively. There is no evidence that any bond other than thc ainide bond in the intact nucleus of penicillin or cephalosporin is broken by the

(V11)

(VIII)

FIG.1. General structure of substrates tind products of the p-lactamase reaction, Substrates: (I) penicillins; (111). ( V ) , and ( V I I ) rephalosporins. Products: (11) penicilloic acids; ( I V ) , (VI), and (VIII) cephalosporoic acids.

26

NATHAN CITRI

enzyme ( 2 1 ) . Thus the catalytic specificity of p-lactamases appehrs at present to be confined to these two closely related families of antibiotics. The term penicillinuse has been retained in common usage to denote p-lactamase preparations which predominantly catalyze the hydrolysis of penicillins. Conversely, preparations which are more active against cephalosporins have been frequently referred to as cephulosporinases. It is expected that this distinction will disappear as more p-lactamases are found to show a continuous spectrum of activity against penicillins and cephalosporins.

C. OCCURRENCE Although @-lactamaseis widespread among the various bacterial species ( 2 , 9 ) ,the levels of p-lactamase activity within each species are extremely variable. The range (22) and significance of such variations have been the subject of a recent discussion by Pollock ( 2 3 ) . Indeed, studies prompted by variations in the penicillinase-forming ability of bacteria led to important advances in our understanding of the transfer and control of genetic information. In the present context the following observations may be relevant: (1) The gene for p-lactamase is readily lost (or acquired) in strains of bacteria (e.g., staphylococci and enteric bacteria) (24) where it is carried by an extrachromosomal particle (18,2 6 ) . (2) The gene expression (i.e., rate of p-lactamase formation) is often controlled by the mechanism of induction ( 2 , 1 1 , 2 7 ) . (3) A single mutational event, resulting presumably in the replacement of a single amino acid residue, may cause a large decrease in activity (3, 23). 21. The suggestion has been made that certain peptides which have been studied as inducers of p-lactamase in Bacillus cereus and in S. atireus may serve as substrates for the enzyme [A. K. Saz and D. L. Lowery, BBRC 15, 525 (1964); 19, 102 (1965)l. However, there is no evidence that such peptides are in fact hydrolyzed by plactamase. 22. J. M . T . Hamilton-Miller, FEBS Letters 1, 86 (196S). 23. M. R . Pollock, Ann. N . 1’. Acncl. Sci. 151, 502 (1968). 24. In enteric bacteria the p-lactamase gene may be acquired by infection with an R factor (cf. Table IV). This led to the pertinent observation (25) that no plactamase should be regarded as “species-specific” until its gene is shown to bc rhromosomal. 25. N. Datta and M. H. Richmond, BJ 98, 204 (1966). 26. R . P. Novick, Ann. N . Y . Acad. Sci. 128, 165 (1965). 27. M . H . Richmond, Nature 216, 1191 (1967).

2.

PENICILLINASE AND OTHER

P-LACTAMASES

27

D. THECATALYTIC REACIYON By definition, p-lactamase catalyzes the hydrolysis of the P-lactam ring in penicillins or cephalosporins as shown in Fig. 1. The product of the reaction with a penicillin has been identified by Abraham et al. (28) as the corresponding penicilloic acid. Although few other cases have been as rigorously investigated, it is generally accepted that the action of a p-lactamase on a penicillin results in the formation of a relatively stable, single product, penicilloic acid [Fig. 1 (11) 1. An analogous reaction has been postulated for cephalosporins (29,30), but it soon became apparent that the presumed primary product is highly unstable (31, 32) except in the case of cephalosporin y-lactones [Fig. 1 (VII) and (VIII)]. In other cephalosporins the nature of the primary product appears to depend on the substituent R [Fig. 1 (III)] (33).When R’ = H, an unstable “cephttloeporoic acid” is formed [Fig. 1 (V) and ( V I ) ] which is a product analogous to the penicilloic acid. In contrast, when R’ is an acetoxy or pyridinium group, the reaction involves the loss of R’ with the formation of an exocyclic double bond and a rearrangement as shown in Fig. 1 (IV). The resulting compound (Amax = 230 nm) is unstable and undergoes fragmentation (33).

II. Molecular Properties

A. PURIFICATION AND PHYSICAL PROPERTIES The numerous reports of p-lactamase activity in various bacterial strains reflect the widespread occurrence and the clinical importance 28. E. P. Abraham, W. Baker, W. R. Boon, C. T. Calam. H. C. Carrington, E. Chain, H. W. Florey, G. G. Freeman, R. Robinson, and H. G. Saunders, in “The Chemistry of Penicillin” (H. T. Clarke, J. R. Johnson, and R. Robinson, eds.). Chaptrr 2. p . 10. Princeton Univ. Press, Princeton, New Jersey, 1949. 29. B. Crompton. M. Jago, K. Crawford, G . G. F. Newton, and E. P. Abraham. BJ 83, 52 (1962). 30. G. G. F. Newton and E. P. Abraham, Nature 175, 548 (1955). 31. L. D. Sabath, M . Jago, and E. P. Abraham, BJ 96, 739 (1965). 32. G. G. F. Newton, E. P. Abraham. and S. Kuwabara, Antimicrobinl Agents

Chemothempy p. 449 (1968). 33. J. M. T. Hamilton-Miller, G . G . F. Newton, and E. P. Abrahani, BJ 116, 371 (1970). 34. Similarly, two distinct p-lactamasee have been observed in pure cultures of Enlerobncter ( 3 6 ) . 35. T. D. Hennessey, J . Gen. Mksobiol. 49, 277 (1967).

TABLE I PURIFIED ~LACTAMASE PREPARATIONS Source of enzyme 1. Bacillus cereus 569/H

extracellular (8-lactamase I) 2. Bacillus cereus 569/H extracellular (8-lactamase 11) 3. Bacillus crrrus 569/H cell bound (8-lactamase 7 )

4. Bacillus cereus 5/B

ex tracellular

5. Bacillus licheniformis 749/C eutracellular 6. Bacillus licheniformis 749/C cell bound

7. Bacillus lirheniformis 6346/C extracellular and cell bound

Main purification steps 1. Selective adsorption on glass 2. Ammonium sulfate fractionation 1. Ammonium sulfate fractionation 2. Ethanol precipitation 1. Sonic disintegration 2. Selective adsorption on cellulose phosphate 3. 1. CJI-cellulose chromatography Acetone precipitation 2. Ammonium srilfate fractionation 3. 1. Ethanol precipitation Selective adsorption on cellulose phosphate 2. Ammonium sulfate fractionation 1. Lyxozyme and trypsin digestion 2. Ammonium srilfate fractionation 3. DEAIGcelliilose chromatography 1. Ammonium sillfate fractionation

Criteria of homogeneity

Ref.

Electrophoretic and sedimentation patterns Crystallized

36

ChZ-cellulose rechromatography

38

Electrophoretic and sedimentation patterns

39

Sedimentation pattern

40

DEAEcellulose rechromatography

40

Electrophoretic and sediment,ation pat terns

40

37

8. Staphylococcus aurcus (types A, B, and C) extracellular

9.

10.

11.

12.

13.

14.

a

1. Selective adsorption on

cellulose phosphate 2. CM-cellulose chromatography 3. Centrifugation on sucrose gradient 4. Electrophoresis on sucrose gradient 1. Ultrasonic disintegration Escherichia coli TEM (R-TEM)o cell bound 2. DEAEcellulose chromatography 3. Gel filtration Escherichia coli K-12 (Glla1)b 1. Release by spheroplasting 2. SE-cellulose chromatography cell bound 3. Chromatography on hydroxylapatite 1. Osmotic release Escherichia coli DB103 (ATSm)a 2. DEAE-cellulose chromatography cell bound 3. Selective adsorption on hydroxylapatite Escherichia coli W3630 ( R G N Z ~ ~ ) ~1. Ultrasonic disintegration 2. DEAE-cellulose chromatography cell bound 3. CM-cellulose chromatography 1. Ultrasonic disintegration Eschwichia coli W3630 (RGN14). 2. Streptomycin precipitation cell bound 3. DEAE-cellulose chromatography Enterobacter cloacae 214 1. Ultrasonic disintegration cell bound 2. Chromatography on CM-Sephadex G-50 3. Gel filtration

CM-cellulose rechromatography ; sedimentation patterns

DEAE-cellulose rechroma tography

41

25,42

Electrophoretic and sedimentation patterns; immunodiff usion Single band on gel electrophoresis

43

Electrophoretic and sedimentation patterns

44

Electrophoretic and sedimentation patterns

44

Electrophoretic pattern and CM-cellulose rechromatography

45

The 8-lactamase gene was acquired by infection with the R factor shown in parentheses. The 8-lactamase gene is chromosomal.

8

30

NATHAN CITRI

of this group of enzymes. However, in most cases no attempts were made t o isolate the enzyme and the available information is too fragmentary to permit characterization a t the molecular level. Table I lists p-lactamase preparations which have been purified and shown to be homogenous. The different purification procedures, summarized in Table I (25,34-45),reflect to some extent differences in the cellular location and in the physical properties of the various preparations. Of particular interest is the isolation of distinct p-lactamases from a single source such as Bacillus cereus strain 569/H, which yields the three P-lactamase preparations listed in Table I (34).This strain is a constitutive mutant of the inducible strain 569 of B. cereus which produces similar levels of all three variants of p-lactamase upon induction (46-49). The relationship between the cell-bound (7-penicillinase) variant and p-lactamase I (a-penicillinase) has been extensively investigated (38, 46, 50-54) but not clarified. Much less is known about the more recently discovered p-lactamase I1 in relation to the two other varieties. One fascinating question which remains open is the interconvertibility of the three varieties (2,.25,37,38). According to a recent report ( 5 5 ) , p-lactamase I of strain 569 can be further resolved by P-cellulose chromatography. The three fractions thus obtained appear to retain their identity on gel electrophoresis, even in the presence of urea, and yield distinctive tryptic digest maps.

36. M. Kogut, M. R . Pollock, and E. J. Tridgell, BJ 62, 391 (1956). 37. S. Kuwabara, and E. P. Abraham, BJ 103, 27C (1967). 38. N. Citri and A. Kalkstein, ABB 121, 720 (1967). 39. M. R. Pollock, A-M. Torriani, and E. J. Tridgell, BJ 62, 387 (1956). 40. M. R. Pollock, BJ 94, 666 (1965). 41. M. H. Richmond, BJ 94, 584 (1965). 42. N. Datta and P. Kontomichalou, Nature 208, 239 (1965). 43. E. B. Lindstrom, H. G. Boman, and B. B. Steele, J. Bucteriol. 101, 218 (1970). 44. S. Yamagishi, K. O’Hara, T. Sawai, and S. Mitsuhashi, J. Biochem. (Tokvo) 66, 11 (1969). 45. T. D. Hennessey and M. H. Richmond, BJ 109, 469 (1968). 46. M. R. Pollock, J . Gen. Microbid 15, 154 (1956). 47. L. D. Sabath and E. P. Abraham, BJ 98, 11C (1966). 48. R. Sheinin, J. Gen. Microbial. 21, 124 (1959). 49. J. D. Duerksen, BBA 87, 123 (1964). 50. J. D. Duerksen and M. L. O’Connor, BBRC 10, 34 (1963). 51. M. R. Pollock and M. Kramer, BJ 70, 665 (1958). 52. N. Citri, BBA 27, 277 (1958). 53. N. Citri, Bull. Res. Council Israel A9, 28 (1960). 54. E. Ron-Zensiper and N. Citri, Nature 198, 887 (1963). 55. J. Imsande, F. D. Gillin, R . J. Tanis, and A. G. Atherly, JBC 245, 2205 (1970).

2.

PENICILLINASE AND OTHER

p-LACTAMASES

31

Several molecular properties of purified P-lactamase preparations are 56-58).I n most cases reported listed in Table I1 (2,25,36,39,40,41,@, the molecular weight is within the range of 28,000-30,000, which seems to be typical for the gram-positive p-lactamases. Similar values have been reported for some but not all, gram-negative p-lactamases. Thus, the values for the enzymes from several strains of E . coli and Salmonella typhinzurium are in the range of 22,00032,000 (43,44, 59,60).In contrast, lower values (14,00C)-17,000) have been reported for other J3-lactamase preparations isolated from E . coli (Table 11, 26, 4 )and from a strain of Enterobacter cloacae (45). The interesting suggestion has been made (61) that the unusually high molecular activity of extracellular p-lactamases (Table 11) may possibly compensate for the dilution of the secreted enzyme in the growth medium. Partial data on the optical rotation (57)and optical rotatory dispersion (55)of native, denatured, and renatured /3-lactamases of B. cereus suggest fairly strong hydrophobic interactions and compact folding. An a-helix content of 30% has been proposed for the native and renatured enzyme of B. cereus 569 (66).

B. COMPOSITION AND SEQUENCE ANALYSES The amino acid composition of the p-lactamases listed in Table I11 (2, 40,41, 43, 45,54, 55, 58, 62-64) underlines the similarities between

preparations derived from closely related strains of bacteria, especially if the combined basic and acidic amino acid content is compared ( 2 3 ) . The staphylococcal enzymes are remarkably rich in lysine, and the lysine-arginine content of the other gram-positive p-lactamases is almost as high. Complete absence of cysteine appears to be characteristic (2,55), although J3-lactamases which contain a t least one cysteine residue have been reported (37,65,66). 56. J. R. Hall and A. G. Ogston, BJ 62, 401 (1956). 57. N. Citri, N. Garber, and M. Sela, JBC 235, 3454 (1960). 58. M. H. Richmond, BJ 88, 452 (1963). 59. C. Lindqvist and K. Nordstrom, J. Bacterial. 101, 232 (1970). 60. H. C. Neu and E. B. Winshell, ABB, 139, 278 (1970). 61. M. R. Pollock, in “The Bacteria” (I. C. Gunsalus and R. Y . Stanier, eds.), Vol. 4,p. 121. Academic Press, New York, 1962. 62. S.Jacobs, Res. Commun. 16th Congr. Quim. Anal., Lisbon, 1966 p . 212 (1956). 62a.M.H.Richmond, personal communication (1966). 63. R. P.Ambler and R. J. Meadway, Nature 222, 24 (1969). 64. R. J. Meadway, BJ 115, 12P (1969). 65. J. T.Smith, Nature 197, 900 (1963). 66. L. D.Sabath and M. Finland, J. Bacterial. 95, 1513 (1968).

TABLE I1 MOLECULAR PROPERTIES OF PURIFIED &LACTAMASES

Source of enzyme"

B. cereus 569 B. cereus 569/H (1) B. cereus 5/B (4) B. lichenifomis 6346 (7) B. Zicheniformis 749/C (6) B. Zicheniformis 749/C (5) S.nureus A ( 8 ) S. aureus B (8) S . aureus C (8) E. wli TEM (9) E. coli G l h l (10)

Molecular weight 31,500 30,800 35,200 28,000 28,000 28,100 29,600 29,600 29,600 16,700

29,OOO

Molecular activityb 1.60 X 1.53 X 1.48 X 2.10 X 1.18 x 1.08 x 2.0 x 2.8 X 1.9 X 2.0 X 2.08 X

106 1W 106 lW 106 106 104 lo3 l(r 10'

1Oa

Sedimentation coefficient (10'8 X ~ 2 0 2.68e 2.5-2.65" 2.82~ 2.63 2.70 2.66 2.62 2.5 2.5 1.85 3.4

EE , ~ )

(nm/mg N) 6.0 6.35 7.35 5.65 5.45 4.75 7.38

Electrophoretic mobility0

(cm*/sec/VX

Asc., 1.68; Desc., 1.70d Asc., 1.73; Desc., 1.65d

Desc., 3.4@ Asc., 4.95; Desc., 4.50/ Asc., 4.33; Desc., 3.65/ -

-

13.1

-

~

Numbers in parentheses are the serial numbers of the preparations in Table I. Moles benzylpenicillin hydrolyzed per mole enzyme per minute at 30". The diffusion coefficients determined (66) for preparations 569, 569/H and 5/B are 8.28, 8.38, and 7.80 (107 x D m J . In glycine buffer, p = 0.2, pH 8.4. e In glycine buffer, p = 0.2, pH 8.15. In veronal buffer, p = 0.1, pH 8.15. u Aw., ascending; l)esc., descending. a

b

Ref.

10-6)

36, 66 36, 66, 67 36, 39, 66 2, 40 2, 40 40 68

41 41 26

43

TABLE I11 AMINOACID COMPOSITION OF B-LACTAMASES E. coli

B. cereus

B. lichenijormis

S. a u r m

E. cloacae Amino acids Lysine Histidine Arginine Aspartic acid Threonine Serine Glutamic Proline Glycine Alanine Valine Methionine Isoleucine Leucine Tyrosine Phenylalanine Cysteine WPhPhan N-term AA Total ammonia amide

214 G l l a l D3lo (Ref. 43) (Ref. 46) 9 2 5

10 8

7 12 9 10 15 10 5 6 10 5 3 ? ?

17 4 10 25 18 16

33 19 23 28 17 4 15 23 10 8 0

-

17 4 9 23 18 14 32 20 22 29 17 5 17 24 12 7 0 6 Ala (22)

5/B

569 569/H (Ref. 2, 62)

31 23 6 6 13 11 33 25 9 12 2 4 25 22 6 10 19 21 30 29 17 15 5 1 22 20 17 19 5 7 7 8 0 0 Aspb Lysc

6346/C (Ref. 40)

749/c (Ref. 63, 64)

21 5 10 31 13 5 22 10 19 33 17 1 22 18 4 6 0

19 1 12 28 18 10 24 9 12 19 10 5 9 21 5

24 1 15 37 21 11 27 11 15 26 15

5

-

0 -

-

LYS

7 3 Lysd (20)

5 14 27 6

TypeA TypeB (Ref. 64,68) (Ref. 4 1 ) 43 2 4 39 13 19 18 9 12 18 16 3 19 22 13 7 0 2 LYS (29)

43 2 5

42 12 17 22 11 16 21 11 2 15 22 9 7 0 -

(28)

TypeC (Ref. 4 1 ) 45 3 4 39 11 17 18 10 15 19 11 2 18 23 14 7 0 -

LYS (32)

The &Iactamase gene was received from G l l a l and both eneymes were purified by identical procedures (Serial No. 11 in Table I). Richmond (@a). c Imsande et al. (66). d See, however, text and Ambler and Meadway (63). a

TABLE IV OF VARIOUSSUBSTRATES (RATEOF HYDROLYSIS OF BENZYLPENICILLIN = 100) RELATIVE RATESOF HYDROLYSIS ~~

w

I@

~

Substrates

Source of enzyme"

Phenoxymethyl- Ampipencillin d i n

Bacillus licheniformis Strain 749/C (3 isozymes) [5] (mutant 77) Strain 6346/C [7] Bacillus cereus Strain 569/H 8-lactamase I [I] plactamase I1 [2] 8-lactamase y [3] Strain 5/B [4] Staphylowccus aureua Types A and B [8] Type C I81 Escherichia w l i Strain TEM (R-TEM) [9] Strain K-12 (Gllal) [lo] Strain W3630 ( h N Z 3 8 ) [I21 Strain W3630 ( h14) ~ [I31 Enlerobacler cloacae 214 [14] a

-

153

64-74

-

-

Cloxacillin

0.45-0.50 4.0 1.0

-

-

-

-

4.6-5.0 5.0 13

0.7 89 <0.1 -

40 9

-

3.5 1.0

10 -

0.5 1.2

5 89 5 4

6-APA

-

100 100

-

1.5 0.6

4.5 1.0

50 -

1 -

-

-

110 4 450

-

-

27 292

-

140

-

-

-

130

86

11

-

-

-

-

171 96

120 64 55

Benzylcephalosporin

Oxacillin

3.0 89 26 1.8

-

Cephalosporin C

Methicillin

-

Numbers in brackets are the serial number of the preparation in Table I.

85 1.2 263

0.3-0.4 0.9.5-1.06 11.0 15 42

17.5-19.8 275 -

Ref.

23 23 40

3 41 17 -

38, 75 37 38 29, 76

-

-

41,58

0.8 920 -

16 -

65 72 3.6

44

-

-

69

44

1830

8200

45

80

7200

<0.1 -

Cephaloridine

-

41 25 43

f

ael

E

2.

PENICILLINASE AND OTHER

p-LACTAMASES

35

The data on terminal amino acids (Table 111) support the accepted notion that every p-lactamase consists of a single polypeptide chain. There are no indications to the contrary, although firm evidence is lacking in most cases. Further observations pertaining to the chemical composition of plactamases and comments on the significance of the reported variations can be found in recent reviews ( d , S , 23). Ambler and Meadway have very recently reported on their studies of the amino acid sequence of p-lactamases from representative strains of S. aureus and B. lichenijormis (63, 6 4 ) . The complete sequence of the p-lactamase protein of S. aureus PCI (type A) was determined by the characterization of the peptides produced by digestion with trypsin, chymotrypsin, or pepsin. The sequence analysis of the p-lactamase protein of B . licheniformis 749/C was performed on both the extracellular and the cell-bound enzyme (cf. serial Nos. 5 and 6, Table I ) . The slight differences between the two forms were confined to the N-terminus and attributed to the difference in the release mechanism. Peptides corresponding to about 90% of the molecule have been characterized and their sequences combined to form five larger fragments. The results of these studies are summarized in Fig. 2, which presents the S. aureus sequence and the B. lichenijormis fragments arranged in such a way as to give the best possible match with that sequence. This arrangement implies that the B . licheniformis molecule is several residues longer a t each end (see also Table 111).The authors estimated that the complete sequence of the B. lichenijormis protein will show about 40% identity with that of S. aureus. This estimate has been most recently confirmed (64), and the homology proposed in Fig. 2 is now firmly established. There can be little doubt that both proteins share a common origin and that their divergent evolution accounts for the differences in the catalytic properties listed in Tables IV and V.

111. Catalytic Properties

A. METHODSOF ASSAY The methods most commonly used for the quantitative assay of plactamase are based on one of the following principles: (1) Manometric or alkalimetric titration of the new carboxyl group formed by hydrolysis of the p-lactam ring; (2) iodometric titration of the product, and (3)

1

8

Lys-

Thr-

26 27 -THR- Lss-Thr-

3

4

5

Glu- Met- Lys28

6 7 Lys- Glu-

Asp- Asp-

8 9 10 11 12 13 14 15 Leu- Asn- Asp- LEV- GLU- Lys- Lys- TyrPhe-

Lys-

LD Leu-

Glu-

Glu-

Glu- Phe-

Ser-

29 Gly-

30 Lys-

Glu-

32 VAL-

Gly-

Thr-

Asn- Arg-

Thr-

Val-

Ah-

Tyr-

Arg-

LC Pro- Asp-

Glu-

51 -Ser-

5’2 Ala-

53 Ile-

54 55 56 Leu- Leu- Glu-

5i Gln-

58 Val-

59 Pro-

60 Tyr-

61 62 Asn- Lys-

63 64 65 Leu- Asn- Lys-

Ser-

Ile

(Glx,

GIx,

Asx.

76 -Ala-

79 80 81 87 83 84 85 86 TYI:- Ser- PRO- ILE- Leu- GLT- LYS- Tyr- VAL- GlyLE Tyr- Asn- Pro- IleThr- Glu- Lys- His- Val- Asp-

87

38 89 Asp- Ile-

31

33 34 35 LD- Phe- Am-

Lys-

8

Ah-

-Val-

??

36

Ser-

id

16 17 18 Asn- ALA- His-

19 He-

20 21 GLT- Val-

Ah-

Leu-

Gly-

Asp-

57 38 39 40 41 42 ASP- Lys- ARG- PHE- ALA- Tyr-

Lys-

Arg-

Asx,

Phe- Ala66 Lys-

L e u ) Arg-

-Tyr-

Jer

126

127 -Oh- Leu-Lys-

Ile-

(Asp.

Asti) Ala-

128

199 130 G L P - ASP- Lys-

106

LCJ- Ile-

112 113 LPS- GluLB Leu- Lys- Gln-

131 13‘7 13) VAL- Thr- Asn-

134 Pro-

135 Val-

136 137 AKG- Tyr-

(Thr.

Gln-

Asn-

Gly-

Asp-

0111- Val

Asx,

Pro,

Glx)

Arg-

Phe-

151 132 153 .Asp- TIIR- Ser-

154 Thr-

155 Pro-

15G Ala-

157 158 ALA- I’he-

159 Gly-

160 Lys-

161 TI r-

162

Srr)

Ala-

Are

Ala-

Val-

Thr-

Scr-

Asx)

Thr

(Tlir,

Leu-

44 45 46 43 ALA-, S E R - T H R - Ser-

Phe

Ah-

Leu- Asp

48 49 47 LTS- ALA- IleAla-

50 Am-

Leu

Ser-

Thr-

Ilc-

Lp-

68 His-

69 Ile-

70 Asn-

71 Lys-

73 74 ASP- ASP- Ile-

75 Val-

Thr-

Tyr-

Thr-

Arg-

Asp-

Asp-

Leu-

9.5 96 Ile- Glu-

07 Ala-

08

Ser-

99 100 Met- Thr-

67 ValLE Ile-

91 92 93 THR-LEU- LTS- Ala90

94 LEr-

7‘7

Asn-

Thr- Gly- Met- Thr- Leu- Lys

107 10rl 109 110 111 ALA- Asn- A S S - L ~ s - I L E - IleAh-

Ile-

23 24 !25 AbA- LEU- ASP-

Phe- Ala-

Ile-

163

Ser!

Leu- Arg.

103 Arg-

124 125 LEV- Lya-

Pro)

Lru-

Glu-

Leu- Arg-

144

145 Ser-

146 Pro-

147 L?s-

148 Stx-

149 Lys-

150 Lys-

(GI?,

Pro,

Thr.

1;1x, Asx,

Asr.

Leu- Ala

119 Val-

(Ser,

140 141 142 143 GLU- LEU- Asn- Tyr-

114 115 116 117 ILE- GLP- GLT- Ile-

138 139 GLU- IleLB Glu- Pro-

Ah,

120 121 122 LJ-s- LTS- Gin-

Glu-

118 Lys-

Lys-

102 103 104 105 -TYR- S E R - Asp- Asn- Thr101

Lys-

Pi! Tyr-

Gly-

Glu-

GI?-

Leu-

1G4 165 166 LEU- A m - Lys- Leu- IleLB Leu- A r e 41a- Phe- A h -

Glu

Tyr-

Gln-

Glu-

Val

16T Ah-

168 Asn-

160

Leu-

Glu-

(Asp.

Lys- Lys-

17‘3

174

GI!--

1TO 171 179 LTS- LEU- Ser-

L?-S.

4:

Asp-

LXS- Leo-

Ser-

Glu-

Pro-

175

1.1.- ASU

Lys-

176 - L p

IT7

178

Lys- Plie-

179 180 LEV- Leu-

181 182 ASP- Leu-

NET- Leu- Asn- Asn- Lys- Ser-

183

Leo-

Asp- Trp-

Met-

184

185

186

187

188

1R9 Gly-

190 Asp-

191 Tlir-

192 193 194 LEV- ILE- Lyr-

195 Asp-

196 197 GLT- Val-

Pro-

193

199 Lys-

200 Asp-

Asn)

Leu- Ile-

Arg-

-4Ia-

Gly

(Asp,

Glu,

Pro.

Trp.

217 .4sn-

919 Val-

220 Ma-

5'11 Phc-

222 Val-

223 224 Tyr- Pro-

225 Lyn-

249

250

LB

Lys- Arg- Asp (Thr. Thr,

Leu-

201 202 -Tyr- Lys-

203 204 Val- Ala-

205 206 207 208 209 ASP- LTS- Ser- CLY- Gln-

Gly,

Ala.

Val,

Val)

Asp-

226

227

228

Gln-

Ser-

229 (;hi-

230 231 Pro- Ile-

-lily.

252 ILE- Ser2.51

-1lew

Ile-

-Are Glu-

Ala-

253 254 GLU- Thr-

Glu-

Ah-

255 Ah-

Thr-

Lys- Thr232 Val-

210 211 ALA- IleLB

Ala-

Gly,

212 213 Thr- Tyr-

214 Ala-

215 216 Ser- Arg-

Thr-

(ily-

Ala-

Ala-

Ber-

Tyr-

Cly-

Arg- Asn- Asp- Ilc-

Aln

233 Leu-

234 Val-

235 Ile-

236 Phe-

237 Thr-

238

239 Asn- Lys-

240 241 ASP- Asn-

242 Lys-

244 Asp-

Ala-

Val-

Leu-

Ser-

Ser-

Arg-

Asp

(Ma,

Lys,

256 257 LYR- Srr-

258 259 260 961 VAL- MET- LYS- Glo-

252

263

264

265

266

967

Phc

Lys-

LA Val-

Leu- Asn-

Met- Asn-

Gly-

Val-

218 Asp-

Met-

Lys- Ala-

243 SerLA Asp,

245 246 LTS- Pro-

L?-s) 1 ~ s - Tyr-

247

248

b s n - A>I'- LYS- LEUAsp-

ASP- Lys-

Leu-

Lys

FIG.2. Amino acid sequences of P-lactamases of Staphylococcus aureus PCI (upper line) and of Bacillw licheniformis 749/C (fragments LA to LE, lower line). Matching residues are shown in block letters (upper line). (From Ambler and Meadway, 63.)

TABLE V APPARENTDISSOCIATION CONSTANT^

(pM)

Substrates Source of enzyme" Bacillus licheniformis Strain 749/C (5) Strain 6346/C (7) Bacillus cereus Strain 569 p-lactamase I* Strain 569/H 8-lactamase I (1) p-lactamase I1 (2) Strain 5/B (4) Staphylococcus aureus Type A (8) Type 80/81b Escherichia coli Strain 214 Tb Strain TEM (9) Strain W3630 ( F ~ 238) N (12) Strain W3630 (&N 14) (13) Aerobacter cloacaeb Pseudomanas pyocyanea

Benzylpenicillin

49 9.5

Ampicillin -

-

Methicillin 0.93 0.23

Oxacillin

Cloxacillin

-

-

b

16.6 16.7

Cephalosporin C

50 50

Cephaloridine

Ref.

-

48

110

410

550

-

-

-

-

60 3300 43

1300 3300

570 1600 330

2300 -

3000 -

-

1100 -

3300

-

460 1600 230

-

77, 79, 80 37 20, 77

5 12

170

16000 5000

5000 -

-

120

100 -

-

41, 58, 81

22 5.1 27.8 8 13

35oc 2.3 15.9 34.0 541 -

>46& 22 26.3 222

-

-

-

1000

600 111 400 69 130

44= 3.2

-

-

-

0.lP

-

1.lC

0.g 13.0 16.5c 0.026

Numbers in parentheses are the serial numbers of the preparations in Table I. For purification procedure see original references. c Based on KI with benzylpenicillin as substrate. a

6-APA

-

-

46, 77, 78

78 82, 83

25

44 44 83 31

2.

PENICILLINASE AND OTHER

P-LACTAMASES

39

colorimetric determination of the hydroxaniate formed by residual substrate. The various applications of these and several other principles have been reviewed elsewhere (67, 6‘8).The assay methods, primarily designed for benzylpenicillin, the standard substrate, are generally suitable for other substrates, although in some cases the procedure has to be modified. Specific problems involved in the replacement of benzylpenicillin with other substrates, especially cephalosporins (69) have been recently discussed ( 2 , 31, 32, 7 0 ) . )

B. KINETICSAND SUBSTRATE SPECIFICITY

It is generally agreed that the &I-lactamase reaction is of zero order in the presence of saturating substrate concentration (71) and that the enzyme displays typical Michaelis-Menten kinetics over a wide rangc of concentrations (72). The slight deviations, which have been observed with crude enzyme preparations only (59, 73) deserve further investigation ( 4 ) . Anomalously high dissociation constants recorded with certain substrate combinations (74) have been correlated with the effect of such substrates on the conformation of p-lactamase (see Section IV,B) . Individual “substrate profiles” of variously related p-lactamases and apparent dissociation constants for several substrates are listed in Tables IV (2.9)25, 29,37, 38,40, 41, 43-45, 58, 75, 76) and V (20, 25, 31, 37, 40) 41, &,46,58,76,77--83), respectively. An interesting attempt to correlate )

67. J. M. T. Hamilton-Miller, J. T. Smith, and R. Knox, J. Pharm. Pharmacol. 15,81 (1963). 68. N. Citri, Methods M e d . Res. 10, 221 (1964). 69. E. P. Abraham, Phamtacol. R e v . 14, 473 (1962). 70. M. R. Pollock, J. Fleming, and S. Petrie, Proc. 2nd Meeting Federation Euro-

pean Biochem. SOC.,1966 p. 139. Pergamon Press, Oxford, 1967. 71. R. J. Henry and R. D. Housewright, JBC 167, 559 (1947). 72. J. E. Banfield, Experientia 13, 403 (1957). 73. W.Rothe, Pharmazie 5, 25 (1950). 74. N. Zyk and N. Citri, BBA 151, 306 (1968). 75. N. Citri and N. Zyk, BBA 99, 427 (1965). 76. S. M. Chaikovskaya and T. G. Venkina, Antibiotiki 9, 329 (1964). 77. N. Citri, N. Garber, and A. Kalkstein, BBA 92, 572 (1964). 78. R. H. Depue, A. G. Moat, and A. Bondi, ABB 107, 347 (1964). 79. F. R. Batchelor, J. Cameron-Wood, E. B. Chain, and G. N. Rolinson, Proc. R o y . SOC.B154, 514 (1961). 80. S. Kuwabara and E. P. Abraham, BJ 115, 859 (1969). 81. M. H. Richmond, Brit. M e d . Bull. 21, 260 (1965). 82. J. M. T. Hamilton-Miller and J. T. Smith, Nature 201, 999 (1964). 83. J. M. T. Hamilton-Miller, J. T. Smith, and R.Knox, Nature 208, 235 (1965).

40

NATHAN CITRI

these values in a physiologically meaningful way has been made by Pollock (40), who pointed out that under typical ecological conditions the enzyme functions at unsaturating substrate concentrations. Thus the “physiological efficiency” of a p-lactamase depends on both the V,,,,, and the K , value and may be conveniently expressed as the ratio VlnaJKm. There can be little doubt that this ratio is an important parameter in comparing 8-lactamases in any context dealing with the ecology, evolution, and physiological significance of the enzyme (2,25,40,84).

C. STRUCTURAL MODIFICATIONS IN SUBSTRATES The effect of replacement of the APA nucleus with an ACA nucleus on the susceptibility of the substrate to enzymic hydrolysis depends on the nature of the p-lactamase ( 2 ) . It has been richly illustrated in the case of the p-lactamase of Pseudomonas pyocganea (31)and analyzed in terms of conformative response (see Section IV,B) in a study of a B . cereus p-lactamase (75).I n that study the role of the nucleus was examined in both catalytic and noncatalytic enzyme-substrate interactions. Little has been published on the effect of modifications within each nucleus, presumably because such modifications lead, as a rule, to the loss of antibiotic activity. Conversion of the carboxyl groups in penicillins confers partial resistance to p-lactamase (85-87).I n cephalosporins, replacement of the acetyl group in position 3 of the dihydrothiazine ring [ R in Fig. 1, (111)] by pyridine causes increased susceptibility to hydrolysis by p-lactamases of Pseudomonas pyocyanea (31,88),Enterobacter cloacae (45),E . coli (42,88),and both the extracellular and cell-bound enzymes of B . cereus (38)* The most common modification is the replacement of the N-acyl substituent (“side chain” R in Fig. 1 ) , especially in penicillins. Indeed, the differences in K , and relative V,,, values among the penicillins listed in Tables IV and V all result from the differences in the nature of the side chain. Analysis of such data (Table VI) (25,@, 4,70,76,77-79, 81, 89,901 84. M. R. Pollock, Antimicrobial Agents Chemotherapy p. 292 (1965). 85. D. E. Cooper and S.B. Brinkely, JACS 70, 3966 (1948). 86. N . J. Huang, T. A. Seto, J. M. Weaver, A. R. English, T. J. McBride, and G. M. Schull, Antimicrobial Agents Chemotherapy p. 493 (1964). 87. E. Grunberg and G . Beskid, Antimicrobial Agents Chemotherapy p. 619 (1968). 88. L. D. Ssbath and M. Finland, Ann. N. Y. Acad. Sci. 145, 237 (1967). 89. R. P. Novick, BJ 83, 229 (1962). 90. J. M. T. Hamilton-Miller, BJ 87, 209 (1963).

2.

41

PENICILLINASE AND OTHEB ~-LACTAMASES

TABLE VI EFFECTOF SIDE CHAINON CATALYTIC CONSTANTS OF PENICILLINS Side chain Phenylacetyl-

V,..:: V,..P 5.3 16.7 2.6 20.0 10.0

Phenylglycyl-

Dimethoxybenzoyl-

K,r: K,P 0.003 0.10 0.02 3.0 20.8

7.7

0.57

1.37

1. o

90.0 0.38 0.77 1.3 1.5 25.0 3.6 1.71 1.07 0.08

0.17 0.67

0.014 0.19 0.13 0.105 0.054 1.40 0.007 0.61 0.15 0.15 37.2 41.5

Source of enzyme

Ref.

S . aureua 524 SC S . aureua 147 B. cereua 569/H B. lichenifomis 749/C (wild type) B. lichenifomis 749/C (mutant 19) B. lichenifomis 6346/C (wild type) B. l i c h e n i f m i s 6346/C (mutant 3) E. coli G l l a l E. wli GN 238 E. coli GN 14 E . euli TEM K . aerogenes 418 S . aurew 147 B. coli G l l a l E . coli GN 238 E. coli GN 14 B. cereua 569/H S . auras 524 SC 8.aureus 147

89 78 79 70

70

70

ro 43

44 44 86

90 81

43

44 44

76, 77, m 89 78

~~

V,..I and K,I are the respective constants for the unsubstituted 6-APA; V,,, K,t are the corresponding constants for the indicated 6-APA derivatives. 0

and

brings out the striking differences in the response of the various p-lactamases to a given side chain. Such differences are clearly evident even in closely related enzyme preparations and in some instances (e.g., the mutants of B . Zicheniformis) may result from the replacement of a single amino acid. The role of the side chain in both catalytic and noncatalytic (76) enzyme-substrate interactions has been investigated in considerable detail (reviewed in reference a ) , and a mechanism has been suggested which may explain the rate-controlling effect of side chains on the catalytic activity of p-lactamase (see Section IV,B).

D. STRUCTURAL MODIFICATIONS IN

THE

ENZYME

No data are available on the effect of defined structural modifications on the properties of p-lactamases. Indirect evidence bearing on the

42

NATHAN CITRI

role of the primary structure in catalytic activity comes from the studies of Pollock (23) on the effect of mutations on the properties of the plactamase of B. licheniformis 749/C. It appears that structural mutations, presumably involving a single amino acid replacement in the enzyme molecule, frequently cause a considerable decrease in thermostability (70) along with a decrease in the catalytic activity ( 2 3 ) . The substrate profile is altered as in the case of the mutant strain 749/C/77 (Table IV) with preferential loss of “penicillinase” activity. The change in thermostability and in response to antibodies (see also Section V) implies an altered conformation in such mutant enzymes (23,7 0 ) . A comparison of spontaneously and experimentally released p-lactamase preparations of B . lichenifomis (40) indicates that the ends of the polypeptide chain can vary without affecting the properties of the enzyme. Although such preparations differ in the C- and N-terminus, they are serologically and catalytically indistinguishable (40,63). Similarly, B. lichenifomis 749/C isozymes, separable by starch gel electrophoresis or DEAEchromatography and believed to differ a t the C- or N-terminus C40),have identical substrate specificities (Table V) . Other indirect (genotypic or phenotypic) modifications of the primary structure (70, 91, 92) as well as reversible conformational transitions (52, 67, 93) have been reviewed by Citri and Pollock ( 2 ) and more recently by Pollock ( 4 0 ) .

E. OTHERFACTORS AFFECTING ACTIVITY 1. Effect of pH and Temperature

Typical pH-activity curves obtained with gram-positive p-lactamases and benzylpenicillin show maxima in the range of p H 6.0-7.0 with a rather sharp decline in the alkaline range ( 2 ) . Recent data on gramnegative p-lactamases (25,31,43,44,59,65,90,94-96)show optimal values scattered over the range of pH 5.0-8.5. A shift in the pH-activity curve has been observed when benzylpenicillin was replaced with 6-APA (78,79) or methicillin (54) but not with cephalosporin C ( 3 1 ) . Few data are available on the temperature dependence of p-lactamase activity ( 2 ) . With benzylpenicillin as the substrate, the optimal tem91. 92. 93. 94. 95. 96.

D. A. Dubnau and M. R. Pollock, J . Gen. Microbiol. 41, 7 (1965). M. H. Richmond, BJ 77, 112 (1960). N. Citri and N. Garber, BBA 30, 664 (1958). J. T. Smith and J. M. T. Hamilton-Miller, Nature 197, 976 (1963). J. M. T. Hamilton-Miller, BBRC 13, 43 (1963). G. A . J. Ayliffe, J. Gen. Microbiol. 30, 339 (1!963).

2.

PENICILLINASE AND OTHER

p-LACTAMASES

43

peratures reported range from 30" (44,97)through 35'40" for B. cereus p-lactamases (71,98)to 45"-55" for several enzyme preparations (&,89,98,99).Optimal temperatures for other substrates have not been reported. It has been pointed out (100) that the activation energy of the catalytic reaction (with benzylpenicillin) is generally higher in the gram-positive p-lactamases (7.2-8.8 x 103 cal/mole) (71,100,101) than in gramnegative enzymes (3.3-5.2 X lo3cal/mole) (31,94,95,100). Replacement of benzylpenicillin with cephalosporin C (31) caused considerable increase in the activation energy. 2. Activators and Inhibitors

As a rule p-lactamase activity does not depend on specific activators or cofactors. The only exception so far is p-lactamase I1 which requires Znz+for stability and activity (37).The metal binding characteristics of this enzyme have been described in a recent study (66).In other cases where the ion content of the assay medium may affect the activity (@,44), no absolute requirements were demonstrated. The search for selective means of inactivation, prompted by clinical considerations, has not been successful, although many substrate analogs act as powerful competitive inhibitors ( 2 ) . On the other hand, irreversible inhibition may result from interaction with substrate analogs which oause labilization of the enzyme (see Section IV,B) . Specific stimulation and inhibition by homologous antibodies is reviewed in Section V. Inhibition of a staphylococcal p-lactamase by certain dipeptides has but it is not clear whether the active site was directly been reported (102), involved. Other remotely related and unrelated compounds as well as metals have been implicated, but the results are largely inconclusive (8,9,20,&). Of the nonspecific inhibitors, the thiol reagents have been most extensively tested and usually found ineffective. This is not surprising in view of the available data on the amino acid composition of p-lactamases (Table 111) which show total absence of cysteine. However, interesting exceptions have been reported. Thiol reagents inhibit the Zn2+-dependent p-lactamase I1 of B . cereus (37, 66) and the &lac97. M. Goldner and R. J. Wilson, Can. J. Microbiol. 7, 45 (1981). 98. E. E. D. Manson, M. R. Pollock, and E. J. Tridgell, J. Gen. Microbiol. 11, 493 (1954). 99. A. Royce, C. Bowler, and G. Sykes, J. Pharm. Pharmacol. 4, 904 (1952). 100. J. T. Smith and J. M . T. Hamilton-Miller, Nature 197, 769 (1963). 101. N. Zyk and N. Citri, BBA 159, 317 (1968). 102. A. K. Saz, D. L. Lowry, and L. J. Jackson, J . Bacten'ol. 82, 298 (1961).

44

NATHAN CITRI

tamases of K . aerogenes and of A . cloacae (65,lOS).I n the case of /3-lactamase I of B. cereus, where the absence of cysteine is reportedly established (Table 111) (lO4),inhibition by PCMB (75)and by Hgz+ ( K , = 1.4 X lo-' M ) (105)has been demonstrated. The former depends on conformative response to substrate analogs (75).The inhibition by Hg2+, which is accompanied by a slight shift in the CD signal in the region of 205-240 nm, is partly reversed by EDTA (105). Inhibition by chelating agents has been reported only in the case of the Zn2+-dependentp-lactamase I1 (47,106). Alcohols may inhibit (107), or stimulate (102).The inhibitory effect of alkyl sulfates (108,109)depends on the length of the alkyl chain (109,110). Thermal inactivation has been studied mainly in the B . cereus enzymes (S8,98,111,llb). The protective effect of certain macromolecules (e.g., gelatin) is observed at loo", but paradoxically not a t 70" (113,114). The effect of various substrates on the thermostability of several has been studied in the context of conp-lactamases (38,77,112,115) formative response (see Section IV,B) .

IV. Conformation and Function

A. NONSPECIFIC CONFORMATIONAL TRANSITIONS The absence of disulfide bridges appears to be one of the few characteristics common to all p-lactamases. There are indications that in one case a t least (the cell-bound 7-type p-lactamase of B. cereus) the tertiary 103. J. T. Smith, BJ 87, 40P (1963). 104. M. R. Pollock and M. H. Richmond, Nature 194, 446 (1962). 105. L. D'Souza and R. A. Day, Abstr., 168th A m , Chem. SOC.Meeting, New York p. 60 (1969). 106. L. D. Sabath and E. P. Abraham, Antimicrobial Agents Chemotherapy p. 392 (1966). 107. E. B. McQuarrie and A. J. Lubmann, A B B 5, 307 (1944). 108. R. Brunner, A. Kraushaar, and E. Prohaska, Antibiot. Ann. p. 169 (1960). 109. Z. C . Kaminski, J. Bacteriol. 85, 1182 (1963). 110. E. Rauenbusch, M e d . Chem., Abhandl. Med.-Chem. Forschungsataetterr Farbenfabriken Bayer A . G . 7 , 466 (1963). 111. E. E. D. Manson and M. R . Pollock, J. Gen. Macrobiol. 8, 163 (1953). 112. N. Citri and N. Garber, BBA 67, 64 (1963). 113. D. H. Williams, 111, A. Bondi, A. G . Moat, and F. Ahmad, J. Bacteriol. 91, 257 (1966). 114. For an interpretation of this and related observations, the original report (11.9) should be consulted.

115. N. Zyk and N. Citri, BBA 159, 327 (1968).

2.

PENICILLINASE AND OTHER

P-LACTAMASES

45

structure of the enzyme is nevertheless stabilized by firm, possibly coI n more typical cases, valent, attachment to the cell membrane (38,48). and certainly in all extracellular P-lactamases analyzed so far, the conformation of the enzyme is not constrained by covalent bonds. A likely consequence is the marked flexibility of the conformation of the extracellular p-lactamases of several bacilli (67,77). Reversible transitions in the conformation (and antigenic structure) have been demonstrated in extracellular p-lactamases of B. cereus 569/H (69,67,93) and of B. cereus (52).Thus, a mild treatment with alkali, urea, or guanidine .HCl resulted in a markedly increased levorotation and in exposure of iodine-sensitive regions. Similar changes, including almost total loss of antigenic identity, were observed when the enzyme was adsorbed to charged surfaces. Surprisingly, the adsorbed enzyme was fully active (68)and this led to an inquiry into the possibility that the substrate may restore the conformation of the active site (93,116).

B. SPECIFIC TRANSITIONS : CONFORMATIVE RESPONSE I n a series of papers on the interaction of several purified p-lactamase preparations with various substrates (75,77,101,118,116,117-120), evidence was presented which indicates a possibly meaningful correlation between the structure of the substrate, its effect on the conformation of the enzyme, and the catalytic activity. Much of the evidence (partly reviewed in reference 8) is based on interactions with penicillins differing in the structure of the side chain (substituent R in Fig. 1). As pointed out above (Section 111) the side chain controls the rate of the catalytic reaction. It was subsequently observed that it also controls the conformative response, i.e., the effect of the substrate on the conformation of the enzyme (76).Significantly, the nature of the conformative response could be correlated with the effect on the catalytic activity. Thus, the conformative response to benzylpenicillin and other penicillins with side chains promoting the rate of catalysis is characterized by increased stability of the enzyme molecule. Conversely, penicillins which carry side chains interfering with the catalytic activity labilize the enzyme and facilitate inactivation by heat, urea, proteolysis, iodination, or photooxidation. Such penicillins act as competitive inhibi116. 117. 118. 119. 120.

N. Citri and N. Garber, BBA 38, 50 (1960). N. Garber and N. Citri, BBA 62, 385 (1962). N. Citri and N. Garber, BBRC 4, 143 (1961). N. Citri and N. Garber, J . Pharm. Pharmctcol. 14, 784 (1962) N. Zyk and N. Citri, BBA 146, 219 (1967).

46

NATHAN CITRI

tors of the hydrolysis of benzylpenicillin. The competitive relationship at the catalytic level is reflected in a corresponding shift in the conformative response. These observations led to the conclusion that the side chain of penicillin affects the rate of the reaction by modifying the conformative response of the enzyme. Subsequent studies led to the suggestion that the modified conformative response is strongly reflected in the binding properties of the enzyme (120).Although lacking direct support, this suggestion provides a single explanation for two sets of discrepancies observed when a comparison is made of enzyme-substrate dissociation constants based on the catalytic activity with those based on the conformative response ( 7 4 ) .

V. Immunological Studies

The immunology of p-lactamases has been studied fairly extensively (16,23,41,4.3,70,115,121-124) and reviewed elsewhere (70,125).At the molecular level, differences in antigenicity could be usually correlated with differences in composition and physical properties ( 7 0 ) . On the other hand, reversible changes in antigenicity-without apparent loss of activity-could be correlated with reversible conformational transitions ( 5 2 ) . The effect of specific antibodies on the catalytic activity of p-lactamase is usually inhibitory, as expected (125).I n some cases, however, it may be stimulatory ( 7 0 , 1 2 2 ) .This surprising effect depends on the structure of the substrate (101, 122) or on structural mutations in the enzyme ( 7 0 ) . It has been suggested (70,101,115) that the stimulation results from the effect of the antibody on the conformation of the enzyme. Indeed, the effect of antibodies on the conformation of p-lactamase shows a quantitative correlation with their effect on the catalytic activity (101). Specifically, homologous antibodies suppress the conformative response and virtually eliminate the effect of the side chain of the substrate on the rate of catalytic activity (116). These results (cf. Section IV,B) imply a plausible mechanism (115) which may account for both the stimulating and inhibitory effect of antibodies on the activity of p-lactamase. 121. 122. 123. 124. 125.

M. R. Pollock, J. Gen. Microbial. 14, 90 (1956). M. R. Pollock, Immunology 7, 707 (1964). H. K. Rhodes, M. Goldner, and R. J. Wilson, Can. J. Microbiol. 7 , 355 (1961). N. Citri and G. Strejan. Nnture 190, 1010 (1961). M. R. Pollock, Ann. N . Y . Acad. Sci. 103, 989 (1963).

Purine. Purine Nucleoside. and Purine Nucleotide Aminohydrolases C . L. ZIELKE

C. H . SUELTER

I . Introduction . . . . . . . . . . A . Historical Background . . . . . . B . Distribution . . . . . . . . . C . Methods of Assay . . . . . . . 11. Adenine Aminohydrolase . . . . . . . I11. Adenosine Aminohydrolase . . . . . . . A . Molecular Properties . . . . . . . B . Catalytic Properties . . . . . . . C . Considerations of Physiological Function . . IV . 5'-Adenylic Acid Aminohydrolase . . . . . A. Molecular Characteristics . . . . . . B . Catalytic Properties . . . . . . . V . Adenine Nucleoside and Nucleotide Aminohydrolast. (Nonspecific) . . . . . . . . . A . Adenosine Aminohydrolase (Nonspecific) . . B . Adenine Nucleotide Aminohydrolase . . . VI . Guanine Aminohydrolase . . . . . . . VII . Guanosine Aminohydrolase . . . . . . .

. . . .

. . . . . . .

.

.

.

.

. . . . . . .

. . . . . . .

. . . . . . .

.

.

.

.

.

.

.

.

.

.

.

.

.

.

.

47 48 49 51 51

54 54 56

63 64 64 66

73 73 75 76 77

.

I Introduction

The purine bases. adenine and guanine. participate in nature by providing structural elements of nucleic acids and numerous cofactors . Tlie deaminated products of adenine and guanine do not participate i n these coenzymic functions except in a relatively few instances . For example. the relative ineffectiveness of inosinc monophosphate (IMP) vs . 47

48

C. L. ZIELKE AND C. H. SUELTER

adenosine monophosphate (AMP) a3 an effeclor of teveral enzymes such as muscle phosphorylase ( I , d ) , heart phosphofructokinase (S),fructose-1,6-diphosphatase ( 4 ) ,and threonine dehydraTe ( 5 ) has been widely documented. Inosine is found in certain restricted positions of several tRNA ( 6 , 7 ) . Other aspects of guanine and adenine nucleotide distribution and their involvement as substrates and regulators of many enzymecatalyzed reactions are discussed elsewhere (8-10).

A. HISTORICAL BACKGROUND The existence of two separate enzymes in animal tissues responsible for the liberation of ammonia from each of the two aminopurines, adenine and guanine, the latter specific for the free purine and the former for the nucleosides, was initially presented by .Jones and his colleagues (11, 12). I n 1928, Schmidt (13-15) demonstrated that AMP aminohydrolase was responsible for the appearance of inosinic acid in muscle and for a t least a portion of ammonia liberated during contraction. He showed not only a marked specificity for deamination of 5’-AMP but also provided the first clue that muscle adenylic acid (5’-AMP) and yeast adenylic acid (3’-AMP) were different compounds. Initial evidence for guanine and adenosine aminohydrolase including aspects of the specificity were also described by Schmidt (16‘). Additional details regarding development of interest in purine aminohydrolases are available in several excellent reviews (17-20). E. G. Krebs, BBA 15, 508 (1954). N. B. Madsen and C. F. Cori, BBA 15, 516 (1954). T. E. Mansour, JBC 238, 2285 (1963). K. Taketa and B. M. Pogell, BBRC 12, 229 (1963). K . W. Rabinowitz, J. D. Shada, and W. A. Wood, JBC 243, 3214 (1968). R. W. Holley, J . Apgar, G. A. Everett, J. T. Madison, M. Marquisee, S. H. Merrill, J. H . Penswick, and A. Zarnir, Science 147, 1462 (1965). 7. J. T. Madison, Ann. R e v . Bwchem. 37, 131 (1968). 8. M. F. Utter, “The Enzymes,” 2nd ed., Vol. 2, Part A, p. 75, 1960. 9. D. E. Atkinson, Ann. R e v . Biochem. 35, 85 (1966). 10. R. M. Bock, “The Enzymes,” 2nd ed., Vol. 2, Part A, p. 3, 1960. 11. W. Jones and C. R. Austrian, Z . Physwl. Chem. 48, 110 (1906). 12. S . Amberg and W. Jones, Z . Physiol. Chem. 73, 407 (1911). 13. G. Schmidt, Z. Physiol. Chefm. 179, 243 (1928). 14. G. Schmidt, Z. Physwl. Chem. 179, 261 (1928). 15. G. Schmidt, in “Biological Phosphorylation” (H. M. Kalckar, cd.), p. 45. Prentice-Hall, Englewood Cliffs, New Jersey, 1969 [English translation of excerpts of Schmidt (IS,1411. 16. G. Schmidt, 2. Physwl. Chem. 208, 185 (1932). 17. P. P. Cohen and G. W. Brown, Jr.. Comp. Biochem. 2, 177 (1960). 1. 2. 3. 4. 5. 6.

3.

PURINE AMINOHYDROLASES

49

B. DISTRIBUTION The virtual lack of adenine aminohydrolase in animal tissues has been confirmed in several laboratories (81-25). The reported presence of this enzyme in milk (21) has not been confirmed (26). Evidence for adenine aminohydrolase i n Saccharom y c e s rerevisine and Candida utilis based on enhanced growth on adenine (27) has been supported by Rousch and his colleagues (28, 29). The direct deamination of adenine by extracts of E . coli (30-32) has not been verified (33). Adenosine aminohydrolase occurs in tissues of both vertebrates and invertebrates. The enzyme has been observed i n the larvae of Drosophila melanogaster ( S 4 ) , the blowfly (35), sea urchin eggs (36),the hapatopancreas of both crayfish and lobster (3‘7), and a variety of animal tissues (38-4lb). Brady and O’Donovan (4%’)examined the distribution 18. M. P. Schulman, in “Metabolic Pathways” (D. M. Greenberg, ed.), Vol. 11. p. 418. .4cademic Press. Xew York, 1961. 19. Q . Schmidt, in “The Nucleic Acids” (E. Chargaff and J. N. Davidson. cds.). Vol. 1. 11. 595. Academic Press, New York, 1955. 20. T.-P. Lee, “The Enzymes,” 2nd ed., Vol. 4, p. 279, 1960; A. Raggi, S. RoncuTestoni. and G. Ronca, BBA 178, 619 (1969). 21. E. J. Morgan, C. P. Stewart, and F. G. Hopkins, Proc. Roy. SOC.B94, 109 (1922-1923). 22. Q. Duchateau, M. Florkin, and G. Frappez, C o m p t . Rend. SOC.B i d . 133, 274 (1940). 23. M. B. Blauch, F. C. Koch, and M. E. Hanke, JBC 130,471 (1939). 24. J. P. Greenstein, Federation Proc. 6, 488 (1947). 25. Y. Wakabayasi, J . Biochem. (Tokyo) 29, 247 (1939). 26. W. F. Williams and C. W. Turner, Proc. SOC.Exptl. Biol. M e d . 94, 196 (1957). 27. F. J. DiCarlo, A. S. Schultz, and D. K.McManus. JBC 189, 151 (1951). 28. A. H. Rousch, A B B 50, 510 (1954). 29. A . H. Rouscli and M. A. Saeed, BBRC 2, 43 (1960). 30. C. Lutwak-Mmn, BJ 30, 1405 (1936). 31. M. Stephenson and A. R. Trim, BJ 32, 1740 (1938). 32. S. Friedman and J. S. Cots, JBC 201, 125 (1953). 33. A . L. Koch and G. Vallee, JBC 234, 1213 (1959). 34. I<. P. Wagner and H. K. Mitchell, A B 17, 87 (1948). 35. F. G. T,cnnox, Australian Council Sci. I d . Res.. Pnmphl. 109, 37 (1941). 36. T. Gustafson and I. Hasselberg. E z p t l . Cell Res. 2, 642 (1951). 37. A. H. Rousch and R. F. Betz, BBA 19, 579 (1956). 38.E. J . Conway and R. Cooke, BJ 33, 479 (1939). 39. D. A . Clark. J. Dnvoll, F. S. Philips, and G. B. Brown, J. Pharmacol. Exptl. ‘1’Aemp. 106, 291 (1952). 40. J . Purzycka, Acta Biochim. Polow. 9, 83 (1962). 41. W. Mak:ircwicz and M. Zydowo. C o n t p . Biocliem. Pliysiol. 6, 269 (1962). 41a. S. J . Mustafn and C. P. Tewari, Eiizymologiu 38, 177 (1970). 41b. S. J. Mustafa and C. P. Tewari, BBA 198, 93 (1970). 42. T. G. Brady and C. I. O’Donovan, P o m p . Biochem. Physwl. 14, 101 (1965).

50

C. L. ZIELKE AND C. H. SUELTER

of this enzyme in 20-23 tissues of rat, guinea pig, mouse, cat, rabbit, and dog, and 10 tissues of the calf. Activity in heart and skeletal muscle was generally low; activity was always high in the spleen. Although activity in the small intestine was consistently high, in other parts of the alimentary tract i t was generally low. Of all tissues examined, the cat placenta, calf intestinal mucosa, cat lung, and cat spleen exhibited the highest activity. Unequivocal proof for existence of adenosine aminohydrolase in any type of higher plant is still lacking (43). AMP aminohydrolase, an enzyme relatively specific for AMP, has been observed in reptiles ( 4 4 ) ,erythrocytes (%), snail (45), unfertilized fish eggs (46), invertebrates (47), a variety of mammalian tissues ( 2 0 ) , and a particulate fraction of pea seeds (48).Evidence suggests that the frog muscle AMP aminohydrolase is located within or just beneath the sarcolemma (49). The rabbit skeletal and heart muscle enzymes were found in the cytoplasm and mitochondria (20, 40, 50, 51), while the enzyme of kidneys and gills of freshwater fish was located in the cytoplasmic fraction (6.2).The enzyme occurs in most areas of the rat (53) and rabbit brain (54). The nonspecific enzyme from several microbial sources deaminates adenosine triphosphate ( ATP) and adenosine diphosphate (ADP) as well as AMP (see Section V ) . Guanine aminohydrolase is present in a variety of animal tissues (55-58), in lobster hepatopancreas (37), in certain bacteria (59), and 43. T. G . Brady :ind V. J. Hrgurty, Nature 209, 1027 (1966). 44. J. 13. Balinsky. G. I,. Murison, A. R. Conradie, and R. E. Santo. S. African J. Med. Sci. 29, 61 (1964). 45. T. Fujiwara and 13. Spencer, BJ 85, 19P (1962). 46. N. M. Abrosimova and R. I. Tatarskaya, Biokhimiya 28, 128 (1963); C A 58, 12895b (1963). 47. J. Uniiastowski. Acta Biochim. Polon. 11, 459 (1964). 48. D. H. Turner and J. F. Turner, BJ 79, 143 (1961). 49. C. R. Dunkley, J. F. Manery, and E. E. Dryden, J. Cellular Physiol. 68, 241 (1966). 50. 1). C. Park and R. J. Pennington, Clin. Chim. Acta 13, 694 (1966). 51. Z. Yu. Nechiporenko and 0. P. Goloborod’ko, Ukr. Biokhim. Zh. 39, 322 (1967) ; C A 67, 105156s (1967). 52. W. Makarewicz, Acta Biochim. Polon. 10, 363 (1963). 53. H. Kluge and V. Wieczorck, Acla Bwl. Metl. Gel. 21, 271 (1968); C A 70, 26875j (1969). 54. M. K . Malyslievu and N. M. Polyukova, U k r . Biokhim. Z h . 37, 360 (1965); C A 63, 12063b (1965). 55. J. P. Greenstcin, C. E. Curter. H . W. Chalkley, and F. M. Leuthardt, J. Natl. Cancer Inst. 7, 9 (1946). 56. W. D. Block and D. V. Johnson, JBC 217, 43 (1955). 57. T. C. Hall, R. Levine, and C. A. Harris, Biochem. Pharmacol. 8, 71 (1961). 58. J. E. Roy, Can. J. Biochem. 44, 1093 (1966). 59. J . Rakosky, Jr. and J. V. Beck, J. Bacterial. 69, 563 (1955).

3.

PURINE AMINOHYDROLASES

51

in yeast Torula utilis (28). The enzyme from rat kidney or spleen but not the brain and liver was localized almost exclusively in the 15,000-g supernatant fraction of isoosmotic sucrose homogenates. The rat cerebellum was devoid of guanine aminohydrolase activity. However, the enzyme from cerebral hemispheres appeared in two isozymic forms: one present a t birth was associated with the 5000-g supernatant while the second, located in the nuclear and mitochondria1 fractions, did not occur until 15-20 days after birth (60).

C. METHODSOF ASSAY Purine aminohydrolases may be assayed by measuring release of ammonia (IS, 14) or directly by the much more convenient spectrophotometric method developed by Kalckar ( 6 1 ) and Rousch and Norris (62).Absorbancy changes resulting from enzymic hydrolysis of the purines or purine derivatives are summarized in Table I (62-75).

II. Adenine Aminohydrolare

The partially purified adenine aminohydrolase (EC 3.5.4.2) from Azotobacter vinelandii catalyzes the anaerobic conversion of adenine to 60. S. Kurnar, V. Josan, K. C. S. Sanger, K. K. Tewari, and P. S. Krishnan, BJ 102, 691 (1967); S. Kumar, ABB 130, 693 (1969). 61. H. M. Kalckar, JBC 167, 445 and 461 (1947). 62. A. H. Rousch and E. R. Norris, AB 29, 124 (1950). 63. Y. Ishida, H. Shirafuji, M. Kida, and M. Yoneda, Agr. Biol. Chem. 33, 384 (1969). M. R. Wolfenden, JACS 88, 3157 (1966). 65. L. N. Simon, R. J. Bauer, R. L. Tolman, and R. K. Robins, Biochemistry 9, 573 (1970). 65a. H. P. Baer, G . I. Drummond, and J. Gillis. ABB 123, 172 (1968). 66. A. Coddington, BBA 99, 442 (1965). 67. R. Wolfenden, J. Kaufman, and J. B. Macon. Biochemistry 8, 2412 (1969). 68. K. L. Smiley, Jr., A. J. Berry, and C. H. Sueltrr, JBC 242, 2502 (1967). 69. R. Wolfenden, T. K. Sharpless, I. S. Ragade, and N. J. Leonard, JACS 88, 185 (1966). 70. B. M. Chassy and R. J. Suhadolnik, JBC 242, 3655 (1967). 71. S. Frederiksen, ABB 113, 383 (1966). 72. J. G. Cory and R. J. Suhadolnik, Bioche'mistry 4, 1733 (1965). 73. S. F. Mason, Ciba Found. Symp. Chem. Biol. Pteridines p. 74 (1954). 74. S. Frederiksen, BBA 87, 574 (1964). 75. S. Minato, T. Tagnwa. M. Miyaki, B. Shimiau, and K. Nakanishi, J . Biochem. (Tokyo) 59, 265 (1966).

52

C. L. Z I E L R E AND C. H. SUELTER

TABLE I ABSORBANCYCHANGES RESULTING FROM ENZYMIC HYDROLYSIS OF ~-SUBSTITLIENT OF PURINE DERIVATIVES

Ae X 1W'M PH A

Substrate Adenine Adenosine N1-Methyladenosine 2'-Deoxyadenosine

AMP 3-(8-~-Ribofuranosyl) adenine 8-hinoadenosine 8-Azaadenosine 8-Hydroxyadenosine 2,6-Diaminopurine ribonucleoside

(nm)

6

7

265 260 265 265 260 265 265 265 285 290 285

-7.8"

-5.2

262 267 6-Hydroxylaminopurine 250 ribonucleoside 269

6-Bromopurine ribonucleoside 6-Iodopurine ri bonucleoside 2-Amino-6-methoxypurine ribonucleoside 2-Amino-6-chloropurine ribonucleoside

270 246 250 250 265 264

9

-

-

+4.16

+2.74

-

-8.33 -8.6b

-8.2"

-

-7.33' +0.300 +o. 120 -7.94

272 280 272 256 284

Na-Methyladenosine

6-Methoxypurine ribonucleoside 6-Chloropurine ribonucleoside

-

8

-8.6* -5.1 -8.86"

-5.37 -7.17 -4.55 +3.90 4-3. llb -11.00 -11.ooo

-

-11.2

-

-

-10.7

-

-

-9.7 -

-

-8.41

$3.65 -8.22

-7.90

-7.00

+3.76

4-3.90

+4.90

+7.02

+5.06

+5.26 -

-

-4.54

+6.3d +5.32 +5.66 -4.0

-

250

4-6.24

250

+6.44

257

+6.35

+6.35

+6.75

256 264

+9.91

+7.28 +9.86

+lO.lO

-

+11.4

Ref.

3.

53

PURINE AMINOHYDROLASEB

TABLE I (Continued)

PH

x Substrate

(nm)

2-Amino-6-iodopurine 255 ribonucleoside 340 7-Amino[l,2,5]oxy diazolo[3,4d] pyrimidine 7-AminoI 1,2,5] 340 thiadiazolo[3,4d] pyrimidine ~ ’ - D I X X ~ I ~ ~ O S ~ M -275 aminopyrazolo[3,4d] pyrimidine 2’-Deoxyribosyl-& 280 azaadenine 2’-Deoxyribosyl-&amino 278 adenine 9-Allyladenine 265 265 Q-(y,y-Dimethylsllyl) adenine 9-Benayladenine 265 4p-~-Glucopyranosyl 265 adenine 245 Guanine 250 255 260 Guanosine 240 8-Azaguanine 245 265

6

7

8

9

Ref.

+7.68 -6.1“

(67,73)

-7.80

(67,73) -8.63d -6.gd

(74)

-8.1

-7.33’ -7.54c -7.69’ -7.05’

-5.85

-7.05 -

-5.85 -5.3

-5.33

-

-3.9 -2.96 -2.22

-3.45 -

4-3.1 +3.1

Determined at pH 6.5.

* Determined at pH 7.4.

Determined at pH 5.6. Determined a t pH 7.2.

hypoxanthine without addition of dialyzable cofactors ( 7 6 ) .The kinetic behavior of enzyme from Candida utilis (77, 78) and Saccharomyces cerevisiae (79) was similar to the A . vinelandii preparation. Of the 76. L. A. Heppel, J. Hurwitz, and B. L. Horecker, JACS 79, 630 (1957). 77. W. D. McElroy, “Methods in Enzymology,” Vol. 6, p. 203, 1963. 78. R. C. Hartenstein and I. Fridovich, JBC 242, 740 (1967). 79. P. Medhat, Dissertation, Microfilm No. 68-71. University Microfilm, Inc., Ann Arbor, Michigan, 1965.

54

C. L. ZIELKE AND C. H. SUELTER

SUBBTRATEB FOR

TABLE I1 ADENINE AYINOHYDROLABE AND Azotobacter vinelandii

OF

Awtobaeter vinelandii

Candidu utilis Compound

K,

Adenine 7-Aminothiazolo[5,4d] pyrimidine 2,bDiaminopurine 6-Hydrazinopurine 6-Iodopurine 2-Amino-6-chloropurine 6-Chloropurine

x

104M

Candida I d d i S

Vrela

K,

x

lWM

VIei4

0.3 3

1.o 1.27

0.1 1.5

1.0 2.51

3.7 9.0

0.06 0.19 2.11 0.29

2.1 3.0 6.0 10 5

0.12 0.29 0.19 3.0 0.62

13 25

a Specific activity of enzyme from C. utilis,0.25 pmole/min/mg and from A , vinelandii 3.6 pmoles/min/mg, 0.01 M potassium phosphate, pH 7.0, 25’.

purine analogs which were substrates, the lowest K , was observed for adenine (Table 11) ; dehalogenation of 2-amino-6-chloropurine was 2-3 times more rapid than deamination of adenine. Guanine, cytosine, guanosine, cytidine, adenosine, 2’-AMP, 3’-AMP, 2’,3’-AMP, 5’-AMP, ADP, ATP, and G M P were deaminated a t less than one-two hundredth the rate for adenine (76). Adenine aminohydrolase of A. vinelandii does not catalyze the back incorporation of products, hypoxanthine or chloride, into 6-chloropurine during the course of hydrolysis when examined over a wide range of pH in contrast to the back incorporation of oxygen-18 into hypoxanthine catalyzed by adenosine aminohydrolase (80) (see Section 111). These results are consistent with a direct displacement of the 6 substituent by water rather than the intermediate formation of purinyl enzyme or chloroenzyme during catalysis. No information is available regarding the physiological function of this nonmammalian enzyme.

111. Adenosine Aminohydrolase

A. MOLECULAR PROPERTIES 1 . Purification

Homogeneous preparations of adenosine aminohydrolase (EC 3.5.4.4) have been obtained from mucosa of calf duodena (81, 82), chicken

3.

PURINE AMINOHYDROLASES

55

duodena (83),calf serum (84),and calf spleen (85).I n addition to the extensively purified preparation from ox heart (86), partially purified preparations fr m brain (87), bovine thyroid (88), Yoshida ascites cells (66), Escheric!t ia coli (33), Mycobacterium tuberculosis (89), Bacillus cereus (M), and cat lung (91)have been reported.

2. Chemical and Physical Properties The overall amino acid compositions of calf duodenal (92,92a) and calf spleen (85) enzyme showing an excess of acidic over basic amino acids are consistent with the isoelectric point of 4.85-5.0 previously reported for the calf and chicken duodenal enzyme (81,83). The N-terminal amino acid of the calf enzyme was not available for reaction with l-fluoro-2,4-dinitrobenzene in 8 M urea (92a).Calf spleen enzyme reportedly contained galactosamine and glucosamine (86) ; other preparations apparently have not been examined for carbohydrate. The molecular weight of the calf duodenal adenosine aminohydrolase determined from sedimentation and diffusion data (91) and by comparative elution from Sephadex gels (93,94) ranges from 31,000-35,OOO (84,91,93,95). A molecular weight of 52,000 from sedimentation velocity and sedimentation equilibrium data has not been confirmed (92). Molecular weight data for other preparations are consistent with additional forms of the enzyme; for example, chromatography of hepatic extracts of amphibia on Sephadex G-200 gave three peaks of adenosine aminohydrolase activity labeled type A, B, and C (96).Types corre~~

~

80. L. G. Howell and P. F'ridovich, JBC 242, 4930 (1967). 81. T. G. Brady and W. O'Connell, BBA 62, 216 (1962). 82. P. M. Murphy, M. Noonan, P. Collins, E. Tully, and T. G. Brady, BBA 171, 157 (1969). 83. V. D. Hoagland, Jr. and J. R. Fisher, JBC 242, 4341 (1967). 84. J. G. Cory, G. Weinbaum, and R. J. Suhadolnik, ABB 118, 428 (1967). 85. N. Pfrogner, ABB 119, 141 (1967); N. Pfrogner, ibid. 119, 147 (1967). 86. M. Rockwell and M. H. Maguire, Mol. Pharmacol. 2, 574 (1966). 87. M. K. Malysheva, U k r . Biokhim. Zh. 35, 764 (1963); C A 60, 4388e (1964). 88. W. Dierick, P. Olislaegers, and J. Stockx, Arch. Intern. Physiol. Bwchim. 75, 623 (1967); C A 68, 38095~(1968). 89. S. R. Guha, R . P. Saxena, and K. L. Arora, J. Sci. Ind. Res. (India) 21C, 66 (1962). 90.J . F. Powell and J. R. Hunter, BJ 62, 381 (1956). 91. P. M. Murphy, T. G. Brady, and W. A. Boggust, BBA 188, 341 (1969). 92. R. Wolfenden, Y. Tomozawa, and B. Bamman, Biochesmktry 7, 3965 (1968). 92a. J. Phelan, F. McEvoy, S. Rooney, and T. G. Brady, BBA 200, 370 (1970). 93. P. Andrews, BJ 96, 595 (1965). 94. J . R. Whitaker, Anal. Chem. 35, 1950 (1963). 95. T. G. Brady and M. O'Sullivan, BBA 132, 127 (1967). 96. P. F. Ma and J. R. Fisher, Comp. Biochem. Physwl. 27, 687 (1968).

56

C. L. ZIELKE AND C. H. SUELTER

sponding to B and C were also identified in chicken liver and rabbit duodena while only type C was found in cat lung and eight other animal duodena (83,91). Approximate molecular weights of the three types determined by comparative elution of standard proteins from Sephadex were: (A) 200,000, (B) lOO,OOO, and (C) 30,000 (96). Calf duodenal enzyme denatured with 8 M urea or 2 M guanidineeHC1, oxidized with performic acid, reduced with sulfite to cleave disulfide bonds and urea denatured carboxymethylated enzyme migrated on Sephadex G-75 as a single peak with a molecular weight near 35,000 consistent with a single polypeptide structure (82). Yet the calf duodenal enzyme (82,8,9,92) as well as the calf serum (84) and the human spleen, liver, heart, lung, and kidney enzyme (97') migrated as more than one band during electrophoresis. Chromatography of the calf duodenal enzyme on cellulose separated the enzyme into five somewhat overlapping peaks with similar kinetic and molecular properties. However, since only two isozymes were present in the mucosa of a single animal when examined immediately after death and also since the calf serum deaminase migrated in two major bands and two ininor bands (84), Murphy et al. (82) suggested that two isozymes of adenosine aminohydrolase are normally present in calf duodena; the additional bands were relegated to artifacts of preparation. No information is available regarding molecular weight differences. All adenosine aminohydrolase preparations have typical absorption spectra consistent with the absence of tightly bound prosthetic groups. No dissociable cofactors are necessary for catalysis since extensive dialysis of enzyme or addition of the metal chelator ethylenediaminetetraacetate (EDTA) did not cause significant inhibition.

B. CATALYTIC PROPERTIES 1. Reaction Parameters

The kinetic parameters K,, VIIIUX, pH optima, and energy of activation (Eact)for deamination of adenosine catalyzed by homogeneous preparations of adenosine aminohydrolase are summarized in Table I11 (8286, 98). Except for the unusually low specific activities observed for both the chicken duodenal and calf serum enzymes, and the significantly higher energy of activation reported for the former, the reaction param97. N. Ressler, Clin. Chim. Acla 24, 247 (1969). 98. 0.P. Chilson and J. R. Fisher, ABB 102, 77 (1963).

3.

57

PURINE AMINOHYDROLASES

TABLE I11 PREPARATIONS* KINETIC PROPERTIES" OF HOMOGENEOUS OF ADENOSINE AMINOHYDROLASE Enzyme preparation

VIW (pmolel (M X 106) min/mg)

K,

E.Ct

Temp. ("C)

pH optima

(kcal/ mole)

Ref.

~

Calf duodenal Isozyme mixture Isozyme 2 3 4 5 6 Calf spleen Ox heartb Calf serum Chicken duodenal

3-5 1.7 1.5 1.5 2.8 44 4.1 3.3 1.3

7.0-8.5 436 330 550

37 37 37

300 374 240 292 1.1 26

37 37

-

-

25 38 38

6.3 6.8-7.2 6.5-8.5 6.5-8.0

7.2

5.6 17.2 [Ref. (98)l

(98) (82)

(86)

(86) (84)

(83)

Kinetic data obtained from linear Lineweaver-Burk plots.

* Kinetic data for the ox heart preparation are included even though definitive data regarding homogeneity are not available.

eters are similar. The lower activities may reflect real tissue or species differences or some other as yet unknown property. I n contrast to the similar ratios of activity with adenosine compared to deoxyadenosine observed for mammalian tissue enzymes (42), lower ratios and higher activation energies were noted for the chicken liver and duodenal adenosine aminohydrolase (99). To define this more concisely Ma and Fisher (100) examined the adenosine-deoxyadenosine activity ratios, activation energies, and K , values of adenosine for the liver and duodenal enzyme from 6 orders of mammals, 5 orders of birds, 3 orders of reptiles, 2 orders of amphibia, 6 orders of bony fish, and 2 orders of cartilaginous fish. This extensive study showed that activation energies ranging from 4-10 kcal/mole, K , from to 6 X M , and adenosine-deoxyadenosine activity ratios from 0.6 to 1.5 are characteristic of vertebrates in general. In confirmation of the previous report (98) and in contrast to vertebrates in general, bird liver and bird duodenal adenosine aminohydrolase have activation energies in the range of 10-17 kcal/mole and lower adenosine-deoxyadenosine activity ratios. 99.J . R. Fisher, P. F. Ma, and 0. P. Chilson. Comp. Bwchem. Physiol. 16, 198 (1965). 100. P. F. Ma and J . R. Fisher, Comp. Biochem. Physiol. 19, 799 (1966).

C.

L. ZIFLKE AND C.

€3. SUFLTER

2. Nature of Active Site Graphical treatment of pKm vs. pH showed a relatively constant K , for adenosine between pH 5.5 and 8.5 for both the calf (65a) and chicken duodenal (98) adenosine aminohydrolase. This plot for the calf enzyme also exhibited inflection points a t pH 5.7-6.3 and pH 9.5-10.2 suggesting dissociable groups in the enzyme substrate complex with analogous pK values. Complete inactivation of the enzyme by equimolar concentrations of p-mercuribenzoate and phenylmercuriacetate and the -protection by 2,6-diaminopurine and 8-azaguanine (101), competitive and uncompetitive inhibitors, respectively, and lack of protection against inactivation by dinitrophenylation of 2-lysine €-amino groups by these same inhibitors (102) suggest but do not prove the involvement of sulfhydryl group(s) in the enzyme-substrate complex. On the other hand, mercuration of one sulfhydryl group of the chicken duodenal enzyme with p-mercuribenzoate produces a stable derivative with 2.5-fold greater activity than the native enzyme ; excess mercurating reagent always resulted in inactivation. Iodoacetamide does not appear to react with the enzyme but N-ethylmaleimide reacts to produce a 1.7-fold activation ( 8 3 ) . Schaeffer et al. (103) have synthesized and also demonstrated enzyme inactivation with two active-site-directed irreversible inhibitors, 9- (p-bromoacetamidobenzyl) adenine and 9- (0-bromoacetamidobenzyl)adenine. Inactivation proceeded by a first-order reaction following formation of an initial reversible inhibitor complex. Inhibition constants M , respectively, for the reversible enzyme M and 43 x of 1.4 X inhibitor complex evaluated from kinetics of inactivation agree with constants obtained by the usual double reciprocal plot of initial velocities. The first-order rate constants for alkylation in the reversible EI complex are 1.1 x min-I and 7.7 x lo-? min-l, respectively. The functional group undergoing alkylation has not been defined. The reversible and competitive inhibition of calf duodenal adenosine aminohydrolase by relatively low concentrations of urea, methyl urea, and 1,3-dimethyl urea is similar to the urea (104) inhibition of several other enzymes (105). The inhibition appears to involve 1-2 molecules of inhibitor in the inhibition complex (106)determined from log[ ( V d v i ) 101. G.Ronca, C.Bauer, and C. A. Rossi, European J. Biochem. 1, 434 (1967). 102. G. Ronca, P. L. Ipata, and C. Bauer, BBA 122, 379 (1966). 103. H.J. Schaeffer, M. A. Schwarts, and E. Odin, J. M e d . Chem. 10, 686 (1967). 104. T.G. Brady and M . O’Sullivan, BJ 80, 17P (1961). 105. K.V. Rajagopolan, I. Fridovich, and P. Handler, JBC 236, 1059 (1961). 106. G. Ronca and S. Ronca-Testoni, BBA 178, 577 (1969).

3.

PURINE AMINOHYDROLASES

59

11 vs. log I plots (107).Mixed inhibition studies (108)with several urea and amidine derivatives coupled with their mutually competitive interaction with 2,6-diaminopurine1 a substrate analog, are consistent with two independent but adjacent binding sites for these compounds (106). I n addition, the similar relationship between the inhibition and basicity of several amidine and purine derivatives (108, 109) suggests an interaction of the amidine derivatives with the purine binding site. The N-alkyl urea derivatives may bind a t a second presumably more apolar site consistent with a previous suggestion of Schaeffer and his colleagues (110).After examination of more than 40 9-substituted alkyl adenine derivatives as inhibitors of calf duodenal adenosine aminohydrolase, particularly the marked inhibition by 9- (1-hydroxymethyldecanyl)adenine, they suggested an active site composed of a purine binding region, an apolar region, and a hydrophillic region (110).In a similar vein, Bloch et a2. (111) concluded after comparison of 43 analogs of adenosine and 2’-deoxyadenosine as substrates or inhibitors of adenosine aminohydrolase that the 1’ and 5’ positions of adenosine were particularly important for substrate activity. The absence of substrate activity of the compounds lacking the 5’-hydroxyl suggests an important role for this group in binding. 3. Mechanism

Calf duodenal adenosine aminohydrolase affects enzymic hydrolysis of a wide variety of 6-substituted purine derivatives as well as analogs of adenosine with alteration in the purine ring and sugar moiety (Table IV) (65,65a,67,70,71,112). Although not noted in Table IV, AMP, ADP, and ATP are not substrates. The hydrolysis of the 6-methoxypurine derivative in H2180 occurs between C-6 and oxygen (113) consistent with the observed back incorporation of 180 from H2l80 into inosine as catalyzed by both calf duodenal and Takadiastase non107. F. H.Johnson, H. Eyring, and R. W. Williams, J. Cellular Comp. Physiol. 20, 247 (1942). 108. I. Fridovich, JBC 239, 3519 (1964). 109. G.Ronca and G. Zucchelli, BBA 159, 203 (1968). 110. H. J. Schaeffer and P. S. Bhargava, Biochemistry 4, 71 (1965); H. J. Schaeffer, D. Vogel, and R. Vince, J. Med. Chern. 8, 502 (1965); H.J. Schaeffer and D. Vogel, ibid. p. 507; H.J. Schaeffer, C. F. Schwender, and R. N. Johnson, J . Pharm. Sci. 54, 978 (1965); H.J. Schaeffer and C. F. Schwender, ibid. 57, 1070 (1968).

111. A. Bloch, M. J. Robins, and J. R. McCarthy, Jr., J. Med. Chem. 10, 908 (1967). 112. J. L. York and G. A. LePage, Can. J. Biochem. 44, 331 (1966). 113. R. Wolfenden and J. F. Kirsch, JACS 90, 6849 (1968).

60

C. L. ZIELKE AND C. H. SUELTER

TABLE IV

KINETICCONSTANTS FOR SUBSTRATES OF CALFDUODENAL ADENOSINE AMIINOHYDROLASE Substrate

Adenosine

(a) Derivatives of Purine Moiety 6.5-7.5 3.5 100

3-(B-D-Ribofuranosyl) adenine (isoadenosine) Adenine N1-Methyladenosine No-Meth yladenosine No-Ethyladenosine 6Hydroxyaminopurine ribonucleoside 6-Methoxypurine ribonucleoside 6-Chloroadenosine 6-Bromoadenosine 6-Iodoadenosine 2-Aminoadenosine 2-Amino-6-chloropurine ribonucleoside 2-Amino-6-iodopurine ribonucleoside 2-Amino-6-methoxypurine ribonucleoside 4-Aminopteridine 7-Amino[1,2,5]oxydiarolo [3,4d]pyrimidine 7-Amino[l,2,5]thiadiszolo 13,4d]pyrimidine &hinoadenosine 8-Oxoadenosine 8-Azaadenosine

6.5

7.7

0.52

6.5 6.5-8.9 7.0 6.8-7.0 7.0 6.8-7.0

15 9.6-10.1 9.6 0.6-1.1 12 7.14.5

6.8 6.8-7.2 7.0 7.0 6.8 7.0 6.8 7.0

3.2 25-70 9.8 30 3.4 6.7 30.5 41

127 59

6.8

5.0

12

6.5 6.5

10.5 38

6.5 7.0 7.0 7.0

333

0.0014 0.17 0.0014 0.16 0.14 23-38 0.45 2140 11 5 25

73

0.58 0.79 7.4

3.7 13.7 10.0 6.27 217 9.6 (b) 9-Substituted Adenine Derivatives 2’-Deoxyadenosine 7.2 2.2 93 3’-Deoxyadenosine 7.2-7.5 2.5-5.2 52 2’,3’-Dideoxyadenosine 7.2-7.5 8.7-7.7 3‘-Amino-2’,3’-dideoxyadenosine 7.2 8.3 3 2’-Amino-2’,3’-dideoxyadenosine 7.2 40 223 3’-Amino-3’-deoxyadenosine 7.2 16.7 103 2‘-O-Methyladenosine 7.0 2.2 53 2’-3’-Isopropylideneadenosine 7.0 4.8 49 4’-Thioadenosine 7.2 3.3 37

3. PURINE AMINOHYDROLASES

61

TABLE IV (Continued) Substrate 9-X ylosy ladenine 9-Arabinosyladenine 9-(3’-Deoxyarabinosyl) adenine 9-Lyxosyladenine 9-(5‘-Deoxylyxosy1)adenine 2’,(3)-O-(N-Benzyloxycarbonyl) glycyladenosine 9(c~-~2’-Deoxyribofuranosyl) adenine 9(@-~-5’-Deoxyxylofuranosyl) adenine

pH

7.5 7.2 7.2-7.5 7.5 7.5 7.5 7.0

K, ( M X I@)

VW

10

57 17 18-25 1.6 1.5 6.5 47

1.9 7.7-14 12 17.1 8 3.5

(rel)

7.0

6.1

8

7.0

6.7

5

Ref.

(c) Other Analogs

2’-Deoxyribosyl-8-azaadenine 2’-Deoxyribosyl-4-aminopyraaolo[3,4d]pyrimidine 2’-Deoxyribosyl-8-amino adenine

7.2 7.2 7.0

12.5 10.7 6.39

460 4.3 2.0

specific enzymes (115). While no rates were given for the very slow nonenzymic exchange into free inosine, the enzymic exchange rates were lower by approximately five orders of magnitude than the limiting rate constants for adenosine deamination by both the calf and Takadiastase enzymes (113). The reverse reaction, i.e., the direct conversion of inosiiie to adenosine catalyzed by both the calf duodenal and the Takadiastase nonspecific adenosine aminohydrolase (Section V) and rncasured as a function of pH with the calf enzyme, was defined by a theoretical curve for the equilibrium, Kc,= ( [inosine] [ NH,] ) / ( [adenosine][ H20]) = 38, with water concentration taken as one and pK, values of 8.8 and 9.2 for inosine and ammonium ion, respectively. The calculated AF = -5400 cal/mole a t pH 7.0 was in reasonable agreement with -6000 cal/mole estimated from the summation of a series of partial reactions (114). Because of the wide specificity of catalysis, Bar and Drummond (115) argued that the basic mechanism of action involves a nucleophilic displacement by hydroxyl at the 6 position of the purine system. Structural 114. R. Wolfenden, JBC 242, 4711 (1967). 115. H. P. Biir and G. I. Drummond. BBRC 24, 584 (1966).

62

C. L. ZIELEE AND C. H. SUELTER

considerations of the substrates suggested an analogy to some esterases and proteolytic enzymes. This comparison appears valid if one considers the amino tautomer of adenosine as the imido form of a carboxylic acid; thus, the 6-chloro, 6-methoxy, and 6-amino derivatives resemble an acid chloride, a methyl ester, and an amide, respectively. Enzymic deamination might also be facilitated by enzyme stabilization of the rare imino tautomer, as contrasted to the 6-amino tautomer which predominates by a factor of more than lo‘ (116). Enzymic activity on 1-methyladenosine (the neutral species of which is already in the imirio form) and on 3- (P-D-ribofuranosyl) adenine are consistent with the formation of the imino tautorner preceding hydrolysis (69). In an extension of the ideas of Bar and Drummond ( 1 1 5 ) ,Wolfenden (116) suggested the rate limiting formation of a tetrahedral intermediate a t the 6 position of purine involving enzyme or enzyme bound water and substrate similar to the type of intermediates generally encountered in nucleophilic aromatic substitution as indicated in ( I ) .

Several lines of evidence are consistent with this mcchanism. First, all known ring-modified substrates can form the tetrahedral intermediate ( 6 7 ) . Second, as previously detailed, both the calf duodenal and the nonspecific Takadiastase enzyme catalyze back incorporation of lSO into inosine. Third, the nearly identical I’,,,:,, for hydrolysis of 6-amino, -1iydrazin0, -chloro, -rnetliylamirio, and -methoxy purine riboside catalyzed by the Takadiastase enzyme when the apparcnt affinity varied by a factor of approximately 600 is indicative of the rate limiting hydrolysis of an intermediate common to all five reactions, one in which the original 6 substituent has been completely displaced (66).However, rates of hydrolysis catalyzed by the calf enzyme were not so similar (see Table I V ) . Furthermore, if a nucleophilic attack by the enzyme (or water bound to the enzyme) on the substrate were rate limiting, then the absence of a primary deuterium isotope effect observed with both enzymes suggests that cleavage of the leaving group before the transition state would be unlikely (117).T h a t is, the rate limiting hydrolysis 116. R. Wolfenden, J M B 40, 307 (1969). 117. R. Wolfenden, Biochemistry 8, 2409 (1969).

3.

63

PURINE AMINOHYDROLASES

of a common purinyl enzyme intermediate would be expected to involve a partial cleavage of water in the transition state. Also, electron withdrawing groups in the ring, for instance, the substitution of an oxygen or sulfur atom for the 8 carbon, enhance the V,,,,, as would be expected if nucleophilic attack constituted a part of the rate-limiting step (67). Deamination of both adenosine and deoxyadenosine is also enhanced by substitution of nitrogen for the 8 carbon (6'5, 74) and substitution of ribose at the 3 or 9 position results in very large rate enhancement as compared to adenine (67). However, Simon et al. (65) could establish no correlation between enhancement of activity and substitution of electron withdrawing groups a t the 8 position of adenosine. However, it was concluded that steric factors were important since those 8-substituted derivatives devoid of activity exhibited greater steric requirements than those which were substrates. Furthermore, Bar and Drummond (115) observed that the order of reactivity of 2-substituted adenosine was H>>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 I<,,&values for adenosine, 5'-AMP, 3'-AMP, 3',5'-cyclic AMP, and 2'-dAMP ranged from 0.1 to 0.5 mM. Except for 2'-dAMP, the relative rates of deaniination for these compounds were similar. The 2'-monophosphate ester was not deaminated (75, 180). Oligomers of AMP such as ApA, ApAp, ApApA, and ApApApA (181) were also substrates; deamination of the first residue of ApAp was more rapid than the second. Deamination of polyadenylic, polydeoxyadenylic, and the terminal residue of E . coli soluble RNA was not observed. TABLE VII SUBSTRATE SPECIFICITY OF ADENINE NUCLEOTIDE AMINOHYDROLASES

D. desulfuricansa

M . audouinic

P . crispatab

Substrate

Vm, (prnolel K, X (d) min/mg) pHopt 106 M

Vmax Km X (rel) pHopt 106 M

ATP ADP AMP Adenosine NAD

0.285 0.30 0.25 0.69 59

0.69 0.72 1.0 0.28 0.49

K,

13.8 12.6 14.2 0.98 30

6.0 6.0 6.8 7.1 5.6

6.6 4.7 4.7 56.0 7.2

5.0 5.0 5.0 5.0

3.3 4.7 8.3 52

Vmax

(re11 1 0.84 0.20

Conditions: 0.05 M sodium phosphate pH 5.8-6.0, temperature not given. Deaminations observed in 0.05 M phosphate buffer, 22";K , and Vmaxwere obtained from Lineweaver-Burk plots. Conditions 0.1 M acetate 25". a

179. S. Minato, J . Biochem. ( T o k y o ) 64, 815 (1968). 180. T. P. Wang, L. Shuster, and N. 0. Kaplan, JBC 206, 299 (1954). 181. R. Wolfenden, T. K . Sharpless, and R. Allan. JBC 242, 977 (1967).

3. PURINEAMINOHYDROLASES

75

Aspects of the mechanism of deamination catalyzed by the nonspecific Takadiastase enzyme are discussed in conjunction with calf duodenal adenosine aminohydrolase (see Section 111).

B. ADENINENUCLWTIDE AMINOHYDROLASE Three highly purified preparations catalyzing the deamination of ATP, ADP, and AMP have been isolated from Porphyra crispata (red marine alga) (182), Microsporum audouini (183), and Desulfovibrio desulfuric a m (grown anaerobically) (184). The enzyme from the red marine alga seaweed, purified 950-fold catalyzes the deamination of compounds noted in Table VII; 2’-AMP, 3’-AMP, NADP, and adenine were not substrates. Although no evidence regarding homogeneity was presented the constant ratios of activity for AMP: ADP: ATP: NAD :adenosine throughout the purification and heat inactivation data are consistent with a single enzyme. The reaction was activated by divalent cations Caz+,Mg2+,and Baz+; Ca2+was twice as effective as Mg2+ and Ba2+.The percent activation by Ca2+ of NAD, ATP, ADP, 5’-AMP, and adenosine deamination was 81, 260, 200, 116, and 0, respectively (182). The enzyme from M . audouini cultured on wheat bran, initially called AMP deaminase (185), has been extensively purified (4000-fold) (183) and renamed ATP aminohydrolase because of greater activity on that substrate. The ultracentrifugal sedimentation pattern exhibited a large and small peak, but no evidence was presented regarding the identity of either. Requirements for catalytic cofactors were negative. In addition to substrates noted in Table VII, 3’-AMP, 3’,5‘-cyclic AMP, dATP, dADP, dAMP, ADPribose, ADPglucose, 5‘-adenosine monosulfate, $-adenosine acetate, 5’-adenosine propionate, NAD, FAD, CoA, and nucleosidin were deaminated at appreciable rates. Deamination of ATP was inhibited by orthophosphate, pyrophosphate, triphosphate, and Fe3+;with adenosine none of these substances were effective inhibitors. Effects of other divalent cations were not reported. In contrast to the cation activation observed with the P . crispata enzyme, the preparation from D. desulfuricans 800-fold purified (184) way not activated by cations nor inhibited by EDTA, cysteine, and Na,S. Substrates other than those noted in Table VII were dATP, dADP, 182. 183. 184. 185.

J. C. Su, C. C. Li, and C. C. Ting, Biochemistry 5, 536 (1966). S. T. Chung and K. Aida, J . Biochem. (Tokyo) 61, 1 (1967). M. G. Yates, BBA 171, 299 (1969). K. Aida, S. Chung, I. Suzuki, and T. Yagi. Agr. B i d . Chem. 29, 508 (1965).

76

C.

L. ZIELKE AND

C.

H.STJELTEB

dAMP, deoxyadenosine, and ADPribose. The enzyme was inhibited by several anions ( K i ): cacodylate (134 mM), phosphate (83 mM), maleate (22 mM), succinate (18 mM) , citrate (6 mM), and &&dimethyl glutarate (3.2

a).

VI. Guanine hinohydrolase

Partially purified preparations of guanine aminohydrolase (EC 3.5.4.3) have been reported from rabbit liver (68,186), rat liver (61), Cbstridium acidurici (187),rat brain (60, 188, 189), and lingcod muscle (IN). The rat brain enzyme (60) occurs in both the mitochondria1 and supernatant fraction; the latter fraction yielded two forms, A and B from DEAEcellulose, which were subsequently purified 70- and 600-fold, respectively. Form B had a specific activity of 290 pmoles/min/mg. Kinetic, immunochemical, and electrophoretic studies revealed that the mitochondrial enriyme was distinct from supernatant enzyme B. A distinction between the supernatant A and B forms was less certain (60). Little is known of the physical properties of guanine aminohydrolase. Elution of rabbit liver enzyme from Sephadex gave a skewed activity peak; the estimated molecular weight of the main component was 170,000, a minor shoulder component, 525,000 (60). Substrates susceptible to enzymic hydrolysis by the lingcod muscle enzyme including K,,, and relative (V,,,,,J are: guanine, 0.033 mM, (1) ; 2-chloro-6-hydroxypurine,0.15 mM, (0.31) ; 2-hydrazino-6-hydroxypurine, 0.04 mM, (0.07); 1-methylguanine, 0.19 mM, (0.8); and 8azaguanine, 1 mM, (8.1) (191). Like adenosine aminohydrolase (see Section 111) guanine aminohydrolase requires no known cofactors and catalyzes both a dechlorination and hydrolysis of a hydrazino group. Likewise the 8-aza derivative of guanine was more rapidly deaminated than the natural substrates. Rat brain mitochondrial enzyme, however, did not affect deamination of 8-azaguanine (60). The enzyme was inhibited by NaCN (0.1 M, 45%), K F (lo-*M ,75%), and p-mercuribenzoate ( lcr4M, 100%); iodoacetate and CuSO,, l W 5 M, did not inhibit (68). The rat and sheep brain preparations exhibited a double pH optima, pH 4-6.5 and 8.0-9.0, with guanine as substrate (188, 186. G. H. Hitchings and E. A. Falco, Proc. Natl. Acad. Sci. U.S . 30, 294 (1444). 187. J. Rakosky, L. N. Zimmerman, and J. V. Beck, J . Bacterial. 69, 566 (1955). 188. M. Mansoor, G. D. Kalyankar, and G. P. Talwar, BBA 77, 307 (1963). 189. S. Kumar, K. K. Tewari, and P. S. Krishnan, J . Neurochem. 13, 1550 (1968). 190. R. Currie, F. Bergel, and R. C. Bray, BJ 104, 634 (1967). 191. J. E. Roy and K. L. Roy, Can. J . Biocbm. 45, 1263 (1967).

3. PUBINE A l U I N O H Y D W W ~

77

189);the rat brain particulate preparation exhibited a single pH 8.0 optimum after solubilization by Triton. The purified supernatant and mitochondria1 enzyme also had single pH optima in the range of pH 7-8 (60).Crude enzyme from lingcod muscle had double peaks of activity at pH 5.6 and 8.5, but after heat treatment during purification it had a single optimum of pH 6.0. While not understood, these double pH optima may reflect pH-dependent mitochondria1 membrane-enzyme associations or other enzyme-protein interactions which are destroyed during purification. A naturally occurring noncompetitive, proteinlike, heat labile inhibitor of guanine aminohydrolase isolated from rat eye lens (192),rat liver (60),or heavy mitochondrial fractions of rat cerebellum (60) may also be responsible. Since guanine aminohydrolase catalyzes the deamination of thioguanine and 8-azaguanine thereby destroying their anti-neoplastic effects, Baker and his colleagues have prepared a series of active site directed irreversible inhibitors to block the enzyme in tumor tissue (193).The most effective inhibitor, 9- (4-methoxy phenyl) guanine, effected a 50% inhibition at 0.38 pill in the presence of 13.3 pill substrate (194). The physiological role of guanine aminohydrolase is uncertain. The catabolic role in uric acid and allantoin production might be emphasized especially with respect to the deficiency of this enzyme in pig liver and spleen which is believed to be responsible for a form of gout observed in pigs in which crystals of guanine are deposited in joints (196).A lack of or deficiency of this enzyme may also account for the interesting distribution of guanine in nature (195). Evaluation of serum guanine aminohydrolase activities has been suggested as a quantitative indication of liver cell damage (196).

VII. Guanosine Aminohydrolase

A guanosine aminohydrolase preparation from Pseudomonas convexa No. 149 free from guanine, adenine and adenosine aminohydrolase and nucleoside phosphorylase activities has been described (63). Of the 22 192. S. Kumar, A B B 126, 964 (1968). 193. B. R. Baker, J . Chem. Educ. 44, 610 (1967). 194. B. R. Baker and W. F. Wood, J . Med. Chem. 10, 1106 (1967). 195. A. Bendich, in “The Nucleic Acids” (E. Chargaff and J. N. Davidson, eds.), Vol. 1, p. 95. Academic Press, New York, 1955. 196. S. McLeod, Can, J . Med. Technol. 29, 60 (1967); E. M. Knights, Jr., Progr. Clin. Pathol. 1, 318 (1966); E. BowkiewicrSurma and J. Krawcznski, Clin. Chim. Acta 16, 26 (1967).

78

C.

L. ZIELKE AND

C. H. SUELTER

purine compounds examined as substrates, guanosine, deoxyguanosine, M, and 8-aaaguanosine were deaminated with K,,, values of 3.6 X 6.2 X M , and 1.22 X lo-' M , respectively. No cofactors were required and optimal activity for guanosine was observed between pH 6 and 6.5. The molecular weight obtained by Sephadex G-100chromatography was 1-2 x 105.

Glutaminases and y.Glutamy1transferases STANDISH C. HARTMAN I . Introduction

. . . . . . . . . . . . . . . . . . . . . . A. Occurrence . . . . . . . . . . . . .

I1. Glutaminase of Escherichia coli .

B . Assay . . . . . . . . . . . . C . Purification . . . . . . . . . . . D. Specificity . . . . . . . . . . . E . Acyl Transfer Reactions . . . . . . . . F. Reactions of Glutaminase with 6-Diarod-oxonorleucine G . Other Inhibitors . . . . . . . . . . H . Effects of pH upon Kinetic Parameters . . . . I . Effect of Temperature . . . . . . . . J . Effect of Deuterium Oxide . . . . . . . K . Relationship to Other Acylases . . . . . . L . Mechanism of Action . . . . . . . . . 111. Other Glutaminases and Glutamyltransferases . . . . . A . Survey . . . . . . . . . . . . B . y-Glutamyltransferase from Agaricaceae . . . . C . y-Glutamyl Transpeptidases from Kidney . . . . D . Glutaminase from Azotobacter agilis . . . . . IV . Concluding Remarks . . . . . . . . . . .

. . . . .

. .

79 80 81 81 82 82 84 85 87

88

.

88

. . .

90 90 90 93 93

. . . . .

.

95

96 97 98

.

I Introduction

Enzymes which catalyze cleavage of the 7-acyl bond in glutamine are classified either as glutamine amidohydrolases (EC 3.5.1.2) (glutamin79

80

STANDISH C. HARTMAN

ases) or as glutamine glutamyltransferases (EC 2.3.2.1), depending upon whether the acyl acceptor is water or some other substance. I n a number of cases, categorization is necessarily somewhat arbitrary since the enzymes involved effect both hydrolysis and transfer to amines. The distinction also tends to obscure the probable mechanistic similarities of these processes. It may be useful therefore to review comparatively enzymes which, in common, transfer the acyl constituents of glutamine and related compounds. The major part of this article will be concerned with the acid glutaminase of Escherichia coli. Representative glutaminases and glutamyl transferases from bacterial, plant, and animal sources will be discussed more briefly. Considerable information on the phosphate-activated glutaminase of animal tissues is given in the previous edition of “The Enzymes” ( 1 ) . Glutamyl transferases, which act exclusively on glutaminyl residues in peptides and proteins such as the calcium-activated enzyme from guinea pig liver ( 2 ) , will be omitted from this discussion as will the class of biosynthetic enzymes which transfer the NH, group of glutamine to various activated acceptors (3).Glutamyltransferase activity dependent upon adenosine nucleotides and phosphate or arsenate, which is apparently associated with glutamine synthetase (4,5), also is not included in this coverage.

II. Glutaminase of Escherichia coli

I n 1955, Meister et al. (6) described the partial purification of an enzyme from E . coli which catalyzed the hydrolysis of L-glutamine to glutamic acid and ammonia and which was maximally active in the acidic region of pH. The enzyme showed a relatively high specificity for Lglutamine since it would not hydrolyze a large number of related amides. I n addition to hydrolysis, the enzyme catalyzed the formation of yglutamyl hydroxamic acid when hydroxylamine was present. More recently the study of this system was undertaken in the author’s laboratory with the expectation that information concerning the enzymic hydrolysis 1. E. Roberts, “The Enzymes,” 2nd ed., Vol. 4, p. 285, 1960. 2. D. D. Clarke, M. J. Mycek, A. Neidle, and H. Waelsch, ABB 79, 338 (1959); J. H. Pincus and H. Waelsch, ibid. 126,34 and 44 (1968) ; J. E. Folk and P. W. Cole, JBC 240, 2951 (1965) ; J. E. Folk and P. W. Cole, ibid. 241, 5518 (1966). 3. A. Meister, “The Enzymes,” 2nd ed., Vol. 6, p. 247, 1962. 4. A. Meister, “The Enzymes,” 2nd ed., Vol. 6, p. 443, 1962. 5. P. K. Stumpf, “Methods in Enzymology,” Vol. 2, p. 263, 1955. 6. A. Meister, L. Levintow, R. E. Greenfield, and P. A. Abendschein, JBC 215, 441 (1955); A. Meister, “Methods in Enzymology,” Vol. 2, p. 380, 1955.

4.

GLCTAMISASES AND 7-GLUTAMYLTRANSFERASES

81

of glutamine would be of value in understanding more complex biosynthetic reactions in which transfer of the NH, group occurs from this substrate to an organic acceptor (7-9).

A. OCCURRENCE Cells of E . coli B are found to produce significant amounts of this enzyme only after the stage of rapid exponential growth has ceased. All attempts to enhance or to repress its synthesis by altering the composition of the growth medium or conditions of culture have been unsuccessful. Glutaminase is not released when the cells are converted to protoplasts in isotonic sucrose, but the enzyme is solubilized upon lysis of the protoplasts. Strain W also possesses the enzyme, but no extensive screening of related species has been done.

B. ASSAY In crude enzyme systems the most sensitive assay depends upon detecting ammonia produced in the reaction: Glutamine -I-HZO -+ glutamate

+ NH,'

(1)

This product can be observed in the presence of glutamine with the use of a modified Nessler's reagent (10). More conveniently, a pH-stat assay may be employed which takes advantage of the fact that in the p H range of enzymic activity protons are taken up during the hydrolysis owing to partial protonation of the glutamate formed ( 7 ) . From the observed pK,' of glutamic acid (4.31 at 25", ionic strength 0.2), the relationship between rate of proton uptake required to maintain any constant pH and the rate of glutamine hydrolysis can be directly obtained. The hydrolysis of other glutamic acid derivatives catalyzed by the enzyme can also be followed by this method, except that ester substrates consume alkali during their hydrolysis. Also, when amides which release partially protonated amines (such as hydroxylamine and O-methylhydroxylamine) are hydrolyzed, a correction must be applied to obtain the proper relationship between titrant addition and net reaction. 7. S. C. Hartman, JBC 243, 853 (1968). 8. R. A. Hammer and S. C. Hartman, JBC 243, 864 (1968). 9. S. C. Hartman, JBC 243, 870 (1968). 10. A. P. Vanselow, Ind. Eng. Chem.. Anal. Ed. 12, 516 (1940).

82

STANDISH

C.

HARTMAN

C. PURIFICATION Purification of the glutaminase involves precipitation of impurities a t low pH, chromatography upon DEAE-cellulose, ammonium sulfate fractionation, gel filtration upon Sephadex G-100, and a heat treatment ( 7 ) . I n more recent preparations the gel filtration step has been omitted. After the heating step, the solution is thoroughly dialyzed against 0.02 M sodium acetate, p H 5.0, clarified by centrifugation, and then passed through a small column of carboxymethyl cellulose previously equilibrated with the same buffer. The enzyme, which passes directly through this column, is concentrated by ultrafiltration. The specific activity at this stage is about 50% of the maximal value observed with samples subjected to zone electrophoresis. The latter preparations appear to be better than 90% pure as judged by analytical gel electrophoresis. A net purification of about 3000-fold is achieved based upon the total soluble protein of the bacterial extracts. The enzyme preparation prior to the electrophoresis step may be stored for many months a t 2" with little loss in activity.

D . SPECIFICITY Kinetic parameters for the known substrates of glutaminase are reported in Table I ( 7 ) , the k, values (turnover numbers per active site) being given by V,,,,,/Eo. Values for active site concentration (E,) were determined from the amount of 14C-6-diazo-5-oxonorleucine(DON), which reacts covalently with the enzyme (see below), with the assumption that the number of molecules of this substrate analog bound is equivalent to the number of substrate-reactive sites. Kagan et al. (If) reported that the enzyme acts on a-methyl-L-glutamine, but rate data comparable to those of Table I are not available. Substances not hydrolyzed at a measurable rate (koless than 0.1 sec-') include L-glutamic 7-ethylamide, D,L-glutamic y-dimethylamide, L-glutamic y-anilide, Lglutamic y-p-nitroanilide, L-glutamic y-2-hydroxyethy tmide, L-glutamic y-2-chloroethylamide, D,L-glutamic y-n-butylamide, L-glutamic diamide, glutaramic acid, N-acetyl-L-glutamine, D-glutamine, D- or L-asparsgine, L-2-amino-4-cyanobutyric acid, ~-2,2,2,-trifluoroethylL-glutamate, L-pyroglutamic acid, S-carbamoyl-L-cysteine, O-acetyl-L-serine and Lisoglutamine. Meister e t al. previously reported negative results with L-homoglutamine as well as with some of the above substances (6). 11. H. M. Kagan, L. R. Manning, and A. Meister, Biochemistry 4, 1063 (1965)

4. GLUTAMINASES

83

AND 7-GLUTAMYLTRANSFERASES

TABLE I KINETICPROPERTIES OF SUBSTRATES Substraten

hb

KC

(sec-l)

(mM)

1265 8.0 14 212 296 5O8Od 645 36 1260 300

0.42 3.3 12 5.1

~~

Glutamine 7-Glutamyl methylamide 7-Glutamyl hydraside 7-Glutamyl hydroxamic acid r-Glutamyl methoxyamide Glutamic acid 7-Methyl glutamate 7-Ethyl glutamate 7-Thio methyl glutamate 7-Thio ethyl glutamate

configuration. numbers per 6-diaeo-5-oxo-~-norleucinebinding site at 25" and pH 5.0. Estimated error in /GO, f 5%. Estimated error in K,, f 20%. The values for glutamic acid are determined from the rates of exchange of 1 6 0 between HZl80and the 7-carboxyl oxygens of this substrate (8);the kinetics of hydroxamate formation yields K , = 2.0 mM. 0

All materials were of the

5.0 2.gd 64 50 10 23

* ko values are the turnover

L

While not hydrolyzed by the enzyme, the following compounds act as competitive inhibitors: L-glutamic y-ethylamide ( K , = 2.8 mM) , L-glutamic y-2-hydroxyethylamide ( K , = 52 mM) , and 2,2,2-trifluoroethyl glutamate ( K , = 67 mill). Other substances related to glutamine which are neither substrates nor inhibitors are L-glutamic diamide, N-acetyl-L-glutamine, glutaramic acid, L-glutamic y-anilide, L-2-amino4-cyanobutyric acid, S-carbamoyl-L-cysteine, 0-acetyl-L-serine, D,L-methionine sulfoxide, D,L-methionine sulfone, D,L-methionine sulfoximine, D,L-allylglycine, and ~,~-2-amino-4-iodobutyric acid. The structural requirements for a substance to be a substrate for the glutaminase are relatively stringent: the L- configuration and a free amino and a-carboxyl group are necessary; also, a carbonyl group on the C-5 atom is apparently essential. I n general, the rate of hydrolysis within a given class of compounds is inversely related to the size of the substituent in the y position (NH, > NHCH, > NHCH,CH,; OH > OCH, > OCH,CH,, etc.), In comparing substituents of similar size, the sulfur and oxygen compounds are cleaved more rapidly than amides (OH > NH,; SCH, > OCH, > NHCH,; etc.). The K , (or K , values are most obviously affected by the chemical nature of the group; the esters and the amides each have a characteristic range of substration parameters. This generalization does not hold in the case of the very (Y-

84

STANDIGH C. EABTMAN

small substituents NH, and OH, which have significantly lower K,,, values than other N or 0 compounds. As the size of the group increases, the hydrolytic rate constant first drops to zero, without a correspondingly large change in K , or K , but after a critical size is reached (e.g., dimethylamide or 2-chloroethylamide) the compound is totally rejected by the enzyme. The contrasting substituent effects upon K,,, and k, indicate that the events of substrate binding and catalysis are governed by largely independent factors. At least for the poorer substrates, the K , values probably approximate dissociation constants of the Michaelis complexes.

E. A c n

TRANSFER

REACTIONS

I n the presence of hydroxylamine, glutaminase catalyzes the synthesis of y-glutamyl hydroxamic acid from all of the substrates in Table I. The rate of this reaction with either glutamine or glutamic acid is linearly related to the concentration of the free base of hydroxylamine over a range of p H values (7).At substrate saturation, a catalytic rate constant for reaction of any substrate with hydroxylamine can therefore be defined as kN = VN/(NH*OH) ( E , ) , where V , is the zero-order rate of hydroxamic acid formation. The relative rates of hydroxylaminolysis and hydrolysis have been determined for a number of substrates (glutamine, glutamic acid, y-methyl glutamate, and 7-thio methyl glutamate), and the ratio kN/k, is observed to be the same for all within experimental error (2.&2.2 M-l) (9). The apparent superiority of hydroxylamine compared to water is even more evident if the rates of hydrolysis are expressed for water in the standard state of moles per liter (i.e., k0/55.5).On a molar basis, hydroxylamine reacts about two orders of magnitude more rapidly than water. Under conditions of experimental measurement, however, when the concentration of NH,OH is on the order of 0.02 M (at pH 5 ) , observed velocities of hydrolysis are much larger than those of hydroxylaminolysis, as noted by Meister et al. ( 6 ) . The equilibrium constant of the reaction RCOOH

+ NHRH

RCONHOH

+ H R ( R = - CH&H.&H(NHs+) COO-)

(2)

as catalyzed by the enzyme is observed to be

when the activity of water is taken as 1, or 1.1 X 10' when it is expressed as 55.5 M.This is a pH-independent constant when the concentrations of the species are given in the ionic forms shown. This value agrees within

4.

85

GLUTAMIMASES AND y-GLUTAMYLTRANSFERASES

a factor of two with that calculated by Ehrenfeld et al. for the similar process catalyzed by an enzyme from Azotobacter agilis (12) or with that found by Jencks et al. for related equilibria of homogeneous phase reactions of carboxylic acids with hydroxylamine (IS). At equilibrium, therefore, partitioning of the acyl group between hydroxylamine and water favors the former by about lo4. I n the reaction catalyzed by E. coli glutaminase, the kinetic partitioning ratio [ kN(HzO)/k,] was seen to be only lo2. Glutaminase also catalyzes acyl transfer from glutamic acid to methanol by the reverse of the reaction of methyl glutamate hydrolysis (7) : Glutamic acid

+ methanol

methyl glutamate

+H a

(4)

The rate constant k, = V,/(CH,OH) ( E o ) is 0.47 M-' sec-' a t 25" and p H 5.0, a value which may be compared to that for the virtual hydrolysis of glutamic acid Ic,/(H,O) of 91.5M-' sec-'. Thus, from the ratio k, (H,O)/lc,, it is seen that the rate with methanol is some 194 times slower on a molar basis than that with water even though methanol is generally observed to be a somewhat better nucleophile than water in similar nonenzymic substitution reactions ( 1 4 ) . These results indicate that the enzyme shows a large preference kinetically for water over hydroxylamine or methanol compared to what would be expected on the basis of the intrinsic reactivities of these substances.

F. REACTIONS OF GLUTAMINASE WITH

6-DIAZO-5-OXONORLEUCINE

In the presence of an excess of the glutamine analog, DON, the enzyme is readily inactivated irreversibly ('7). This substance behaves as an active-site-directed alkylating agent in a number of enzymes which utilize glutamine as substrate ( 3 ) . With the use of 6-14C-DON, the amount of inhibitor covalently bound to protein can be directly measured; and, as shown in Fig. 1, this amount is linearly correlated with the extent of inhibition of catalytic activity. The simplest explanatioii of this behavior is that irreversible reaction of one molecule of DON with the enzyme inactivates one catalytic site, probably by combination with an essential group a t that site. With this assumption the concentration of active sites in an enzyme preparation may be calculated for the 12. E. Ehrenfeld, S. J. Marble, and A. Meister, JBC 238, 3711 (1963). 13. W. P. Jencks. M . Caplow, M. Gilchrist, and R. G. Kallen, Biochemistry 2, 1313 (1963). 14. M. L. Bender, G . E. Clement, C. R. Gunter, and F. J. KBsdy, JACS 66, 3697 (1964).

86

STANDISH C. HARTMAN

4

c

0 3 x

-e0

E i

:2 = n 0 z

x I

I

I

5

10

’

15

11.0

DON added, prnoles X I O ~

FIG.1. Titration of glutaminase with “C-DON. The amount of labeled DON bound to 20.7 p g of enzyme protein is shown. (0) DON bound; (0) enzymic activity.

purpose of expressing kinetic values on an absolute basis. In studies with the essentially pure enzyme it is found that one mole of DON binds to 50,000 g of protein. Comparison with an approximate molecular weight for the protein of 100,000, estimated by gel filtration on a calibrated column, indicates that two identical catalytic subunits comprise the molecule. It may be seen from the proportional region of Fig. 1 that only a small fraction, about 1/70, of the DON added to the enzyme becomes covalently bound. Further analysis of this phenomenon has shown that glutaminase catalyzes the hydrolytic cleavage of the ketone and that the rate of this reaction is 70 times that of the irreversible inactivation of the enzyme. Once alkylation of all reactive sites is complete, the further catalytic hydrolysis of DON is totally blocked. After reaction of 6-14C-DONwith glutaminase, glutamic acid ( 1 equivalent) and I4C-methanol (0.75 equivalent) were isolated from the reaction mixture. The initial product formed from the C-6 atom appears to be diazomethane ( 1 4 ~ )If. relatively high concentrations of a benaoic acidbenzoate buffer are included in the reaction mixture, labeled methyl benzoate is formed in addition to the methanol. Further, the relative 14a. S. C . Hartman and T. McGrath, unpublished observations (1969).

4.

87

GLUTAMINASES AND y-GLUTAMYLTRANSFERASES

amounts of methyl benzoate and methanol formed are identical to those observed when 14C-diazomethane itself is added to the benzoate buffer system a t the same p H and concentration. The reactions are viewed as occurring as follows:

+

+

Enzymic: RCOCHNz HzO + RCOOH CHZNZ Nonenzymic: CHZNZ HzO -+ CH,OH Nz CHZNZ beiizoic acid + methyl benzoate

+ +

+

(5)

+ Iyz

(6)

(7)

The C-C bond cleavage catalyzed by the enzyme appears to be very similar to the reactions of all other derivatives of glutamic acid. A precedent for this C-C bond cleavage exists in the action of chymotrypsin upon ethyl 5- (p-hydroxyphenyl)-3-oxovalerate (15).Which of the two modes of reaction occurs between DON and glutaminase may depend upon whether a catalytic group on the enzyme attacks C-5 of the analog, leading to hydrolysis, or C-6, leading to irreversible alkylation. The fact that l’C-DON labeled in positions 1-5 gives rise to labeled enzyme and, after cleavage with trypsin, to peptides identical to those derived with the use of 6-14C-DON indicates that the diazomethane released is not the primary inhibitor. The site of covalent attachment of the inhibitor moiety has not yet been defined. G. OTHERINHIBITORS Glutaminase is sensitive to inhibition by heavy metal ions and their derivatives. Mercuric ion and p-mercuribenzoate inhibit completely a t 0.1 mM; silver, lead, and cupric ions are effective a t somewhat higher concentrations. The presence of glutamine prevents inhibition by p-mercuribenzoate. Other divalent metal ions, including magnesium, manganese, zinc, cadmium, cobalt, ferrous, and calcium, neither inhibit nor activate a t 1 mM. The metal chelating agents ethylenediaminetetraacetate (EDTA) , o-phenanthroline, and dithiothreitol do not inhibit but instead result in slight activation. Despite the inhibition by heavy metals, iodoacetate, iodoacetamide, and N-ethylmaleimide, which generally react with sulfhydryl groups, are not inhibitory when the enzyme is exposed to them a t either pH 5 or 7. Glutaminase is similarly insensitive to diisopropylphosphorofluoridate (DFP). Phthalein dyes, such as bromocresol green, strongly inhibit glutaminases from kidney (1) and from Clostn’dia (16),but they do not affect the E . coli enzyme. 15. D. G. Doherty, JACS 77, 4887 (1955). 16. D. E. Hughes and D. H. Williamson, BJ 51, 45 (1952).

88 H. EFFECTS OF pH

STANDISH C. HARTMAN

UPON

KINETICPARAMETERS

The E . coli glutaminase functions effectively only in the acidic pH region. Values of K , and k , have been evaluated over the whole pH range of interest for a good ester substrate (methyl glutamate) and for a poor amide substrate (glutamyl methylamide), as well as over a more restricted ranges for glutamic acid and glutamine (9). I n all cases the maximum velocity for any substrate is essentially independent of pH over the entire accessible range. The K , values are also constant from below pH 4 to about 5.3. (The fact that the K,,, for glutamic acid is also constant in this range indicates that the protonated form of glutamic acid rather than glutamate ion is the true substrate.) Above about pH 5.3, K , values rise as a very steep function of pH as shown in Table 11; thus, when a pH above 5.8 is reached, substrates virtually cease to be bound by the enzyme. As Peller and Alberty have shown, the dependence of IC,/K, upon p H indicates the dissociation of essential acidic groups on the free enzyme (17).I n the present case this ratio changes with pH four to five times more steeply than would be expected if a single acidic group were required in the free enzyme. The results are consistent with the view that the state of the enzyme capable of binding substrate is stabilized by the cooperative action of several acidic groups. Dissociation of any one of these groups transforms the enzyme into the inactive form as the cooperative stabilization is lost. Since there is no additional effect of p H upon k,, it appears that all catalytic groups are in their proper state of dissociation in the ES complex and are not permitted to change while the complex exists.

I. EFFECT OF TEMPERATURE The variations of velocity with temperature over the range 5 " 3 5 " were measured for several substrates (9). Linear Arrhenius plots of these data were obtained which allowed calculation of the empirical activation energy and AH^. A F ~values a t 25" were calculated from the first-order rate constants ko expressed in units of sec-'. These values and the entropies of activation are given in Table 111. The K , values for glutamine have also been determined over this temperature range and found not to vary from the value a t 25" (4.32 mM) by an experimentally significant amount. 17. L. Peller and R. A. Alberty, JACS 81, 5907 (1959).

4.

89

GLCTAMINASES AND y-CLUTAMTLTRANSFERASES

TABLE I1 KINETIC PARAMETERSO

EFFECTOF pH

UPON

~

K, (mM)

Substrate and pH

4.00 4.50 5.00 5.30 5.40 5.50 5.60 5.70

Methyl glutamate 58 64 60 70 122 270 555 >2000

3.50 4.00 4.50 5.00 5.30 5.40 5.50 5.60

Glutamyl methylamide 3.35 3.20 3.41 3.30 3.70 4.65 5.50 14.3

4.00 4.50 5.00 5.40

Glutamine 0.53 0.48 0.42 0.92 Glutamic acid 2.0

5.00 4.30 a

b

ko (sec-l)

537 615 645 603 670 683 634 b 7.4 8.1 8.0 8.0 8.7 8.4 9.2 9.0 1250 1220 1265 1190

5080 4830

All values are at 25" and ionic strength 0.2 M. Indeterminate. TABLE I11 ACTIVATION PAR.4METERSa

~

~~

ko

AF:

AH:

Substrate

(sec-l)

(kcal mole-')

(kcal mole-l)

AS: (eu)

Glutamic acid Glutamine ?-Methyl glutamate 7-Ethyl glutamate 7-Glutamyl methylamide

5080 1265 645 36 8.0

12.4 13.2 13.6 15.4 16.2

8.2 7.9 8.1 10.2 14.2

-14.1 -17.8 -18.4 -17.4 -6.7

a

Values were determined at 25",ionic strength 0.2M . Estimated error in AH:, f 0.5

kcsl mole-'; in AS:, f 2 eu.

STANDISH C. HARTMAN

J. EFFECT OF DEUTERIUM OXIDE Rates of hydrolysis of several substrates in water and in D,O have been compared (9). Average values of VH20/VD,0were, for glutamine, 1.16; for methyl glutamate, 1.38; for glutamyl methylamide, 1.10; and for glutamic acid, 1.50.

K. RELATIONSHIP TO OTHERACYLASES A majority of the known enzymes capable of hydrolyzing carboxylic derivatives (acylases) can be placed into four categories according to their mode of action or key functional groups (18): (1) serine esterases, typified by chymotrypsin and subtilisin; (2) thiol acylases such as papain ; (3) metal dependent enzymes, including carboxypeptidase and various aminoacylases; and (4) acid proteases such as pepsin. Glutaminase catalyzes reactions similar to those of the above enzymes, including the ability to act on various types of carboxylic derivatives and to transfer the acyl groups to water and other acceptors. A more detailed comparison, however, strongly indicates that the glutaminase does not fall into these previously recognized categories. Table IV summarizes certain characteristics of representative enzymes from these classes and those of glutaminase (19-96). The distinctions between glutaminase and these enzymes are clear enough, it is felt, to warrant the conclusion that primary differences must exist in the fundamental aspects of the catalytic process. The strong possibility that glutaminase acts upon carboxylic derivatives by a novel mechanism and with a catalytic efficiency substantially greater than that of other acylases make it an intriguing system for further study.

L. MECHANISM OF ACTION Hydrolytic enzymes may be regarded as catalysts of group transfer processes. It is often the case in such processes that the group being 18. B. S. Hartley, Ann Rev. Biochem. 29, 45 (1960). 19. G. P. Hess, “The Enzymes,” 3rd ed., Vol. 111, p. 213, 1971. 20. E. L. Smith, “The Enzymes,” 3rd ed., Vol. 111, p. 501, 1971. 21. W. Lipscomb and J. A. Hartsuck, “The Enzymes,” 3rd ed., Vol. 111, p. 1, 1971. 22. J. S. Fruton, “The Enzymes,” 3rd ed., Vol. 111, p. 119, 1971. 23. B. Zerner, R. P. M. Bond, and M. L. Bender, JACS 86, 3674 (1964). 24. J. R. Whitaker and M. 2, Bender, JACS 87, 2728 (1965). 25. W. 0. McClure, A. Neurath, and K. A. Walsh, Biochemistry 3, 1897 (1964). 26. K. Inouye and J. S. Fruton, JACS 89, 187 (1967).

P 0

TABLE IV COMPARISON O F E. COli GLUTAMINASE WlTH Chymotrypsina Principal functional groups pH optimum Selective inhibitors

Serine, histidine >7 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).

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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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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.

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JOHN C. WRISTON, JR.

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).

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JOHN C. WRISTON, JR.

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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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.

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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.

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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).

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JOHN C. WRISTON, JR.

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).

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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

JOHN C. WRISTON, JR.

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'

>

<30 min

Very short

a-2-Globulin behavior. Cellulose acetate strip electrophoresis. 0 Isoelectric focusing, Mashburn and Landin (86). Broome (26). ' Mashburn and Landin (86). J

Protein synthesis is the only known major role for asparagine in mammalian systems. Both Sobin and Kidd (99) and Broome (91) have shown that the capacity of asparagine-dependent tumors to synthesize protein is impaired in the absence of this amino acid. Nevertheless, the question is not yet resolved as to whether the dramatic effect of asparaginase on leukemic cells can bc ascribed solely to interference with this process or whether other metabolic functions are affected indirectly. Ryan and Sornson (100)have shown recently that the administration of asparaginase depressed the level of tumor glycine specifically in 98. J. D.Broome, Trans. N . Y . Acad. Sci. [21 30, 690 (1968). 99. L. H.Sobin and J. G . Kidd, Proc. SOC.Ezpll. Bid. M e d . 119, 325 (1965). 100. W.L. Ryan and H. L. Sornson, Science 167, 1512 (1970).

120

J O H N C. WRISTON, J R .

mice, and they suggested that the loss of cellular glycine may be more important than loss of asparaginc because of the rcquiremeiit for glycine in purine synthesis. A decrcase in glycine in animals treated with asparaginase might arise if tlic tumor synthesizes it from glyoxylic acid and asparagine, a route to glycinc which has been sho~vnI)y RIcister to occur in liver (1). Several factors arc involved in thc widc variation in tumor inhibitory activity of asparaginases from different sources (98). One obvious possibility is the rate of clearance of tlic enzyme from the circulation of the host animal, a i d Broonie (26) was tlie first to obtain cvidence iniplicating half-life as a factor iii antitumor effectiveness. Guinea pig serum asparaginasc, for example, has a half-life time of 11-19 hr, while a partially purified yeast asparaginase preparation without antilyniphoma activity is almost completely cleared within 30 min. Differences in half-life alonc cannot explain the differences in antitumor activity in all cases, however; and the question still remains as to what structural features are responsible for rapid or slow clearance. Mashburn and Landin (85) have suggested that differences i n half-life time may be related to the isoclectric point of the enzyme, and evidence also exists to hupport the itlen that t l i o tumor inliibitory activity of sonic asparaginases is related to their K,,, values (59, 6 7 ) . An additional factor has recently been shown to affect the clearance time of the E . coli asparaginase and its pattern of tumor inliibitory activity. Mashburn et al. (101) and Boyse et al. (873 observed that E. coli asparaginase was far less effective than the GPS enzyme when atlministered a t the same time that mice were inoculated with tumor cells but was more effective than the GPS enzyme against established tumors. An explanation for this apparently paradoxical finding was provided by the work of Riley et al. (102) and of Notkins and Scheele (103),who had shown earlier that a “lactic dehydrogenase elevating virus” ( L D H virus) is carried by 6C3HED lymphosarcoma cells and leads to a decrease in the rate a t which certain enzymes are removed from the blood. The virus does not affect the half-life of all enzymes (I&), but it has been shown (41, 59, 105) that the L D H virus does have a pronounced 101. I,. T.Mashburn, E. A. Boyse, H. A . Campbell, and L. J. Old. I’roc. Soc. E x p l l . Biol. M e t / . 124, 568 (1967). 102. V. Rilry. J. L. Lovelcss, M . A . Fitzmuwice. and W. M . Siler, Lije Sci. 4, 487, 1965; V. Riley, Methods Cancer Res. 4, 493 (1968). 103. A. L. Notkins and C. Srheele, J. Nntl. Cnncer Inst. 33, 741 (1964). 104. B. W. J. Mahv. K . E. K . Bowsrn, and C. W. Pam, J . E z p t / . M e d . 125, 277 (1967). 105. V . Riley, Nulure 220, 1245 (1968).

5.

L-ASPARAGINASE

121

effect on the half-life time of E . coli asparaginase, increasing it in one set of experiments, for example, from the control value of 2.6 hr in normal mice to 19.7 hr (59). The status of the extensive clinical trials of L-asparaginase against a number of forms of cancer, especially acute lymphatic leukemia, has been thoroughly treated in several recent reviews (106-109) and will not be dealt with here.

106. R. H. Adumson and S. Fabro, Crrncer Chemotherapy R e p . 52, 617 (1968). 107. R. E. Handschumacher, Ann. R e v . M e d . 21, 433 (1970) ; Ann. R e v . Pharmacol. 10, 421 (1970). 108. P. Laboureur, Pathol. Biol. Semaiiie H o p . 17, 885 (1969). 109. H. Marquardt, ArzneimitteZ-Forsch. 18, 1380 (1968).

This Page Intentionally Left Blank

Enzymology of Pyrrolidone Carboxvlic Acid MARIAN ORLOWSKI

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 . . . . . . . . . . , . A. D-Glutamic Acid Cyclotransferase . . . . . . B. Glutamine Syntbetase and y-Glutamylcysteine Synthetase C. L-Glutamic Acid Cyclotransferase (L-Glutamic Acid . . . . . . . . . . . Dehydrase) D. Conversion of L-Glutamate to Pyrrolidone Carboxylate by Rat Liver Nuclear Preparations . . . . . . VI. Enzymic Formation of Pyrrolidone Carboxylic Acid from Glutamine and Glutaminyl Peptides . . . . . . A. L-Glutamine Cyclotransferase . . . . . . . B. y-Glutamyl Transpeptidase and . . . . . . . y-Glutamyl Cyclotransferase VII. Enzymic Formation of Pyrrolidone Carboxylic Acid from y-Glutamyl Amino Acids . . . . . . . . A. y-L-Glutamyl Cyclotransferase (y-Glutamyl Lactamase) . VIII. Enzymic Formation of Derivatives of Pyrrolidone Carboxylic Acid . . . . . . . . . . . . IX. Pyrrolidone Carboxylyl Peptidase . . . . . . . . X. Pyrrolidone Carboxylate Metabolism . . . . . . .

123

124 125 127 130 133 133 136 138 138 139 139 141 142 142 146 147 149

124

M. ORLOWSKI AND A. MEISTER

1. Introduction

Haitinger ( I ) , who seems to have been the first to describe pyrrolidone carboxylic acid (2-pyrrolidone-5-carboxylic acid ; pyroglutamic acid) , reported in 1882 that when he heated glutamic acid a t 180-190" it was converted to a new compound which differed from glutamic acid in having lost a molecule of water; Haitinger named this compound pyroglutamic acid. A decade later, Menozzi and Appiani ( 2 , 5 ) reported similar but more detailed studies and gave the correct structure. Thus, pyrrolidone carboxylic acid has been known for about 90 years as a cyclic compound derived from glutamic acid. It has been found in the free state in many plark and animal tissues and also in peptide linkage as the N-terminal group of a number of peptides and proteins. Its role in metabolism is as yet undefined, and it is not yet certain in a number of instances whether its presence in proteins and peptides can be ascribed to nonenzymic modification of an N-terminal glutamine or glutamate residue during isolation or whether it is present as such in the native molecule. Pyrrolidone carboxylic acid was reported as a product of glutathione metabolism in kidney extracts in 1942 ( 4 ) ; however, the first clearly defined enzymic reaction in which pyrrolidone carboxylic acid is a product was described as recently as 1956 by Connell and Hanes ( 5 ) . These workers described an enzymic activity (y-glutamyl lactamase ; 7-glutamyl cyclotransferase) in liver that catalyzes the conversion of 7-glutamyl amino acids to pyrrolidone carboxylic acid and the corresponding amino acid. Subsequently, the formation of L- and D-pyrrolidone carboxylic acid from the corresponding isomers of glutamic acid was found to be catalyzed by glutamine synthetase in the absence of ammonia (6, 7 ) . Other enzymes that catalyze pyrrolidone carboxylate formation include (1) an activity found in the kidney and liver of a number of animals which catalyzes the conversion of D-glutamate to Dpyrrolidone carboxylate (8),(2)a bacterial enzyme that forms L-pyrrolidone carboxylate from L-glutamate ( 9 ) , and (3) an enzyme found in 1. L. Haitinger, Monatsh. Chem. 3, 228 (1882). 2. A. Menozzi and G. Appiani, Gazz. Chim. Ital. 22, 14 (1892). 3. A. Menozzi and G. Appiani, Gazz. Chim. Ital. 24, 370 (1894). 4. G.E.Woodward and F. E. Reinhart. JBC 145, 471 (1942). 5. G.E. Connell and C. S. Hanes, Nature 177, 377 (1956). 6. P. R. Krishnaswamy, V. Pamiljans, and A. Meister, JBC 235, PC39, (1960). 7. P.R. Krishnaswamy, V. Pamiljans, and A. Meister, JBC 237, 2932 (1962). 8. A. Meister and M . W. Bukenberger, Nature 194, 557 (1962). 9. S. Akita, K. Tanaka, and S. Kinoshita, BBRC 1, 179 (1959).

6.

ENZYMOLOGY OF PYRRQLIDONE CARBOXYLIC ACID

125

papaya latex that forms pyrrolidone carboxylate from free glutamine and glutaminyl peptides (10).The formation of pyrrolidone carboxylate from glutathione in kidney extracts ( 4 ) may now be ascribed to the cornbined action of two enzymes: ( 1 ) y-glutamyl transpeptidase and (2) y-glutamyl cyclotransferase (11-13). Preparations and extracts of nuclei have been reported to catalyze the conversion of L-glutamate to L-pyrrolidone carboxylate, and it appears that this reaction is linked to an energyyielding oxidation reaction (14-16). I n this chapter the enzymic reactions mentioned above will be reviewed. I n addition, consideration will be given to the detection, determination, and natural occurrence of pyrrolidone carboxylic acid, the noneneymic formation of pyrrolidone carboxylic acid, and to other information now available about the metabolism of this compound.

II. Detection and Determination of Pyrrolidone Carboxylic Acid

Ellfolk and Synge (17) have reviewed methods that have been used for the identification and isolation of pyrrolidone carboxylic acid. A number of workers have used extraction with ethyl acetate, chloroform, methanol, or ethanol. A typical procedure is extraction of an aqueous solution (pH 2.4-2.6) with ethyl acetate (4, 18-22) ; after removal of the solvent, the extracted pyrrolidone carboxylic acid is hydrolyzed with hydrochloric acid and quantitatively determined as glutamic acid. This method suffers from the disadvantage that other conjugated forms of glutamic acid and indeed other conjugated amino acids may be extracted into the solvent (17); furthermore, pyrrolidone carboxylic acid may be formed from glutamine during extraction ( 2 1 , 23). The separation of pyrrolidone carboxylic acid from other organic 10. M. Messer, Nature 197, 1299 (1963). 11. E. E. Cliffe and S. G. Waley, BJ 69, 649 (1958). 12. E. E. Cliffe and S. G. Waley, BJ 79, 118 (1961). 13. M. Orlowski, P. G. Richman, and A. Meister, Biochemistry 8, 1048 (1969). 14. T. Niwaguchi, N. Motohashi, and H. J. Strecker, Biochem. 2. 342, 469 (1965). 15. T. Niwaguchi, N. Motohashi, and H. J. Strecker, BBA 82, 635 (1964). 16. T. Niwaguchi and H. J. Strecker, BBRC 16, 535 (1964). 17. N. Ellfolk and R. L. M. Synge, BJ 59, 523 (1955). 18. H. Wilson and R. K. Cannan, JBC 119, 309 (1937). 19. H. G. Bray, S. P. James, I. M. Raffan, and W. V. Thorpe, BP 44, 625 (1949). 20. G. W. Pucher and H. B. Vickery, I n d . Eng. Chem., Anal. Ed. 12, 27 (1940). 21. F. Leuthardt, 2. Physiol. Chem . 265, 1 (1940). 22. P. J. Fodor, A. Miller, and H. Waelsch, JBC 202, 551 (1953). 23. S. Ratner, JBC 152, 559 (1944).

126

M. ORLOWSKI AND A. MEISTER

acids (including glutamic acid) can be achieved by paper chromatography in a variety of solvent mixtures (5-7, 64-27). High voltage paper electrophoresis has also been used for the separation of pyrrolidone carboxylic acid from amino acids (6, 7 , 28, 2 9 ) . The use of “C-labeled substrates (e.g., glutamate, glutamine, y-glutamyl amino acids) facilitates the quantitative study of enzymic reactions. Pyrrolidone carboxylic acid does not give a positive ninhydrin reaction but may be detected on paper chromatograms by application of indicators such as bromcresol green or bromphenol blue. A more sensitive procedure was introduced by Ellfolk and Synge ( l 7 ) ,who used the chlorine-starch-iodide reaction of Rydon and Smith (30)to render pyrrolidone carboxylic acid visible on paper chromatograms. This procedure or one of its modifications (3134a) is very useful for detection of pyrrolidone carboxylic acid on papers after chromatography or electrophoresis. Pyrrolidone carboxylic acid may be quantitatively determined as the methyl ester by gas-liquid chromatography (35); this procedure distinguishes pyrrolidone carboxylic acid, from pyrrolidone, a-methylpyrrolidone carboxylic acid, and piperidone carboxylic acid. The gas chromatographic procedure has been used for determination of D-glutamic acid cyclotransferase activity (35) and for identification of pyrrolidone carboxylic acid in human urine ( 3 6 ) .Zamir and Lichtenstein (37) determined pyrrolidone carboxylic acid colorimetrically after heating samples with hydroxylamine followed by treatment with the ferric chloride reagent of Lipmann and Tuttle (38). Ramakrishna and Krishnaswamy (39)used this reaction to render pyrrolidone carboxylic acid visible on paper chromatograms ; they cut out 24. L. Levintow, A. Meister, G. H. Hogeboorn, and E. L. Kuff, JACS 77, 5304 (1955). 25. W. J. Le Quesne and G. T. Young, J. Chem. Soc. p. 594 (1952). 26. J . B. Stark, A. E. Goodban, and H. S. Owens, Anal. Chem. 23, 413 (1951). 27. A. Meister, M. W. Bukenberger, and M. Strassburger, Biochem. 2. 338, 217 (1963). 28. D. Gross, Nature 178, 29 (1956). 29. G. E. Connell and A . Szewczuk, Clin. Chim. Acla 17, 423 (1967). 30. H. N. Rydon and P. W. G. Smith, Nature 169, 922 (1952). 31. F. Reindel and W. Hoppe, Naturwissenschajtsn 40, 221 (1953); Chem. Ber. 87, 1103 (1954). 32. A. Meister, I,. Levintow, R. E. Greenfield, and P. A. Abendschein, JBC 215, 441 (1955). 33. S. C. Pan and J. D. Dutcher, Anal. Chem. 28, 836 (1956). 34. D. P. Schwartz and M. J. Pallansch, Anal. Chem. 30, 219 (1958). 34ii. It. H. Mazur, B. W. Ellis. and P. S. Cammarata, JBC 237, 1619 (1962). 35. P. Polgar and A. Meister, AM^. Biochem. 12, 338 (1965). 36. R. Tham, L. Nystrom, and B. Holmstedt, Biochem. Pharmacol. 17, 1735 (1968). 37. A. Zarnir and N. Lichtenstein, A n d . Chim. Acta 12, 577 (1955). 38. F. Lipmann and L. C. Tuttle, JBC 159, 21 (1945). 39. M. Rarnakrishna and P. R. Krishnaswamy, Anal. Biochem 19, 338 (1967).

6.

ENZYMOLOGY OF PYRROLIDONE CARBOXYLIC ACID

127

the spots containing y-glutamyl hydroxamate and after elution with ethanol estimated the hydroxamate colorimetrically. The procedure suffers from the disadvantage that it also yields color with glutamine, asparagine, and glutathione; furthermore, the low sensitivity of this test restricts its usefulness. It is known that peptides and proteins exhibit a higher absorbance a t wavelengths below 240 nm than the component amino acids (40, 41) and that the absorbance a t 205 nm of peptides and proteins is approximately proportional to the number of peptide bonds. Pyrrolidone carboxylic acid, which has an internal peptide bond, exhibits this type of absorbance, which may be used under appropriate conditions for its quantitative determination. Thus, Orlowski et al. (13) followed the activity of y-glutamyl cyclotransferase by this procedure; after incubation of the enzyme with 7-glutamyl amino acids, the reaction mixture was passed through a small column of Dowex 50 (H’) in order to remove interfering substances, and the absorbance of the eluate a t 205 nm was then determined. A number of attempts have been made to determine pyrrolidone carboxylic acid after reduction with sodium in alcohol, zinc and HCl, Na2S204,and other reducing agents; in general, these were not successful (42). Recently, the reduction of an N-terminal pyrrolidone carboxylic acid residue of proteins with diborane in tetrahydrofuran or tetramethyl urea was attempted with some success; the N-terminal pyrrolidone carboxylic acid residue may thus be identified by its conversion to proline ( 4 3 ) . Special procedures (44) are required for the demonstration of Nterminal pyrrolidone carboxylic acid residues in peptides and proteins; frequently, pyrrolidone carboxylyl-peptides can be detected by the procedure of Rydon and Smith (SO). Discovery of enzymes capable of cleaving pyrrolidone carboxylic acid from the N-terminus of peptide chains should facilitate studies of this type (45, 46) (see below).

111. Natural Occurrence of Pyrrolidone Carboxylic Acid

Pyrrolidone carboxylic acid has been found in the free state in a wide variety of vegetables, fruits, and grasses. Thus, it has been found in 40. 41. 42. 43. 44. 45. 46.

J. S. Ham and J. R. Platt, J . Chem. Phys. 20, 335 (1952). L. J. Saidel, ABB 54, 184 (1955). G. 0. Schlutz, Biochem. 2. 324, 295 (1953). S. Takahashi and L. A . Cohen, Biochemistry 8, 864 (1969). B. Blomback, “Methods in Enzymology,” Vol. 11, p. 398, 1967. R. F. Doolittle and R. W. Armentrout, Biochemistry 7, 516 (1968). R. U‘.Armentrout, BBA 191, 756 (1969).

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M. ORLOWSKI AND A. MEISTER

TABLE I N-TERMINAL PYRROLIDONE CARBOXYLYL (PYR-)RESIDUES OF PEPTIDES AND PROTEINS No. 1 2 3 4 5

6

7 8 9

10 11 12

Peptide or protein

Sequence

Eisenine Fastigiatine Eledoisin Physalaemin Thyrotropin releasing hormone? Fibrinopeptides B Man Bovine Reindeer Elk Muntjak Mule deer Cape buffalo Water buffalo Gastrin Peptides from snake venoms Heavy chain of human immunoglobin (Eu) Heavy chain of pathological human immunoglobin IgG Heavy chains of .abbit IgG

Pyr-Glu-AlaOH Pyr-Glu-GlnOH Pyr-Pro-SerPyr-Ala-AspPyr-His-Pro-NH2

A-Type Bence Jones proteins Type BO Type Ha Mouse A chains a,acid glycoprotein of human plasma a2chain of rat collagen &Galactosidase from E. wli

Reference

Pyr-G1y-ValPyr-Phe-ProPyr-Leu-AlaPyr-His-SerPyr-His-SerPyr-His-LeuPyr-Phe-ProPyr-Phe-ProPyr-Gly-ProPyr-Gln-Try-OH Pyr-Asn-Try -OH Pyr-Val-GlnPyr-Val-ThrSeveral Pyrpeptides Pyr-Ser-AlaPyr-Ser-ValPyr-Ala-ValPyr-Pro-LeuRyr-TVr-SerPyr-?

Italian rye grass that has been stored a t -20" ( 1 7 ) . However, it seems to be absent in extracts of fresh grass but appears after drying and in silage (17, 4 7 ) . It is of interest that while fresh tomato juice does not contain appreciab 'e quantities of pyrrolidone carboxylic acid, this compound represents about 30% of the total organic acid in stored tomato juices (48). These and other reports which have appeared in the literature 47. H. T. Macphenon and J. S. Slater, BJ 71, 654 (1959). 48. A. C. Rice and C. S. Pederson, Food Res. 19, 106 (1954).

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ENZYMOLOGY OF PYRROLIWNE CARBOXYLIC ACID

129

suggest that most plant materials contain little if any free pyrrolidone carboxylic acid ; however, the amount increases, often considerably, during processing or storage probably a t the expense of glutamine, 7glutamyl amino acids, or y-glutamyl peptides. Pyrrolidone carboxylic acid is formed nonenzymically from glutamine during prolonged incubation of proteins with proteolytic enzymes, and in proteins in which the N-terminal amino acid is glutamine, cyclization may precede cleavage of the residue leading to the formation of a pyrrolidone carboxylic-peptide. Sanger et al. (4.9) were able to demonstrate that a pyrrolidone carboxylic acid residue was formed from a glutamine residue in positions 5 and 15 of the A chain of insulin; thus, they obtained two peptides having the same amino acid sequence but differing only in the N-terminal component; one peptide had a pyrrolidone carboxylic acid residue and the other a glutamine residue. Similar observations were made in studies on ribonuclease (50).Pyrrolidone carboxylic acid formation has also been observed during enzymic hydrolysis of enolase, albumin, casein, egg albumin, and other proteins (51-53).In a number of instances it has been difficult to determine whether the Nterminal pyrrolidone carboxylic acid residue formed during isolation or whether it is present in the native peptide or protein. The subject has been extensively reviewed recently (44).A list of some of the peptides and proteins which have been found to contain N-terminal pyrrolidone carboxylic acid residues is given in Table I (54-73).The methods 49. F. Sanger, E. 0. P. Thompson, and R. Kitai, BJ 59, 509 (1955). 50. D. G. Smyth, W. H. Stein and S. Moore, JBC 237, 1845 (1962). 51. J. A. Winstead and F. Wold, Biochemistry 3, 791 (1964). 52. K. Narita and J. Ishii, J. Biochem. ( T o k y o ) 52, 367 (1962). 53. S. Aqvist and A. Wretlind, Acta Physiol. Scand. 39, 147 (1957). 54. B. Blomback, M. Blomback, P. Edman, and B. Hessel B B A 115, 371 (1966). 55. E. M. Press, P. J. Piggot, and R. R. Porter, BJ 99, 356 (1966). 56. C. A. Dekker, D. Stone, and J. S. Fruton, JBC 181, 719 (1949). 57. A. Anastasi and V. Erspamer, ABB 101, 56 (1963). 58. V. Erspamer, A. Anastasi, G. Bertaccini, and J. M. Cei, Ezperientiu 20, 489 (1964). 59. J. Boler, F. Ensmann, K. Folken, C. Y. Bowers, and A. V. Schally, BBRC 37, 705 (1969). 60. B. Blomback, M. Blomback, and P. Edman, Acta Chem. Scand. 17, 1184 (1963). 61. B. Blomback and R . F. Doolittle, Acta Chem. Scand. 17, 1816 (1963). 62. B. Blomback and R. F. Doolittle, Acla Chem. Scand. 17, 1819 (1963). 63. G. A. Mross and R. F. Doolittle, ABB 122, 674 (1967). 64. H. Gregory, P. M. Hardy, D. S. Jones, G. W. Kenner, and R. C. Sheppard, Nature 204, 931 (1964). 65. H. Kato, S. Iwanaga, and T. Suzuki, Ezperientia 22, 49 (1966). 66. G. M. Edelman, B. A. Cunningham, W. E. Gall, P. D. Gottlieb, U. Rutishauser, and M. J. Waxdal, Proc. Natl. Acad. Sci. U . S. 63, 78 (1960).

130

M. ORLOWSKI AND A. MEISTEH

employed for the isolation of some of these peptides (e.g., eisenine, fastigiatine, and gastrin) involve extraction with hot or boiling water or water-alcohol mixtures, procedures that might be expected to lead to cyclization of N-terminal glutamine residues. On the other hand, the isolation of fibrinogen (54) and immunoglobin (55) were carried out a t neutral pH and low temperature; it seems unlikely that under these conditions appreciable cyclization of glutaminyl residues occurred and thus that the native protein probably contains N-terminal pyrrolidone carboxylic acid. It remains to be determined whether pyrrolidone carboxylic acid can be incorporated directly into protein or whether this residue is formed by cyclization of a glutaminyl residue on the tRNA, growing peptide, or completed peptide. Some experimental efforts have been made in this direction (74, 7 4 a ) . It is of interest in this connection that the enzyme from papaya latex ( 7 6 ) ,which catalyzes the cyclization of glutamine and glutaminyl peptides also catalyzes the formation of pyrrolidone carboxylyl-tRNA from E. coli glutaminyl-tRNA ( 7 6 ) . It would be of interest to determine whether glutamine cyclotransferase or a similar enzyme with activity toward glutaminyl-tRNA and glutaminyl peptides is present in animal tissues.

IV. Nonenzymic Formation of Pyrrolidone Carboxylic Acid from Glutamic Acid, Glutamine, and Other Compounds

Abderhalden and Kautzch (77) reported that the heating of dry glutamic acid to 180"-200" led to the formation of essentially optically inactive pyrrolidone carboxylic acid. They obtained optically active Lpyrrolidone carboxylic acid by heating L-glutamic acid a t 150"-160" followed by fractional crystallization of the product. These workers also observed that treatment of optically active pyrrolidone carboxylic acid with strong mineral acid led to its conversion to optically active glutamic 67. J. M. Wilkinson, E. M. Press, and R. R. Porter, BJ 100, 303 (1966). 68. F. W. Putnam, T. Shinoda, K. Titani and M. Wikler, Science 157, 1050 (1967). 69. L. Hood, W. R. Gray, and W. J. Dreyer, JMB 22, 179 (1968). 70. E. Appella and R. N. Perham, JMB 33, 963 (1968). 71. T. Ikenaka and K. Schmid, Proc. Soc. EzptZ. Biol. M e d . 120, 749 (1965). 72. A. H. Kang, P. Bornstein. and K . A. Piez, Biochemistry 6, 788 (1967). 73. R. P. Erickson and E. Steers, Jr., BBRC 37, 736 (1969). 74. B. Moav and T. N. Harris. BBRC 29, 773 (1967). 74a. C. Baglioni, BBRC 38, 212 (1970). 75. M. Messer and M. Ottesen, BBA 92, 409 (1964). 76. M. R. Bernfield and L.Nestor, BBRC 33, 843 (1968). 77. E. Abderhalden and K. Kautzch, 2. PhysioZ. Chem. 68, 487 (1910).

6. ENZYMOLOGY OF PYRROLIDONE CARBOXYLIC ACID

131

acid. The marked difference between the melting points of L-pyrrolidone carboxylic acid (160"-161") and D,L-pyrrolidone carboxylic acid (180"183") (78) led to the erroneous identification of the former compound as 3,6-diketopiperazine-2,5-dipropionicacid (79). Wilson and Cannan (18) reported detailed observations on the equilibrium and velocity constants in the glutamic acid-pyrrolidone carboxylic acid system in dilute aqueous solution. They found that the conversion of glutamic acid to pyrrolidone carboxylic acid follows the equation for a reversible first-order reaction. The equilibrium constant and the rate a t which the equilibrium is achieved depend on the pH of the solution and the temperature. I n neutral solutions, the equilibrium favors almost complete conversion of glutamic acid to pyrrolidone carboxylic acid; however, the rate of the reaction is very slow and thus only 1% conversion occurs after 2-3 hr a t 100".I n weakly acid (pH 4) and alkaline (pH 10) solutions, the conversion of glutamic acid to pyrrolidone carboxylic acid is much faster and about 98% conversion occurs in less than 50 hr. In strong acid (2 N HCl) and base (0.5 N NaOH) the conversion of pyrrolidone carboxylic acid to glutamic acid proceeds rapidly and virtually to completion. Other studies have shown that the conversion of glutamic acid to pyrrolidone carboxylic acid can be carried out within 2 hr a t 142" with little alteration of optical rotation (80). Using the data of Wilson and Cannan (It?), Cleaves (81) was able to show that the rate of formation of pyrrolidone carboxylic acid from glutamic acid in aqueous solution depends directly on the concentration of the ionic species of glutamic acid in solution. Thus, the reactive species are (I), (11),and (IV), while (111) is relatively unreactive. Protonation of the amino group and dissociation of the 7-carboxyl group thus makes these groups less reactive ; carboxylate ion resonance apparently hinders nucleophilic attack by the amino nitrogen. The stabilizing effect of the carboxylate ion resonance is also evident YOOH

78. 79. SO. 81.

y

2

y

2

COOH

yOO-

yo-

7Hz

y i 2

7%

I

C HNH:

7% C HNH:

COOH

COO-

COO-

coo-

(1)

(11)

(UI)

(IV)

I

7% CHNH:

A . F. Beecham, JACS 76, 4613 (1954). J. A . King and F. H. McMillan, JACS 74, 2859 (1952). A. C. Kibrick. JBC 174, 845 (1948). D. W. Cleaves, JBC 186, 163 (1950).

7% CHNH,

132

M. ORLOWSKI AND A. MEISTER

from the rapid cyclization of glutamic acid y-derivatives in which this effect is weakened or abolished. For example, glutamine, 7-esters of glutamic acid, and y-glutamyl peptides are known to cyclize much more rapidly than glutamic acid. Chibnall and Westall (82) found that when glutamine was boiled in solution (p H 3-10) substantial amounts of ammonia form while a t the same time the amount of amino nitrogen decreases. Thus, the main product of "hydrolysis" of glutamine under these conditions is pyrrolidone carboxylic acid (83).Archibald (84) has called attention to the fact that when glutamine was heated at 186" for 30 sec it is quantitatively converted to ammonium pyrrolidone carboxylate. When solutions of glutamine in phosphate buffer (pH 6.5) are placed a t loo", there is 99% conversion to pyrrolidone carboxylate in 90 min; in contrast, cyclization of glutamate under these conditions is very slow. Hamilton was the first to observe that phosphate exerted an accelerating effect on the cyclization of glutamine ( 8 5 ) . These studies were confirmed and extended by Gilbert et a2. (86) who showed that arsenate and carbonate exhibit a similar effect. The nonenzymic cyclization of glutamine is substantial even a t 37",10% being converted to pyrrolidone carboxylic acid after 24 hr (87). Melville has shown that peptides containing glutamine in the N-terminal position exhibit a lability similar to that of glutamine (88). When the amino group is substituted, considerable stabilization is observed ; thus, C-terminal glutamine and glutamine residues within the peptide chain are much more stable than is free glutamine (82, 89). 7-Glutamyl hydroxamate is much more labile than glutamine ; thus, it is virtually completely converted to pyrrolidone carboxylic acid a t pH 7 after heating a t 100" for 7.5 min; less than 10% of glutamine is cyclized under these conditions ( 2 4 ) . a,y-Diesters of glutamic acid (90) are also susceptible to rapid cyclization while a-esters are much more stable (91). Attempts to prepare glutamine from y-methylglutamate or 7-ethylglutamate by ammonolysis fail because cyclization proceeds very rapidly, so rapidly indeed that ammonolysis is not effective. Thus, treat82. A. C. Chibnall and R. G. Westall, BJ 28, 122 (1932). 83. H. B. Vickery, G. W. Pucher, H. E. Clark, A. C. Chibnall, and R. G. Westall, BJ 29, 2710 (1935). 84. R . M. Archibald, Chem. R e v . 37, 161 (1945). 85. P. B. Hamilton, JBC 158, 375 (1945). 86. J. B. Gilbert, V. E. Price, and J. P. Greenstein, JBC 180, 209 (1949). 87. G. L. Tritsch and G. E. Moore, Exptl. Cell Res. 28, 360 (1962). 88. J. Melville, BJ 29, 179 (1935). 89. H. Thierfelder and E. von Cramm, 2. Physiol. Chem. 105, 58 (1919). 90. E. Abderhalden and A. Weil, 2. Physiol. Chem. 74, 445 (1911). 91. A. Neuberger, BJ 30, 2085 (1936).

6.

ENZYMOLOGY OF PYRROLIWNE CARBOXYLIC ACID

133

ment of y-ethylglutamate with aqueous ammonia leads to almost quantitative conversion to pyrrolidone carboxylate (92-94). y-Thioesters of glutamic acid decompose rapidly a t neutral or slightly acid p H to yield pyrrolidone carboxylic acid (95), and it is of interest that phosphate enhances this reaction. Hopkins (96) found that heating of aqueous solutions of glutathione led to formation of pyrrolidone carboxylic acid; cleavage is complete in 120 hr a t 62" (97). Other y-glutamyl peptides exhibit similar behavior (25). However, the nonenzymic cyclization of y-glutamyl amino acids and related compounds proceeds more slowly than that of glutamine; thus, 7-glutamylmethyl amide, y-glutamyldimethylamide, y-glutamylethylamide, and y-glutamylglycine were not significantly affected by incubation in 0.5 M potassium phosphate buffer a t p H 7.8 after 3 days, while under the same conditions the deamidation of glutamine, a-methylglutamine, homoglutamine, and y-methylglutamine had gone virtually to completion (98). It is of interest that the deamidation of glutamine and of a-methylglutamine occur a t approximately the same rates while y-methylglutamine is deamidated considerably more rapidly than glutamine (98). Similarly, y-methyleneglutamine cyclizes more rapidly than glutamine (99). The same product was obtained in the cyclization of 7-methyleneglutamic acid ; this reaction occurs more rapidly than that of glutamic acid. The cyclization of y-hydroxy-ymethylglutamic acid (99) and of p-hydroxyglutamic acid (100) has also been described.

V. Enzymic Formation of Pyrrolidone Carboxylic Acid from Glutamic Acid A.

D-GLUTAMIC ACID CYCLOTRANSFERASE

The discovery of D-glutamic acid cyclotransferase was foreshadowed by the observations of Ratner (23) who fed rats D,L-glutamate labeled with I5N in the amino group and deuterium attached to the a- and pM. Bergman and L. Zervas, 2.Physiol. Chem. 221, 51 (1933). D. Coleman, J. Chem. SOC.p. 2294 (1951). A. F. Beecham, JACS 76, 4615 (1954). H. Sachs and H. Waelsch, ABB 69, 422 (1957). F. G. Hopkins, JBC 84, 269 (1929). E. C. Kendall, H. L. Mason, and B. F. McKenzie, JBC 88, 409 (1930). 98. A. Meister, JBC 210, 17 (1954). 99. J. Blake and L. Fowden, BJ 92, 136 (1964). 100. C. R. Harington and S. S. Randall, BJ 25, 1917 (1931).

92. 93. 94. 95. 96. 97.

134

M. ORLOWSKI AND A. MEISTER

carbon atoms. The animals excreted more 15N in the urine than could be accounted for by urea and ammonia; the extra 15N was found in the form of D-pyrrolidone carboxylic acid, which contained the same concentrations of isotopes as that of the glutamic acid fed. Ratner recovered 73% of the administered D-glutamic acid as urinary D-pyrrolidone carboxylic acid, and since the concentration of deuterium was unchanged she concluded that no reaction involving the a- and phydrogen atoms of D-glutamic acid had occurred. These findings were subsequently confirmed by Wilson and Koeppe (101) who found that injection of l'C-~-glutamate to rats led to the excretion of more than 50% of the radioactivity as D-pyrrolidone carboxylic acid. These authors also found that incubation of D-glutamate with slices of rat liver and kidney led to the formation of D-pyrrolidone carboxylate. The enzyme responsible for the conversion of D-glutamic acid to D-pyrrolidone carboxylic acid was subsequently demonstrated in homogenates of kidney and liver of the mouse, rat, and man (8). Somewhat lower levels of activity were subsequently found in the kidney and liver of the guinea pig and rabbit and in rat spleen, skeletal muscle, brain, pancreas, and testes (102). The enzyme was purified approximately 50-fold from mouse kidney, which appears to be the richest source of this enzyme. Studies on rat and mouse liver and kidney indicated that about two-thirds of the initial activity remained in the supernatant solution when homogenates of these tissues were centrifuged a t 100,000 g for 2 hr. Purification was accomplished by centrifugation of the homogenates followed by ammonium sulfate precipitation (3&60% of saturation) ; the enzyme was then passed through a column of Sephadex G-50and lyophilized. A moderate increase in specific activity was achieved by absorption of impurities on alumina C-y gel. The enzyme requires either manganese or magnesium ions for activity, but no other cofactor requirements were found. With 0.033 M concentrations of metal ions, the activity was more than three times higher with manganese than with magnesium ions. Dialysis of the enzyme against tris-HC1 buffer (pH 7.5) or water a t 0" led to inactivation and attempts a t reactivation were unsuccessful. Optimal activity was observed in the pH range 7.8-8.3 (tris-HC1 buffers). The apparent K , value for Dglutamate is 1.5 x 1CP M . The reaction is reversible and the equilibrium was approached from both directions in the presence of magnesium ions; an equilibrium value of 94% of D-glutamate was converted to D-pyrrolidone carboxylate a t (pH 8.3). The equilibrium constant, K,,= pyrrolidone carboxylate/glutamate = 16. 101. W. E. Wilson and R. E. Koeppe, JBC 236, 365 (1961). 102. A . Meister, M. W. Bukenberger, and M . Strassburger, Baochem. 2. 338, 217 (1963).

6.

135

ENZYMOLOGY OF PYRROLIDONE CARBOXYLIC ACID

The product of the enzymic reaction was isolated in crystalline form and identified as D-pyrrolidone carboxylic acid. The enzyme does not act on L-glutamic acid nor does it catalyze the synthesis of L- or Dglutamine or of the corresponding 7-glutamyl hydroxamates. Studies on the specificity of the enzyme indicated that a number of racemic glutamic acid derivatives are substrates. Thus, as indicated in Table 11, various methyl and hydroxy derivatives of glutamic acid are attacked; with the racemic substrates 50% was utilized. It is of interest that although D-e-aminoadipic acid is not a substrate, both diastereoisomers of D,L-a-aminotricarballylate are attacked. The ability of D-glutamic acid cyclotransferase to act on a-aminotricarballylate has made it possible to obtain enzymic evidence for the configuration of the D-isocitric acid of the Krebs tricarboxylic acid cycle (103). The available data suggest that the amount of D-glutamic acid cyclotransferase in mammalian kidney and liver is sufficient to explain the appearance of D-pyrrolidone carboxylate in the urine of animals that TABLE I1 SPECIFICITY OF D-GLUTAMIC ACID CYCMTRANSFERASEO

Amino acid D-Glutamatd D,LGlutamatee L-Glutamate

,

D b y -Hy droxygluta-

mate (isomer A)" D,ty-Hydroxyglutamate (isomer B) D,tp-Hydroxyglutamate (isomer B) D, bfl-MethylglutaD,ba-Methylglutamate Da-Methylglutamate

Disappearance of amino acid (pmole/hr)

3.8 3.8 0 2.6

0.80

Amino acid D-y-Methylglu tamatd D,ty-Methylglutamatec D,tAllo-a-aminotricarballylatdvc D,bcu-Aminotncarballylate**C tGlutamine

Disappearance of amino acid (pmolelhr)

3.0 2.9 3.6 1.8

0 0

1.8 3.2

D- Aspartate

0

0.80 0.80

D-a-Aminadipate D-Homoglutamine

0 0

a The reaction mixtures contained initially mouse kidney enzyme (0.5 mg), b o r D-amino acids (5 pmoles), D,tamino acids (10 pmoles), MgC12 (4 pmoles), and 2-amino-2-(hydroxymethyl)-1,3-propanediol-HCl buffer (33 rmoles; pH 8.3) in a final volume of 0.2 ml; 37";analysis by procedure C. * The product was identified by paper chromatography [from Meister et al. (IOS)]. c On prolonged incubation (2-6 hr) disappearance of amino acid was close to but did not exceed 5070 of the total amino acid added.

103. A. Meister and M. Strassburger, Nature 200, 259 (1963).

136

M. ORLOWSKI AND

A.

MEISTER

have been fed D-glutamic acid. Substantial amounts of D-pyrrolidone carboxylic acid have been found in human urine (10%);it is not known whether this arises from D-glutamic acid of dietary origin or whether it is produced by bacterial metabolism in the intestine. It might conceivably arise from dietary D-pyrrolidone carboxylic acid (104). The finding of D-pyrrolidone carboxylic acid and D-glutamic acid in the urine after feeding of human tumors to dogs and rats (105,1 0 6 ) was cited as evidence that tumors contain D-glutamic acid; this subject has been reviewed by Miller (107).The physiological role of D-glutamic acid cyclotransferase may be to detoxify D-glutamate formed by the intestinal flora or present in the diet. D-Glutamic acid is one of the most abundant bacterial D-amino acids and it is a very poor substrate for D-amino acid oxidase. The enzymic cyclization of D-glutamate may be considered as a special case of an intramolecular acylation reaction, similar to the physiological acylation of benzoate (as hippuric acid) and phenylacetate (as phenylaceturic acid and phenylacetylglutamine) .

B. GLUTAMINE SYNTHETASE AND 7-GLUTAMYLCYSTEINE SYNTHETASE The enzymic synthesis of glutamine takes place according to the following reaction: ena. M g * +

Glutamate

+ ATP + NH, 7’ glutamine + ADP + Pi

Glutamine synthetase (from brain and peas) catalyzes the synthesis of L-glutamine and D-glutamine from the respective optical isomers of glutamic acid [for reviews, see Meister (108-1IO)].When glutamine synthetase is incubated with ATP, magnesium ions, and either L- or Dglutamate (in the absence of ammonia) the corresponding isomer of pyrrolidone carboxylic acid is formed : Glutamate

+ ATP

en5, Mg’+ A

pyrrolidone carboxylate

+ ADP + Pi

The formation of pyrrolidone carboxylic acid under these conditions has been interpreted to indicate the enzymic formation of an activated glutamate intermediate. The available evidence supports the view that glutamate and ATP interact on the active site of the enzyme to yield en104. E. Abderhalden and R. Hanslian, 2.Physiol. Chem. 81, 228 (1912). 105. F. Kogl, T. J. Barendregt, and A. J. Klein, Nature 162, 732 (1948). 106. G. Hillman, A. Hillman-Elies, and F. Methfessel, 2. Naturforsch, (1954). 107. J. A. Miller, Cancer Res. 10, 65 (1950). 10s. A. Meister, “The Enzymes,” Vol 6, p. 443, 1962. 109. A. Meister, Harvey Lectures 63, 139 (1969). 110: A. Meister, Advan. Enzymol. 31, 183 (1968).

Qb,

660

6.

ENZYMOLOGY OF PYRROLIDONE CARBOXYLIC ACID

137

zyme-bound 7-glutamyl phosphate. In the absence of ammonia, the enzyme-bound acyl phosphate is released from the enzyme as pyrrolidone carboxylic acid. Whether cyclization occurs on the enzyme or after dissociation of the acyl phosphate from the enzyme is not yet certain. It is of interest that the rate of enzymic formation of pyrrolidone carboxylic acid from L-glutamate is approximately the same as that of D-glutamate; however, the cyclization reactions occur much more slowly than that of the synthesis of glutamine catalyzed by the same enzyme. Similar cyclization reactions take place with the optical isomers of a-aminoadipic acid. I n this reaction, piperidone carboxylic acid is formed (6, 7, 111). Although the cyclization reactions catalyzed by glutamine synthetase are of interest in relation to the mechanism of action of this enzyme, it seems unlikely that this relatively slow reaction can be of physiological significance. Purified preparations of y-glutamylcysteine synthetase can also catalyze the formation of pyrrolidone carboxylic acid ( I l l a , b ). This enzyme catalyzes the synthesis of y-glutamylcysteine according to the following reaction: L-Glutamate

+ A T P + L-cysteine enz, Mgz, ---+

L-y-glutamyl-L-cysteine

+ A D P + Pi

When L-cysteine is replaced by L-a-aminobutyrate, L-y-glutamyl-L-aaminobutyrate is formed. The enzyme does not catalyze the synthesis of glutamine nor does glutamine synthetase catalyze the formation of y-glutamyl amino acids. Nevertheless, the reactions catalyzed by these enzymes are analogous and appear to involve activation of the y-carboxyl group of glutamate. When y-glutamylcysteine synthetase purified from rat kidney was incubated with L-glutamate, ATP, and magnesium ions in the absence of an amino acid acceptor pyrrolidone carboxylic acid was formed. The formation of pyrrolidone carboxylate requires magnesium ions and ATP and is associated with the cleavage of ATP to stoichiometric quantities of ADP and inorganic phosphate. The enzyme also catalyzes the formation of D-y-glutamyl-L-a-aminobutyrate; however, even in the presence of L-0-aminobutyrate, cyclization of D-glutamate takes place. It is of interest that r-glutamylcysteine synthetase catalyzes the cyclization of D-glutamate about five times more rapidly than Lglutamate. Presumably the cyclization of D-glutamate and L-glutamate catalyzed by this enzyme reflects, as in the case of the analogous reactions catalyzed by glutamine synthetase, activation of the y-carboxyl group of glutamate, i.e., formation of 7-glutamyl phosphate. 111. V. P. Wellner, M. Zoukis, and A. Meister, Biochemistry 5, 3509 (1966). l l l a . M.Orlowski and A. Meister, Biochemistry 10, 372 (1971). l l l b . M.Orlowski and A. Meister, unpublished data (1970).

M. ORLOWSKI AND A. MEISTER

C. L-GLUTAMIC ACID CYCLOTRANSFERASE (L-GLUTAMIC ACID DEHYDRASE) Akita e t al. (9,112) found L-glutamic acid cyclotransferase in a variant of Pseudomonas cruciviae (var. ovalis). This organism utilizes peptone, L-glutamate, L-pyrrolidone carboxylate, and intermediates of the tricarboxylic acid cycle ; while C1-carboxylic acids are good carbon sources, glucose, sucrose, acetate, and ribose are not utilized for growth. The enzyme was purified from lyophilized cells by a procedure which involved sonication of the cells in 0.066 M potassium phosphate buffer (pH 8) followed by centrifugation. After differential heat denaturation of impurities, the enzyme was fractionated with ammonium sulfate and then further purified by acetone fractionation a t -20" and lyophilized. This procedure gave about a 20-fold increase in specific activity. The purified enzyme was reported to exhibit a single component on electrophoresis in phosphate buffer a t pH 8. The enzyme is not active with D-glutamate. At p H 8 and 50") 98.2% of the added L-glutamate was converted to pyrrolidone carboxylic acid a t equilib-rium. The reaction was found to be reversible. The maximal rate of reaction occurred a t 50". The enzyme exhibited a pH optimum a t 8.0 in potassium phosphate buffers. No cofactor requirements were reported. Activity was inhibited by ethylenediaminetetraacetate (EDTA) , sodium sulfide, and L-cysteine. The apparent K , value for L-glutamate was 1.5-1.6 x 10-1 M . It is of interest that the K , value for D-glutamate with D-glutamic acid cyclotransferase (see above) is also relatively high. L-Glutamic acid cyclotransferase was reported to be inactive toward N-acetyl-L-glutamate, L-glutamine, L-glutamic acid y-ethyl ester, D,L-aspartic acid, 7-aminobutyric acid, L-ornithine, L-lysine, glycylglycine, a-aminoadipic acid, diaminopimelic acid, and glutathione.

D. CONVERSION OF L-GLUTAMATE TO PYRROLIDONE CAREOXYLATE BY RAT LIVERNUCLEAR PREPARATIONS Niwaguchi et al. (14-16) reported that when rat liver homogenates were incubated in a system containing '*C-~-glutarnate,glucose, D P N , adenosine triphosphate (ATP), magnesium ions, cytochrome c, and fumarate, an acidic compound was formed which could be identified as pyrrolidone carboxylic acid by infrared spectroscopy, electrophoresis, 112. S. Akita. K. Tanaka, and S. Kinoshita, Koso Kagaku Shimpoziumu 15, 130 (1961).

6.

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ENZYMOLOGY OF PYRROLIDONE CARBOXYLIC ACID

paper chromatography, and melting point determination. About 70% of the initial activity of the homogenate was found to reside in the nuclear fraction while the supernatant fractions were essentially devoid of activity. It was subsequently found that pyruvate or intermediates of the tricarboxylic acid cycle plus ATP and magnesium ions were able to substitute for the multicomponent system described above. Furthermore, a soluble extract of the nuclear fraction was capable of catalyzing the reaction. The formation of pyrrolidone carboxylic acid required oxygen and was inhibited by compounds that inhibit A” synthesis, e.g., cyanide, areenate, 2,4-dinitrophenol, aside, Antimycin A, dicumarol, fluoroacetate, and Amytal. The observation that ATP plus an ATP-regenerating system did not substitute for a carbohydrate subtrate suggested that formation of pyrrolidone carboxylic acid is coupled to an energy-yielding oxidation reaction. The mechanism of pyrrolidone carboxylic acid formation in this system is not yet clear. The possibility that the reaction involves the synthesis of a y-glutamyl amino acid catalyzed by y-glutamyl cysteine synthetase followed by cyclization catalyzed by 7-glutamyl cyclotransferase (although not excluded) seems unlikely since 7-glutamyl cyclotransferase has been found almost exclusively in the supernatant fractions of cells. Furthermore, the nuclear preparations studied did not catalyze the synthesis of glutathione or of 7-glutamyl cysteine. Niwaguchi et al. (14) considered the possibility that a “high energy” intermediate derived from glutamate is formed. Further work is necessary to elucidate the mechanism and physiological significance of this interesting reaction. VI. Enzymic Formation of Pyrrolidone Carboxylic Acid from Glutamine and Glutaminyl Peptides

A. L-GLUTAMINE CYCLOTRANSFERASE This enzyme, which was discovered in papaya latex by Messer ( l o ) , catalyzes the conversion of glutamine and glutaminyl peptides to pyrrolidone carboxylic acid or pyrrolidone carboxylyl peptides, respectively : CON& I

f: H*

7%

KC-CH, 1

C-NHCHCONH.. I

6

1

O&,N,CHCNHCHCONH.

CHNH,

R

.

H

II O

I R

-- +

NH,

140

M. ORLOWSKI AND A. MEISTER

Damodaran and Ananta-Narayanan (113) observed that the rate of formation of ammonia from protein amide groups during hydrolysis by crude preparations of papain was considerably higher than when other proteolytic enzymes were used for hydrolysis. This observation was explained by the discovery (10, 75) that crude papain contains L-glutamine cyclotransferase. Neither chymopapain nor papain catalyzes this cyclization reaction ; indeed, neither of these enzymes liberates ammonia when incubated with wheat gluten. It is known that treatment of wheat gluten with crude papain abolishes the toxic effects of the gluten for patients with celiac disease (114, 115). It has been suggested that the toxicity of gluten is related to the presence of a peptide possessing an N-terminal glutaminyl residue ; presumably this is cleaved by an intestinal peptidase which is lacking in patients with celiac disease. The purification of glutamine cyclotransferase from papaya latex has been carried out by Messer and Ottesen (116). A batch procedure was used for the removal of impurities by passage of a papaya latex extract through a thin layer of carboxymethyl-Sephadex ; the active protein was separated by selective elution. Additional purification was achieved by chromatography on a column of carboxymethyl-Sephadex and by gel filtration on Sephadex G-100. The purified enzyme was homogeneous by the criteria of paper electrophoresis, ultracentrifugation, and gel filtration on Sephadex G-100 columns and chromatography on carboxymethyi- or DEAE-Sephadex. The electrophoretic behavior of the enzyme indicates that it is a basic protein with an isoelectric point near 9.0. The sedimentation coefficient is 3.0 S. The approximate molecular weight of the enzyme was determined by gel filtration (117)to be about 25,000. The amino acid composition of the enzyme was determined. Enzymic activity has been conveniently determined using L-glutaminyl-L-asparagine as the substrate by following the formation of ammonia. The pH optimum was 8.4 (Verona1 buffer) ; with L-glutamine as substrate, optimal activity was observed a t pH 9.4. The apparent K , value (with L-glutaminyl-L-asparagine) is 0.2 mM. The enzyme was found to be inactive toward D-glutamine, L-glutamic acid, glycyl-L-glutamine, L-asparagine, L-yglutamylhydrazide, glutathione, and y-L-glu113. M. Damodaran and P. Ananta-Narayanan. BJ 32, 1877 (1938). 114. M. Messer, C. M. Anderson, and L. Hubbard, Gut 5, 295 (1964). 115. H. G. Krainick, G. Mohn, and H. H. Fischer, Helv. Paediat. Acla 2, 124 (1959). 116. M. Messer and M. Ottesen, Compt. Rend. Truv. Lab. Carlsberg 35, 1 (1965). 117. P. Andrews, BJ 91, 222 (1964).

6.

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tamylglycine. L-Glutamine methyl ester is a substrate and is converted to pyrrolidone carboxylate methyl ester. The relative activity of the enzyme toward several substrates is given in Table 111. The enzyme also catalyzes the conversion of glutaminyl-tRNA to pyrrolidone carboxylyltRNA (76).

B. 7-GLUTAMYL TRANSPEPTIDASE AND 7- GLUTA MYL CYCLOTRAN SFERASE An enzyme possessing the properties of L-glutamine cyclotransferase has not as yet been found in animal tissue. However, the conversion of glutamine to pyrrolidone caboxylic acid by a two-step enzyme-catalyzed pathway has been observed (IS). Thus, glutamine may be converted to y-glutamyl glutamine by y-glutamyl transpeptidase: 2-~-Glutamine

y-glutarnyl

* 7-L-glutamyl-L-glutamine

transpeptidase

+ NHs

7-L-Glutamyl-L-glutamine is a substrate for 7-glutamyl cyclotransferase (see below), which converts it to pyrrolidone carboxylic acid and glutamine.

TABLE I11 SUBSTRATE SPECIFICITY OF GLUTAMINE CYCLOTRANSFERASE~ Substrate LGlutamine D-Glutamine L-Glutamic acid diamide LGlutaminyl-L-asparagine LGlu taminyl-cleucine L-Glutaminyl-Lasparaginyl-8-benzyl-lrcys teinyl-Lprolyl-bleucylgly cinamide L-Glutaminyl-lrasparaginyl-8-benzyl-bcys teinyl-b prolyl-N-tosyl-blysyl-glycinamide Glycyl-bglutamine L-Asparagine

Relative activity 0.02 0 0.7 1.0 1.0 1.4 1.5 0 0

The experiments were done in Conway dishes witahfraction 5 and 2.5 mM substrate in 0.1 M Verona1 buffer, pH 8.4. For other details, see Messer and Ottesen (116).

142

M. ORLOWSKI A N D A. MEISTER

VII. Enzymic Formation of Pyrrolidone Carboxylic Acid from 7-Glutamyl Amino Acids

A. Y-L-GLUTAMYL CYCLOTRANSFERASE (y-GLUTAMYL LACTAMASE) I n the course of studiea on the hydrolysis of glutathione by kidney extracts, \l'oodwarcl :ind Rcinliart 4 I rioted that 1)yrrolidone carboxylic acid was formed. Pyrrolidone carboxylic acid was the predominant product a t relatively alkaline yalucb:, of pH, while at pH values lower than 6.6, free glutamic acid was tlie major product. The formation of pyrrolidone carboxylic acid from glutathione in the presence of a more purified sheep kidney enzyme preparation waa later reported by Fodor et al. (22).The formation of pyrrolidone carboxylic acid in these systems may be ascribed to thc action of two enzymes, i.e., y-glutaniyl transpeptidase and 7-glutamyl cyclotransferaae. Hanes et al. (1181 were the first to demonstrate enzymic tranbfer of the y-glutamyl moiety of y-glutamyl peptides (or y-glutamyl amino acids ) to acceptor peptides and amino acids with the formation of a new 7-glutamyl linkage. Subsequent studies with purified preparations of y-glutamyl transpeptidase (119, 120) clearly demonstrated that this enzyme does not produce pyrrolidone carboxylic acid wlic~n inculxited with 7-glutarnyl amino acid3 or y-glutamyl peptides. The (libcowry by Connell and Hanes ( 5 1 of an enzyme that catalyzes the forinatioii of pyrrolidone carhoxylic acid from y-glutamylglycine, y-glutamylglutamate, and (more slowly) from y-glutamyl phenylalanine and glutathione thub seems to explain a number of earlier observations on the formation of pyrrolidone carboxylic acid in systems containing glutathione. Connell and Hancs (5) did not detect forination of free glutainic acid and revereibility of the reaction could iiot be demonstrated. 7-Glutamyl cyclotransferase activity wab later observed by Cliffe and Waley (1 1 ) in preparations of calf lens and rabbit liver. Extracts of these tissues catalyzed the cleavage of y-Lglutamyl-L-a-aminobutyrate and 7-L-glutamylglycine to pyrrolidone carboxylic acid and the corre+onding amino acids. The supernatant fraction of rat liver hornogenateb also catalyzed the conversion of ophthalmic acid (7-glutamyl-a-aminobutyrylglycine) to pyrrolidone carboxylic acid, but glutathione wab only slightly attacked. On the other hand, prcyarations from lens did not exhibit detectable action on the tripeptides. Studies 118. C. S. Hance. I;. J. R. Hid. :ind I;. A. Isherwood, BJ 51, 25 (1952) 119. F. J. H. H i d :ind P. H. Springell, BJ 56, 417 (1954). 120. M. Orlowski und A. Mcister. JBC' 240, 338 (1965).

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ENZYMOLOGY OF PYRROLIDONE CARBOXYLIC ACID

on the specificity of highly purified y-glutamyl cyclotransferase (IS) indicates that this enzyme differs considerably from that of 7-glutamyl transpeptidase (120).While y-glutamyl transpeptidase attacks virtually all y-glutamyl peptides and 7-glutamyl amino acids, the specificity of y-glutamyl cyclotransferase is much narrower. Thus, purified 7-glutamyl cyclotransferase does not attack glutamine, oxidized glutathione, or reduced glutathione. However, when both y-glutamyl transpeptidase and 7-glutamyl cyclotransferase are present, pyrrolidone carboxylic acid is formed from glutathione. These findings suggest that the actual substrate of y-glutamyl cyclotransferase is the corresponding y-glutamyl amino acid and that the formation of pyrrolidone carboxylic acid from glutathione proceeds in two steps as follows: 7-glutamyl

2-Glutathione

7-glutamyl glutathione

transpeptidase y-glutamyl

7-Glutamyl glutathione

+ cysteinylglycine

’ pyrrolidone carboxylic acid

cyclotransferase

+ glutathione

The formation of pyrrolidone carboxylic acid from glutamine (in the presence of both enzymes) probably takes place in an analogous fashion. In contrast to y-glutamyl transpeptidase, which is almost entirely bound to particulates, y-glutamyl cyclotransferase is present in the supernatant fractions of liver and other tissues. Studies on the distribution of the enzyme shows that it is present in most animal tissues but that there are some species differences (Table I V ) . Relatively high enzymic activity is found in kidney, liver, brain, and skin. In man, the highest activity is found in the brain. The unusually high activity in TABLE IV 7-GLUTAMYL CYCLOTRANSFERASE ACTIVITY OF SEVERAL ANIMALTISSUEP Tissue

Mouse

Guinea Pig

Rat

Skin Testes Kidney Brain Heart Liver Spleen Skeletal muscle Intestine Lung

185 141 83 79 63 52 57 23 14 14

426 60 133 66 30 101 86 29 64 54

157 39 286 12 11 77 17 5 89 17

Man 76 63 150 27 45 28 27 38

4 Mean values (units/mg protein) of several determinations (unpublished data obtained in the authors’ laboratory, 1969-1970).

144

M. ORLOWSKI A N D A. MEISTEK

skin is of interest and may be related to the observation that skin contains a rather high concentration of pyrrolidone carboxylic acid (121123) . Connell and Hanes ( 5 ) achieved a 20-fold purification of y-glutamyl cyclotransferase from pig liver by a procedure involving ammonium sulfate fractionation; the active protein precipitated in the fraction between 70 and 85% of ammonium sulfate saturation. A 14-fold purification of a similar enzyme from rat liver was reported by Kakimoto et al. (124).This enzyme was named y-glutamyl glutamine lactamase and was thought to be different from that isolated by Connell and Hanes ( 5 ) because i t was found to be inactive toward y-glutamylglycine and glutathione. However, since both the preparations obtained by Connell and Hanes (5)and Kakimoto e t al. (124)were of relatively low purity, it is difficult to be certain as to whether the reported differences in specificity reflect different enzymes; the presence of additional enzymes in these preparations which might catalyze interfering reactions might possibly account for the observed differences. The most highly purified preparations of y-glutamyl cyclotransferase that have thus far been obtained have been isolated from sheep and human brain (13).The method of isolation involves preparation of an acetone powder of brain followed by extraction and precipitation of inactive proteins at pH 4.2. Further purification is carried out by ammonium sulfate fractionation, heat denaturation of impurities, gel filtration on Sephadex G-75 columns, and chromatography on carboxymethyl cellulose and DEAE-cellulose columns. Several active protein fractions differing in isoelectric point have been found in the course of purification of sheep brain acetone powder extracts. The enzyme from human brain has been purified more than 2000-fold; the preparation obtained catalyzed the formation of about 250 pmoles of pyrrolidone carboxylic acid per milligram of protein per minute using the model substrate L-y-glutamyl-L-7-glutamyl-p-nitroanilide. At least three catalytically active proteins were obtained from human brain. During purification on Sephadex G-75 columns, the activity separated into two different distinct fractions. The lower molecular weight component comprised approximately 75% of the overall activity; after isolation it was shown by DEAEcellulose chromatography and by electrofocusing in a pH gradient (125, 126) to consist of two components differing somewhat in isoelectric 121. G. Pascher, Arch. Klin. Exptl. Dermatol. 203, 234 (1956). 122. G. Leonhardi, I. V. Glasenapp, and G. Bruhl, 2. Physiol. Chem. 292, 89 (1953). 123. K. Laden and R.Spitzer, J. SOC. Cosmetic Chemists 18, 351 (1967). 124. Y. Kakimoto, A. Kanazawa, and I. Sano, BBA 132, 472 (1967). 125. H. Svensson, Acta Chem. Scand. 15, 325 (1961) ; 16, 456 (1962). 126. H. Svensson, ABB 1, Suppl. 132 (1962).

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ENZYMOLOGY OF PYRROLIDONE CARBOXYLIC ACID

point (PI 4.06 and 4.25). The isolated enzyme was found to be stable a t 0" for a t least one month, and it was also stable in the lyophilieed state when stored a t -20". The pH optimum lies in the range 7.8-8.2 (borate buffer). The apparent K,,, value for L-y-glutamyl-L-y-glutamyl p-nitroanilide is 4 X lo-* M . Table V gives data on the specificity of the purified 7-glutamyl cyclotransferases obtained from human and sheep brain; the two enzymes exhibited similar specificity. The model substrate, L-7-glutamyl-L-7-glutamyl-p-nitroanilide was the most active substrate ; the corresponding L,D-, D,L-, and Dpisomers were inactive. The enzyme preparations also TABLE V SPECIFICITY OF

THE

7-GLUTAMYL CYCLOTRANSFERASES ISOLATED FROM HUMANAND SHEEPBRAIN" Formation of pyrrolidone carboxylate (pmole/mg/min)

Substrate

Human brain ~

y-~-Glutamyl-y-Lglutamyl-p-ni troanilide y-tGlutamyl-rcglu tamine 7-tGlutamy1-L-a-alanine y-rcGlutamylgly cine y-bGlutamy1-Lcu-aminobutyric acid 7-rcGlutamy1-S-met hyl-I,-cysteine y-rcGlu tamyl-Lvaline 7-rcGlutamyl-rcleucine y-rcGlutamy 1-rctyrosine 7-rcGlutamy1-Lp henylalanine 7-LGlutamylglycine ethyl ester 7-L-Glutamyl-8-aminoisobutyric acid Glutathione (reduced) Glutathione (oxidized) y-LGlutamyl-palanine LGlutamine 7-rcGlutamy1-p-nitroanilide y-~-Glutamyl-p-nitroanilide y-D-Glu tamyl-y-D-glutamyl-p-nitroanilide 7-D-Glutamyl-7-Lglu tamyl-p-ni troanilide y-bGlutamyl-y-D-glu tamyl-p-ni troanilide

150 (156)* 6 4 . 5 (68.8) 84.7 (77.6) 2 1 . 6 (22.5) 16.9 (18.5) 7 . 3 (15.3) 1.32 (9.7) 1 . 2 8 (7.95) 1.41 (13.5) 0.75 (8.95) 0 . 4 8 (6.75) 0.00 (0.31) 0 . 5 3 (18.5) 0 (7.1) 0 (0.79) 0 (2.96) 0 (59.2) 0 (0) 0 (0) 0 (0) 0 (4.85)

Sheep brain ~

~

100 (102)b 7 6 . 5 (81.3) 2 7 . 7 (38.8) 9 . 4 5 (15.9) 6 . 2 (9.45) 0 . 6 8 (7.15) 0 . 1 7 (7.15) 0 . 1 1 (4.10) 0.08 (3.10) 0 . 3 1 (5.95) 0 . 0 5 (0.11) 0 . 2 8 (17.8) 0 (10.5) 0 (0.71) 0 (2.01) 0 (22.6) 0 (0) 0 (0) 0 (0) 0 (1.97)

a The reaction mixtures contained substrate (4 pmoles; 2 rmoles for y-Lglutamyl-pnitroanilide), tris-HCl buffer (pH 8.0, 10 rmoles), and y-glutamyl cyclotransferase (0.1 unit; human brain enzyme specific activity 150 units/mg; sheep brain enzyme specific activity 100 units/mg) in a final volume of 0.25 ml. The mixtures were incubated a t 37" for 10-120 min, after which the formation of pyrrolidone carboxylate was determined as described in the text. * The values given in parentheses were obtained in experiments in which 1 unit of y-glutamyl transpeptidase was added to the reaction mixture [from Orlowski et al. (IS)].

146

M. ORLOWSKI AND A. MEISTER

exhibited activity toward L-y-glutamyl-L-glutamine, L-7-glutamyl-L-aalanine, L-y-glutamyl-L-a-aminobutyric acid, L-y-glutamylglycine, and L-7-glutamyl-S-methyl-L-cysteine. Although a number of other closely related compounds were not active or only slightly active, when 7-glutamyl transpeptidase was added, substantial formation of pyrrolidone carboxylic acid was observed with almost all of the y-glutamyl compounds. I n many cases activity was significantly increased ; notable are the values obtained with glutathione, L-y-glutamyl-p-nitroanilide, L-glutamine, L-y-glutamyl-L-phenylalanine, and L-y-glutamyl-L-valine. The enzyme did not act on L-a-y-glutamyl diglycine, L-a-glutamyl-Lalanine, L-y-glutamyl-P-alanine, L-y-glutamyl-P-aminoisobutyric acid. y-Glutamyl cyclotransferase activity is widely distributed and this observation suggests that the enzyme is of metabolic significance in many mammalian tissues. However, the function of this enzyme as well as that of a variety of y-glutamyl amino acids that have been found in nature remain to be elucidated. (See Section X.)

Vlll. Enzymic Formation of Derivatives of Pyrrolidone Carboxylic Acid

Studies on the transamination of glutamine with a-ketoacids (127132’) have shown that a-ketoglutaramic acid is formed as a product. a-Ketoglutaramic acid has been found to exist in solution in equilibrium with the cyclic ketolactam form: CONH,

At pH 7.5 less than 1% of the compound exists in the open-chain a-ketoacid form while a t pH 9 approximately 3% is in a form that reacts as a typical a-keto acid. Glutamine transaminase also catalyzes the transamination of glutamic acid y-A‘-methylamide ; the expected transamination product, a-keto-N-methylglutaramic acid has not yet been isolated from a transamination reaction mixture. However, this compound was 127. A . Meister and S. V . Tim, JBC 187, 173 (1950). 128. A. Meister, JBC 197, 309 (1952). 129. A . Meister, JBC 200, 571 (1953). 130. T. T. Otani and A. Meister, JBC 224, 137 (1957). 131. A . Meister, “The Enzymes,” Vol. 6, p. 193, 1962. 132. A. Meister, Science 120, 43 (1954).

6.

147

ENZYMOLOGY OF PYRROLIDONE CARBOXYLIC ACID

prepared by enzymic oxidation of glutamic acid y-N-methylamide; the product isolated exhibited the properties of 5-hydroxy-N-methylpyrrolidone carboxylic acid; it was not possible to obtain the a-keto analog of glutamic acid-y-N-methylamide in a form possessing a reactive keto group (129).The observation that glutamic acid-7-N-methylamide is a substrate for glutamine transaminase (127) and that methylamine is apparently formed (in the presence of both glutamine transaminase and a-ketoglutaramic acid amidase) suggests that the open-chain form (i.e., a-keto-N-methylglutaramic acid) is demethylamidated rapidly under these conditions, probably more rapidly than its rate of cyclization to 5-hydroxy-N-methylpyrrolidone carboxylic acid. The formation of a-keto-y-methylglutaramic acid by an analogous transmination reaction with y-methylglutamine has also been demonstrated. The a-ketoglutaramic acid amidase present in liver (and other tissues), which catalyzes the deamidation of the open-chain form of a-ketoglutaramic acid, and apparently a-keto-A'-methylglutaramic acid, has no activity toward aketo-y-methylglutaramic acid. Hersh et al. (133) have studied an enzymic reaction between methylamine and a-ketoglutarate which is catalyzed by cellfree extracts of a Pseudomonas; the product of this reaction was shown to be 5-hydroxyN-methylpyrrolidone carboxylic acid. Although the mechanism of this reaction is not as yet understood, it was suggested that the synthesis of 5-hydroxy-N-methylpyrrolidonecarboxylic acid involves two steps. The first of these is presumably enzymic and leads to the formation of the y-N-methylamide of a-ketoglutaric acid: (Y-

In the second postulated step, the product spontaneously cyclizes to yield 5-hydroxy-N-methylpyrrolidone carboxylic acid.

IX. Pyrrolidone Carboxylyl Peptidase

Doolittle and Armentrout (46) recently described an enzyme that selectively cleaves pyrrolidone carboxylyl residues from polypeptide chains. The enzyme was obtained from a strain of Pseudomonas fluores133. L. B. Hersh, L. Tsai, and E.

R. Stadtman, JBC 224, 4677

(1969).

148

M. ORLOWSKI AND A. MEISTER

cens which was isolated from soil. The organism is capable of growing on free pyrrolidone carboxylic acid as the sole source of carbon and nitrogen [see also Maruyama and Nomura ( 1 3 4 ) l . Although the physiological function of this enzyme is not clear, it is clearly of value in the characterization of many proteins and peptides which possess N-terminal pyrrolidone carboxylyl residues. This peptidase has also been found in Bacillus subtilis and in other bacteria ( 1 3 5 ) , and a similar activity (cleavage of the amide linkage of pyrrolidone carboxylic acid amide) has been found in Vibrio dunbar preparations (136). Pyrrolidone carboxylyl peptidase activity has been followed using pyrrolidone carboxylyl L-alanine (or other pyrrolidone carboxylyl-amino acids) as substrate by following the amount of amino acid released as determined by the ninhydrin method (13'7).I n another assay procedure the release of p-naphthylamine from L-pyrrolidone carboxylic-pnaphthylamide (135) is determined by the diazotization procedure of Bratton and Marshall (138) as modified by Goldberg and Rutenberg (139). Armentrout and Doolittle (140) purified the enzyme from a sonicate of Pseudomonas fluorescens by a procedure involving high speed centrifugation of the sonicate, precipitation of impurities by protamine sulfate, ammonium sulfate fractionation, Sephadex G-200 gel filtration, chromatography on DEAE-Sephadex, and preparative gel electrophoresis. A 200-fold increase in specific activity was obtained. These authors stabilized the enzyme during purification by addition of a relatively high concentration (0.1 M ) of 2-pyrrolidone. The enzyme was completely inhibited by brief exposure to sulfhydryl reagents, e.g., iodoacetamide and p-mercuriphenylsulfonate. The most purified preparation was reported to exhibit two components on gel electrophoresis. Studies on the specificity of the enzyme (141) showed that pyrrolidone carboxylyl-L-alanine (Pyr-Ala) was the best substrate. The influence of the penultimate amino acid on the rate of hydrolysis of Lpyrrolidone carboxylyl-L-amino acid dipeptides is considerable. The order of the relative hydrolysis rates varied in the sequence: Pyr-Ala, Pyr-Ile, Pyr-Val, Pyr-Leu, Pyr-Phe, Pyr-Tyr. L-Pyrrolidone carboxylyl 134. Y. Maruyama and M. Nomura, J. Biochem. (Tok y o) 43, 327 (1956). 135. A. Seewczuk and M. Mulceyk, European J. Biochem. 8, 63 (1969). 136. U. Senji, 0. Hiwatashi, and 0. Kimio, Tohoku J. ExpZ. M e d . 42, 46 (1942). Chem. Abstr. 42, 5067 (1948). 137. C. H. W. Him, S. Moore, and W. a.Stein, JBC 219, 623 (1956). 138. A. C. Bratton and E. K. Marshall, Jr., JBC 128, 537 (1939). 139. J. A. Goldbarg and A. M. Rutenburg, Cancer 11, 283 (1958). 140. R. W. Armentrout and R. F. Doolittle, ABB 132, 80 (1969). 141. J. A. Uliana and R. F. Doolittle, P B B 131, 561 (1969).

6. ENZYMOLOGY

OF PYRROLIDONE CARBOXYLIC ACID

149

L-proline is not detectably hydrolyzed. The enzyme exhibits considerable optical specificity ; thus, D-pyrrolidone carboxylyl L-alanine is not hydrolyzed while L-pyrrolidone carboxylyl D-alanine is hydrolyzed about 30 times more slowly than the corresponding L-L isomer. Pyrrolidone carboxylyl peptidase from B. sirbtilis (135)exhibits properties similar to the enzyme obtained from Pseudomonas. It is of interest that the enzyme is inactivated by shaking with air a t room temperature for 5 min and that reactivation is readily achieved by brief incubation with 2-mercaptoethanol and EDTA. Studies on pyrrolidone carboxylyl peptidase have shown that the enzyme can catalyze the removal of the terminal pyrrolidone carboxylyl residue from fibrinopeptides, fibrinogen (135,l4O), and human seromucoid (136). A recent report describes the presence of pyrrolidone carboxylyl peptidase in rat liver (46).A partial purification of the enzyme has been obtained by ammonium sulfate fractionation and gel filtration on Sephadex G-200. The rat liver eiizyme is similar to that obtained from Pseudomonas; thus, it is protected against inactivation by 2-pyrrolidone and exhibits similar substrate specificity.

X. Pyrrolidone Carboxylate Metabolism

There is relatively little definitive information in the literature about the metabolism of pyrrolidone carboxylic acid; although several reports indicate that pyrrolidone carboxylic acid can be utilized by mammals (104,142-1467, and certain microorganism (45,134, 147-1521, information relating to the enzyme-catalyzed utilization of pyrrolidone carboxylic 142. R . Muraclii, Acta Schol. Med. Univ. Kioto 7 , 445 (1925). 143. R. M . Rctlikr and H. Strcnback, JBC 58, 105 (1923). 144. S. Prdrrscn and H. B. L(.wis. JBC 154, 705 (1944). 145. L. D. Greenberg and C. L. A . Schmidt, Univ. Calif. Berkeley Publ. Physiol. 8, 129 (1936) ; Chem. Abstr. 30, 8388 (1936). 146. J. S. Butts. H. Hlunden, and M. S. Diinn. JBC 119, 247 (1937). 147. P. Sirrionnrt and K. Y. Chow. Antonze vrcu Leeuwenhoek 1. Microbiol. Serol. 19, 34 (1953) ; Chem. Abstr. 47, 8173 (1953). 148. M. Forbes and M. G. Sevag, ABB 31, 406 (1961). 149. T. Toss : i n t i I. Chibuta, J. Btrcteriol. 89, 919 (1965). 150. Y. Ihwai, Y. Kawai, and T. Urrnurn, Agr. Biol. Chem. ( T o k y o ) 29, 395 (1'365). 151. Y. Kanai and T. Yrmura, Agr. Uiol. ('hem. ( T o k y o ) 30, 438 (1966). 152. Y. Kawai, K. Aida, and T. Ucinura, Agr. Biol. Chem. ( T o k y o ) 33, 212 (1'369).

150

M . ORLOWSKI AND A. MEISTER

acid has been slow in accumulating. Recently, Ramakrishna et al. (153) reported that when l4C-~-pyrro1idonecarboxylic acid was injected intraperitoneally into rats more than half of the radioactivity appeared in the expired carbon dioxide within 30 min; these workers also found that liver and kidney slices converted 14C-pyrrolidone carboxylic acid to 14C0, and to 14C-glutamate.I n independent studies in the author's laboratory it was found that slices of rat kidney, liver, heart, muscle, skeletal muscle, and brain oxidized "C-pyrrolidone rarboxylate to "Co, ; analysis of the intracellular amino acid pool in the experiments with kidney slices revealed that 70% of the initial radioactivity was present in glutamate and glutamine (154). Rush and Starr (155) observed in the course of studies on the incorporation of pyrrolidone carboxylic acid into tRNA that a soluble rat liver preparation catalyzed the conversion of 14C-pyrrolidone carboxylate to "C-glutamate and "C-glutamine in the presence of ATP. Recent studies conducted by the authors have shown that when rat kidney extracts were incubated with "C-L-pyrrolidone carboxylate in the presence of ATP, magnesium ions, and an ATP-generating system, more than 80% of the radioactivity was converted to 14C-glutamine; the remainder was present as '*C-glutamate. When such incubations were carried out in the presence of L-methionine sulfoximirie [an irreversible inhibitor of glutamine synthetase (156)1, all of the radio-

FIG. 1. The y-glutamyl cycle. I, y-Glutamylcysteine synthetase ; 11, glutathione synthetnse ; 111, y-glutamyl transpeptidase; IV, y-glutnrnyl cyc1otr:tnsferase; V. peptidase. PCA = pyrrolidone carboxylic acid ; AA = amino acid. 153. M. Ramakrishna, P. R. Krishnaswamy, and Rajagopal D. Rao, BJ 118, 895 (1970). 154. P. Van Der Werf. M . Orlowski. and A . Meister. unpublished data (1970). 155. E. .4. Rush and J. I,. Stnrr, BBA 199, 41 (1970). 156. R. .\. Ronzio :ind A . Mrialrr. Ploc. S o t l . Acntl. Sci. I ' . S. 59, 164 (1968).

6.

151

ENZYMOLOGY OF PYRROLIDONECARBOXYLIC ACID

activity was found in glutamate (154). It thus appears that rat kidney (and probably other animal tissues) contains an enzyme that converts pyrrolidone carboxylic acid to glutamate in a reaction that requires ATP and magnesium ions. Evidence has been presented that rat kidney contains enzymes that catalyze the utilization and synthesis of glutathione, i.e., y-glutamyltranspeptidasc, y-glutamyl-cyclotransferase, y-glutamylcysteine synthetase, and glutathione synthetasc. The reactions catalyzed by these and two other enzymes, which involvc the uptake and release of amino acids from y-glutamyl linkage and the formation and utilization of pyrrolidone carboxylic acid, constitute a cyclical process (Fig. 1) which has been termed the “7-glutamyl cycle” (157).It has been suggested that the y-glutamyl cycle may function in the transport of amino acids.

157. M. Orlowski and A . Meister, Proc. iVatl. Acnd. Scz.

U. S . 67,

1248 (1970).

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Stabhv lococcal Nuclease X-Ray Structure F. ALBERT COTTON

EDWARD E. HAZEN, JR.

I . Introduction . . . . . . A. Why Staphylococcal Nuclease?

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. B. Introduction to thc Crystnllographic Stiidics . 11. The Conformation of the Peptidc Chain . . . .

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111. The Binding of Thymidinc-3’,5’-Diphosphate and Calcium Ion . . . . . . . . . . . IV. Some Correlations with Stirdics in Solution . . . . V. Some Tentative Comments on Mechanism and Plnns for Future Studies . . . . . . . . . . A . A Warning . . . . . . . . . . B. Mechanism . . . . . . . . . . C. Proposed Future Studies . . . . . . .

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153 153 156 159 163 172 174 174 174 175

I. Introduction

A. WHYSTAPHYLOCOCCAL NUCLEASE? I n the second edition of this work t h e subject of the extracellular nuclease of Staphylococcus a w e u s , ribonucleate (deoxyribonucleate) 3’nucleotidohydrolase, EC 3.1.4.7,was quite adequately covered in some six-hundred words ( 1 ) . I n this edition there are two chapters jointly containing more than a n order of magnitude more words about this forinerly obscure, nonspecific riuclease with its high pH optimum (pH 9-10) and 1. M. Laskowski, Sr.. “The Enzymes,” 2nd ed., Vol. 5, p. 142. 1963 153

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F. A. COTTON A N D E. E. H A Z E N , J R .

requirement for relatively high (0.01 41) calcium ion concciitration for full activity ( 2 ) . Earlier work on this cnzymc has been well rcvicwetl (1, 3-5). I n recent ycars work on the cnzymc in solution has been reported from only four 1al)oratorics. Von Hippel and Fclscnfelrl (6)and Wingert and voii Hippel (7) have examined some parameters influencing the mode and rate of hydrolysis of deosyribonucleic acid (IINA) by the enzyme. 1,askows;ki :ind his collaborators havc rontinuetl thcir intcrcst in the nuclease as a tool in nuclcic acid rcsearch with further studies of its purification, specificity, and mode of action (8-12). ,Jardetzky and hie colleagues are in the process of publishing their cxtcnsive nuclear magnetic resonance studies of native as well as specifically dcuteratcd derivatives of the enzymc (13-18). The most intcnsivc and comprehensive studies of the nuclease arc being executed in C. B. Anfinsen’s laboratory, with particular recent emphasis on factors influencing chain folding. Tliese may be considered as a natural extension of earlier work on conformtitional aspects of bovine pancreatic ribonucleasc (19, 2 0 ) . In the present case, the system is free of complications arising from the prcscncc of disulfide cross-links for, in its single chain of 149 amino acid residues, the nuclease has no covalent intrachain cross-linking nor, for that matter, any sulfhydryl groups (21-23). A description of iiivestigations on the 2. P. Cuatrctcasns. S. Fuchs, and C. B. Anfinsen, JBC 242, 1541 (1967). 3. M. Laskowski. Sr.. Atlvtrn. Enzymol. 29, 166 (1967). 4. M. Privat dc Garilhr, “Enzymes in Nucleic Acid Research.” p. 170. H(miann. Paris, 1967. 5. P. Cuatrecasas, H. Taniuchi, and C. B. Anfinsen. Brookhaven Symp. B i ~ l 21, . 172 (1968). 6. P. H. von Hipprl and G . Fclsenfcld. Biocheniis/ry 3, 27 (1964). 7. I,. MTingrrt :ind P. H. von Hipprl. U B A 157, 114 (1968). 8. E. Sulkowski and M. Laskowski, Sr., JBC 241, 4386 (1966). 9. E. Sulkowski and M. 1,askowski. Sr.. JBC 243, 651 (1968). 10. E. Sulkowski and M. Laskowski, Sr., JBC 243, 4917 (1968). 11. E. Sulkowski and M. Idaskowski, Sr., JBC 244, 3818 (1969). 12. A. J. Mikulski, E. Sulkowski, L. Stasiuk, and M. Laskowski, Sr., JBC 244, 6559 (1969). 13. D. H. Meadows, J. L. Markley, J. S. Cohen, and 0. Jardetzky, l’roc. Nnll. Acad. Sci. U . S. 58, 1307 (1967). 14. J. L. Markley, I. Putter, and 0. Jmdetzky, Z. Anal. Chem. 243, 367 (1968). 15. J. L. Markley, I . Putter, and 0. Jardetzky. Scieme 161, 1249 (1968). 16. I. Putter, A . Barreto, J. L. Markley, and 0. Jardetzky, Proc. N a t l . Acnd. Sci. U.S. 64, 1396 (1969). 17. I. Putter, J. L. Markley, and 0. Jardetzky. Proc. N n l l . Acad. Sci. U. S. 65, 395 (1970). 18. J. I,. Markley, M. W. Williams. and 0. Jardetzky, Proc. Natl. Actcd. Sci. C.. S. 65, 645 (1970). 19. C. B. Anfinsen. Harvey Lectures 61, 95 (1967). 20. C. 13. Anfinsen, Develop. Biol. S u p p l . 2, 1 (1968).

7. STAPHYLOCOCCAL

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structure, function, and synthesis of staphylococcal nuclease is given in earlier reviem ( 5 , 241 and i i i Chapter 8 by Anfinmi et al. in this volume ( 2 5 ). One observation liab 1)ecn particularly critical in the X-ray crystallographic investigation of the enzyme. It was found that thymidine-3’,5’-diphospliate ( pdTp) i n the presence of calcium ions is a potent in1iil)itor of the nucleasc ( 2 6 ) .The obligate ternary complex (27) of nuclea~e-ldTI)-Ca’+ha. hoiiie Iwopcrtic. nhicli differ markedly from those of the uninhibited nucleabc ; the differenceb are probahly only in part the result of ~ n i a l conformational l change* (5.18,30,3 1 ) .T h e inhibited nuclease is more rehi&iiit to urea ( 5 ) and heat ( 5 ,9, 3 2 ) denaturation, has a distinctly altered pattern of reactivity of its tyrosine residues toward chemical inodification (32,3 3 ) , has ahout 34 fewer easily exchangeable protonb (34, 3 5 ) , and is inore rebistant t o liydrolybis by the proteolytic enzyme, trypsin, which now cleaves it specifically in only two places (between residues 5 and 6 and I)etneen 48 and 4 9 ) . The combination of 6 4 8 and 49-149 fragiiients has bigiiificant (about 10% 1 enzymic activity (36-38). X-ray crybtallographic study was undertaken to supply a structural 21. H. Taniuclii. C. U. Anfinsen. and il. Sod.i:i, JBP 242, 4752 (1967). 22. C . I.. Ciisuniano. H. T:iniuclii, :ind C. B. ilnfinsen, JBC 243, 4769 (1968). 23. H. Tnniuchi. C. L. Cusumano. C. b. Anfinsen, and J. L. Cone, JBC 243, 4775 (1968). 24. C. B. Anfinsen, Pure Appl. Chem. 17, 461 (1968). 25. C. B. Anfinsen. P. Cuatreciisns. and H. Taniuchi, “The Enzyrnrs.” 3rd ed., Vol. IV, p. 177, 1971. 26. P. Cuntrec:isas, S. Fuchs. and C. B. Anfinsen, JBC 242, 1541 (1967). 27. Although gel filtration ( 2 s ) and sl~t~ctroscol)ic (29) ni(~asurmientsof riuclcotidc binding indirate nn nbsolut,e rcclriirement for cnlciurii ion and virc vc’rsa. the N M R studies (13-15, 1 s ) cuggcst thnt both C‘a” and nuchotides c:in indrlwndently bind to the nucleasr but that further rh;ingrs takr pl:icr in the NMR spcctruni when all three (Ca”, pdTp. and nuclease) are present,. As Markley et al. pointed out (15) this aliliarent discrrpancy is probably only a mattcr of rnethodological sensitivity and tightness of binding. 28. P. Cuntrcrasas. S. Fuchs, and C. 13. Anfinscn. JBC 242, 3063 (1967). 29. P. Cuatrecasas. S. Furhs, and C. B. rlnfinsrn, JBP 242, 4759 (1967). 30. P. Cuatrecasas, H. Edelhoch, and C. B. ilnfinscn, Proc. Nntl. Acad. Sci. 7,’. S . 58, 2013 (1967). 31. A. Arnone. C. J. Birr, F. A . Cotton, E. E. Hazen, Jr., 11. C . Richardson, and J. S. Richardson, Proc. N n t l . Acnd. Scz. G. S. 64, 420 (1969). 32. 1’. Cuatrecasas, S. Fuclis, :ind C. 13. Anfinsen. HUA 159, 417 (1968). 33. 1’. Cuatrrcasas, S. F L I C ~and S . C. B. Anfinsen. JBC 243, 4787 (1968). 34. A . N. Schechter, L. Moravek, and C. €3. Anfinsen. Proc. N n f l . Acrrd. Sci. U . S . 61, 1478 (1968). 35. A . N. Schcchter, L. Moravek. and C. B. Anfinsen, JBC 244, 4981 (1969). 36. H. Taniuchi, C. B. Anfinsen. and A. Sodja, Proc. Nutl. Acrtd. Sci. U . S. 58, 1235 (1967). 37. H. Taniuchi and C. B. Anfinsen, JBC 243, 4778 (1968). 38. H . Taniuchi, L. Moravek. and C. B. Anfinsen, JBC 244, 4600 (1969).

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basis for understanding these and other properties of the nuclease as well as to help elucidate the mechanism of its action.

B. INTRODUCTION TO THE CRYSTALLOGRAPHIC STUDIES We began our investigation (39) of the crystal structure of the staphylococcal nuclease with attempts to crystallize the enzyme in the spring of 1963. At approximately the same time that Sulkowski and Laskowski crystallized the nuclease from ammonium sulfate (8J, we obtained suitable crystals for X-ray diffraction purposes (40) and began active crystallographic work about the beginning of 1966. All our work has been done on the nuclease from the Foggi strain of Staphylococcus aureus ( 2 2 ) . All crystals have been grown from low ionic strength buffers ( p N 0.01) with 2-methyl-2,4-pentanediol as the precipitant ( 4 1 ) . The detailed conditions for the growing of crystals, the preparation of heavy-atom derivatives, the collection of data, and the production of electron density maps are given elsewhere (31, 40, 4 2 ) . Anomalous dispersion measurements were made on all heavy-atom derivatives ; this allows an electron density map to be calculated with the data obtained from a single heavy-atom derivative ( 4 3 ) . We have studied crystals of both the uninhibited nuclease and the nuclease-pdTp-Ca’+ complex and treated them as independent structures. The crystals of uninhibited nuclease, hereafter referred to as Type I, have unit cell parameters a = b = 47.75 A, c = 63.5 A ; the ternary complex (inhibited nuclease) crystals, hereafter called Type I I , have unit cell parameters a = b = 48.3 A, c = 63.3 A ; the space group of both is P4, (31). We shall show views of the structure of the nuclease a t three ‘‘levels of resolution”; these structures were determined as follows : (1) At 6 A resolution, Type I crystals phased from derivatives prepared by soaking in PtCli-, p-chloromercuribenzenesulfonic acid (PCMBS) and p-acetoxymercurianiline (PAMA) and using all the data within a 6 A sphere in reciprocal space. 39. The others n.ho have been associated with this project, in order of appenrance, nre D. C. Richardson, J. S. Richardson, A. Amone, C. J. Bier, V. W. Day, A . Yonath, and S. Lnrsen. 40. F. A. Cotton, E . E. Hasen, Jr., and D. C. Richardson, JBC 241, 4389 (1966). 41. M. V. King. BBA 79, 388 (1964). 42. A. Arnonc, C. J. Bier, F. A. Cotton, E. E. Hazen, Jr., D. C. Richardson, J. S. Richardson, V. W. Day, and A. Yonath, JBC (1971) (in preparation). Some additional details will be found in the P1i.D. thesis of A. Amone, Massachusetts Institute of Technology, 1970. 43. A. C. T. North, Acta Cryst. 18, 212 (1965).

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STAPHYLOCOCCAL NUCLEASE X-RAY STRUCTURE

157

(2) At 4 A resolution, again using all the data. (a) Type I crystals phased with the same heavy-atom derivatives. (b) Type I1 crystals phased with the single substitution of an iodine atom for a methyl group. (3) At high resolution, Type I1 crystals phased with iodine for methyl and barium for calcium ion substitutions using all the data t o 4 A but only that 35% of the reflections from 4 to 2 A with the highest intensities. I n the Type I1 crystal the iodine atom was introduced by substituting 5-iododeoxyuridine-3,5’-diphosphate for pdTp when growing the crystals. For the other derivative the bound calcium ion was replaced by a barium ion by soaking, since it proved difficult to grow large well-formed bariumcontaining crystals de novo. Our results indicate that the iodine atom replaces the methyl group, and the Ba’+ the Cazf with full occupancy and high isomorphicity. There is an indication that the Ba‘+ position is somewhat displaced from that of the Ca‘+ (4.2). Although the determination of protein structures by X-ray crystallography has had immense impact on the field and although these crystal structures appear to be excellent approximations of the structures in solutions ( 4 4 ) ,the unsophisticated use of this information is not without its dangers. If it has not already happened, it is almost inevitable that a protein-crystallographic group will someday make a gross error-minor errors or uncertainties are commonplace-in the interpretation of their electron density map. Indeed, our private attempts to fit our 4-A resolution maps located the path of much of the peptide chain correctly but were completely out of register with the sequence. I n the determination of the structure of small molecules by X-ray crystallography, the resolution is sufficient to reveal individual atoms, thus allowing, for one thing, the application of prior knowledge of reasonable values for bond lengths and angles. I n addition it is possible, with the aid of the computer, to apply reasonably rigorous tests to the correctness of the structure. Although the ability to test the fitting and correctness of a protein electron density map is growing (45-48), we believe these maps, not because we can be sure they are correct but because they satisfy our notions of what a protein should look like, were obtained with valid 44. J. A . Rupley. in “Structure and Stability of Biological Macromolerules” (S.N . Timashcff and G. D. Fasman, eds.). p. 291. Marccl Dckker, New York, 1969. 45. F. M. Richards, J M B 37, 225 (1968). 46. R. Diamond, Actn Cryst. 21, 253 (1966). 47. R . Diamond, Acta Cryst. A25, 5189 (1968) (Abstr. 81h Intern. Congr. I n t s i n . Union Crystallog.). 48. M. Levitt and S. Lifson, J M B 46, 269 (1969).

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techniques by competent people and are, after all, the only electron density maps we have of proteins. The recent dispute over the interpretation of the structure of DNA (49-53) is not without relevance to protein structures, even though the isomorphous replacement method allows the independent determination of magnitudes and phases. Indeed, Donohue's comments on resolution, disorder, and the R value would appear most apt (49). To come to our point, it may be very worthwhile for students to build Kendrew models of enzymes from lists of coordinates, but for the investigator whose work depends critically on the knowledge of an enzyme structure, complete reliance on such a model is risky; the electron density map itself should also be consulted where crucial questions of

FIQ.la. 49. J. Donohue, Science 165, 1091 (1969). 50. M. H. F. Wilkins, D. A. Marvin, and L. D . Hamilton. Science 167, 1693 (1970) 51. F. H. C. Crick, Science 167, 1694 (1970). 52. S. Amott, Science 167, 1694 (1970). 53. J. Donohue, Science 167, 1700 (1970).

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STAPHYLOCOCCAL NUCLEASE X-RAY STRUCTURE

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atomic configurations arise. In the first place, a set of coordinates is a secondary source of data involving someone else’s partially subjective interpretation of the primary data; in the second place, there is considerably more information in the electron density map than t.here is in a model. The lack of coordinates in this chapter should, however, not be taken as implying that we disapprove of their publication but only that we have not yet measured a set with sufficient precision to justify inclusion here.

II. The Conformation of the Peptide Chain

Figure l a is the front view of a Styrofoam model of the nuclease made from the 6-A electron density map of Type I crystals. As we speak, subsequently, of the front, back, top, bottom, etc., of the molecule, we are

Fic:. 1 . (a) Front virw of the 6-A Styrofoam model of the uninhibited nuclease (Typca I crystals); (b) view of thr samr Styrofoam model from the left side.

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P. A. COTTON A N D E. E. HAZEN, J R .

FIG.2a.

viewing it as it is shown here with right and left meaning the viewer’s right and left. Figure l b is the same model viewed from the left-hand side. Figures 2a and 2b show the front and right views, respectively, of the conformation of the peptide backbone of the nuclease, based on the high resolution structure of the Type I1 crystals. The latter photographs, which are black and white reproductions of the original color slides, are taken with UV illumination of fluorescent dye-filled, transparent tubing tied to each a-carbon of a Kendrew model of the nuclease (54).Figure 3 is a drawing of the peptide chain showing the approximate positions of the a-carbon atoms. Figure 4 is the amino acid sequence of the Foggi strain of Staphylococcus aureus ( 2 2 ) . As one views the nuclease from the front, the N-terminal residue is a t the upper left rear. From this point the chain goes diagonally down 54. These photographs were made by C. B. Anfinsen, D. C. Richardson, and J. S. Richardson. This technique is particularly effective when the flow of dye through the tubing is vicwed as a function of time in a motion picture.

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FIG.2. (a) Front view of the conformation of the peptide chain of the inhibited nudeme at high resolution (Type I1 crystals). ( b ) Right side view of the chain conformation a t high resolution. The curlicue at the upper left center represents the inhibitor pdTp.

across the back of the molecule until at residue 12 there begins a nearly vertical, inwardly curving section of antiparallel /?-pleated sheet made up of three strands which run approximately front to back. This sheet, consisting of some 24 residues, forms the upper right flank of the nuclease molecule. Between residues 37 and 44 the chain loops through the center of the molecule and then between residues 44 and 53 forms the loop which can be seen extending out into solution in the side view of the chain. Residues 54 through 67 form some three and one-half turns of helix which runs from front to back. From the end of the helix the chain rises up the back of the molecule, comes to the front a t the top right, at T y r 85 turns again toward the back and then drops down the back to the start of another helical section (residues 99-107) of two and one-half turns that comes toward the front bottom of the molecule, slanting down a t an angle of about 20" from the horizontal. From

162

FIG.3. Drawing of

F. A. COTTON AND E. E. HAZEN, JR.

:I

front view of the chain conformation showing the locations

of the a-carbon atoms along the chain. The positions of the 3’- and 5’-phosphatr groups of pdTp are indicated by X and the Caz+by 0 .

10

20

FIG.4. The amino acid sequence of the staphylococcal nucleuse, Foggi strain.

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this point, the chain rises directly up forming the left front of the molecule, loops toward the rear with Pro 117 a t the peak of the loop, and drops down to the start of another helical section a t residue 122. At a n angle of 40” with the horizontal, this helical section runs down and forward to residue 134 near the bottom left front of the molecule. The chain ends in a complex loop near the bottom left of the molecule. Despite the fact that the phases for the high resolution map were obtained from only two heavy-atom derivatives ( 5 5 ) ,the electron density map was quite interpretable with the aid of the sequence along most of the peptide chain. This is illustrated in Fig. 5a which shows the high resolution map of the bottom right helix, one of the best resolved sections. Figure 5b shows the same section phased only with the high resolution iodine data; Fig. 5c, the Type I1 crystals phased with iodine and barium at 4 A ; and Fig. 5d, the Type I crystals phased with the three heavy-atom derivatives a t 6 A resolution. The five N-terminal residues and the six or seven C-terminal residues cannot be seen in the high resolution electron density map, and the loop referred to above, formed by residues 44 to 53, appears a t only one-third to one-half the amplitude of the well-resolved parts of the map. The lack of clarity in theae three regions might possibly result from poor phasing or some other crystallographic factor, but we consider it more likely that these predominantly hydrophilic sections of the peptide project in a disordered way into the solvent. In this connection, it is interesting that in the presence of Ca’+ and pdTp trypsin cleaves inhibited nuclease a t only two points: between residues 5 and 6 and between residues 48 and 49 (36-38) which are a t the very extremity of the loop. It also seems relevant that ribonuclease S also shows lack of clarity a t the ends of the peptide chains and in the region of a relatively exposed loop (66)* 111. The Binding of Thymidine-3’,5’-Diphosphate and Calcium Ion

In the front view of the Styrofoam model (Fig. l a ) a distinct pocket is seen in the upper left quadrant of the model. Figure6a shows a view from directly above this pocket in the 4-A electron density map of the 55. I n addition, not only are iodine and barium relatively light “heavy” atoms. but they are replacing a carbon atom (the 5-methyl of thymine) and a calcium ion. respectively, giving net changes of 47 and 36 electrons, respectively. 56. H. W. Wyckoff. D. Tsernoglou, A. W. Hanson, J. R. Knox, B. Lee, and F. M. Richards, JBC 245, 305 (1970).

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F. A. COTTON AND E. E. HAZEN, JR.

(b)

FIG.5a and b. Type I (uninhibited) crystals. I n a rather marked contrast, Fig. 6b shows the same view of the 4-A electron density map of the Type I1 (inhibited) crystals. A mass of electron density now appears in the pocket, and there also appears to be some added density a t the left side of the perimeter of the pocket itself. It should perhaps be reemphasized here that these structures of the Type I and Type I1 crystals were determined completely independently using one set of heavy atoms and unsubstituted data for the uninhibited nuclease and another entirely different combination for the nuclease-inhibitor complex. However, the fact that heavy-atom positions appear very distinctly when intensities from Type I crystals are combined with Type I1 phases (and vice versa) indicates that the nuclease undergoes no gross conformational changes with pdTp and Caz+binding (31) ; thus, the kind of comparison presented

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FIQ.5. (a) The helical section a t the lower right of the nuclcase as seen from dircctly above. This section is from the high resolution map phased with both Ba" and I in the Type I1 crystal. The heaviest lines are meant to indicate the groups closest to the viewer. (b) The samc helical section a t high resolution but phased only with I in Type I1 crystals. (c) The same helical section a t 4 A resolution phased with both Ba2+and I in Type I1 crystals. (d) The same helical section a t 6 A resolution in Type I crystals phascd with three heavy-atom derivatives.

in Fig. 6 is justified. From its dimensions and amplitude distribution, the electron density seen in the pocket in Fig. 6b may be attributed to the pdTp molecule with the pyrimidine ring seen approximately edge-on in the upper part and the two phosphate groups nearly superposed in the lower part. This interpretation is strikingly and unequivocally confirmed by

166

F. A. COTTON AND E. E. HAZEN, JR.

FIG.6a.

the electron density contours shown in Fig. 7a ( 5 7 ) . These are in planes which are vertical and are viewed from the right side (directions relative to the canonical orientation of the molecule as defined earlier). According to the interpretation suggested for Fig. 6b, we should now be viewing the pdTp molecule approximately perpendicular to its mean plane. Figure 7b shows a space-filling model of pdTp, to the same scale as the electron density sections of Fig. 7a. It is clear that the pyrimidine ring of the pdTp lies within the pocket formed by the enzyme with the phosphate groups exposed to solution at the enzyme surface, the 3‘-phosphate toward the top, the 5’-phosphate toward the bottom, and very close to dead front-center of the model in Fig. l a . It does not appear to be possible to fit the inhibitor to the electron density maps with any significantly different orientation or configuration of the pdTp. The high resolution map has revealed the qualitative character of the binding of the pdTp and Ca2+,but the accurate fitting of map to model has, unfortunately, not proceeded quite far enough a t this writing to permit a quantitative discussion. The calcium ion is almost directly below the $-phosphate and is complexed by the carboxylate side chains of 57. The map in this figure was calculated with phases from both the Ba” and I substitutions. We show a very similar map in Fig. 6 of Arnone et al. (SO),but that one-the original view-was phased with a single I, indicating that useful inform:ition can be obtained from maps calculated with the aid of a single derivative.

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FIQ.6. (a) Top view of the area of the pocket from the 4-A electron density map of the Type I crystals; X marks the position of the 5-methyl of pdTp in the Type I1 crystals. (b) Same view of the pocket area. but now the 4-A electron density map of the Type I1 (inhibitor complex) crystals. The phasing is based on a single I.

Asp 21, Asp 40, and Glu 43. However, it appears almost certain that the Ca‘+ is too far from the phosphate for even a weak electrostatic bonding, nor have we found any indication of a bridging water molecule in the electron density map. The 5’-phosphate interacts with the guanidinium terminus of Arg 35, which in turn also appears to interact with the peptide carbonyl group of Leu 36 and perhaps with the peptide carhonyl of Val 39. The guanidinium group of Arg 87 also interacts on one side with the 5’-phosphate and on the other side with the carboxylate side chain of Asp 83, which in turn may form a bond to the peptide nitrogen of Gly 86. There may also he interaction between the guanidinium group and the oxygen atom of the furanose ring of the deoxyribose. The T y r 113 ring plane is parallcl to thc pyrimidine ring of pdTp but probably too distant to warrant a postulate of any significant ring-ring interaction. However, it does appear that its phenolic hydroxyl group is in position to interact with the oxygen of the 5’-phosphate ester bond. The %-phosphate group appears definitely to interact with the phenolic hydroxyl group of T y r 85 and possibly, but less clearly, with the c-amino group of Lys 84. The lining of the pocket is formed by a complex intertwining of the

(b)

FIG.7. (a) View of the electron density map shown in Fig. 6b as if looking from the right side of the Styrofoam model of Fig. la, but looking only at the added electron density in the pocket region and on the left side of the pocket. I, determined position of the iodine atom in the crystals containing 5-iododeoxyuridine diphosphate analog of pdTp; B and R, the pyrimidine ring and deoxyribose moiety of pdTp, respectively. The positions of the 3’- and 5’-phosphates and the Ca2+ are also shown. T, a group thought a t the time to be the benzene ring of a tyrosine residue, later identified as Tyr 113. Again the heaviest lines are meant to show the nearest groups (67). (b) A space-filling model of pdTp.

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STAPHYLOCOCCAL NUCLEASE X-RAY STRUCTURE

(01

(b)

Fro. 8a and b.

169

170

F. A . COTTON A N D E. E. HAZEN, JR.

FIG.8c and d.

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STAPHYLOCOCCAL NCCLEASE X-RAY STRUCTURE

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side chains of hydrophobic and neutral residues with the main peptide chain. The exception to this is Asp 83 lying on the upper right side, but this residue is tightly folded back along the main chain and probably has its charge partially neutralized by Arg 87. We must not fail to mention that the c-amino of Lys 71 from a neighboring nuclease molecule in the unit cell appears to interact with the 5’-phosphate. There is no wholly adequate substitute for a detailed study of the three-dimensional electron density map, but Figs. 8a-e show two-dimensional projections of interactions discussed above.

FIG.8. (a) The side view of the pdTp from the high resolution map, this time with the density associated with Tyr 113 omitted. R, deoxyribose; B, the pyrimidine ring. The labels indicate approximate position of the side chains of the indicated residues. (b) View of the pocket from the top in the plane of the calcium ion with the positions of some side chains indicated. Again the dark lines are densities nearest the eye. The depth of the section is 5.1 A. (c) View of the pocket from the top in the plane of the %-phosphate. The depth of the section is 5.1 A. (d) View of pocket from the top cut off just above the deoxyribose and below the 3’-phosphate. The depth of the section is 3.2 if. R, deoxyribose; B, the pyrimidine ring. (e) View of the pocket from the top running from just below the 3’-phosphate to the top of the enzyme molecule. The depth of the section is 3 8 A. B, top of the pyrimidine ring.

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IV. Some Correlations with Studies in Solution

From differences between the spectral (29) and fluorescence (30)properties of the inhibited and uninhibited nuclease and from differences in the number of groups susceptible to acetylation (32) in the two forms of the enzyme, Cuatrecasas et al. (33) concluded that tyrosyl residues were involved in binding pdTp. This led them to some very interesting studies of the specific modification of certain tyrosyl residues with tetranitromethane and of the properties of these modified forms of the nuclease which we will discuss below in the context of this paper. Briefly, the pattern of relative reactivity was found to be Uninhibited nuclease: Tyr 85 >> 54 2 113 > 27 >> 115 Inhibited nuclease: Tyr 115 >> 54 2 113 >> 85 ‘v-27

Tyrosine 91 and 93 are not nitrated even a t high molar excess of reagent unless the enzyme is denatured. Tyrosine 85 and 115 are so highly reactive that it proved possible to isolate nuclease derivatives in which only Tyr 85 was nitrated (NT,,Nuc) or only T y r 115 was nitrated (NT,,,Nuc) and, by reduction, the corresponding aminotyrosine derivatives (AT,,Nuc and ATlISNuc). First of all let us deal with the question of the high reactivity of Tyr 85 in the uninhibited nuclease and of Tyr 115 in the inhibited. One of the characteristics of electron density maps of protein structures (and information that is not available in models) is that the clarity with which amino acid side chains can be seen varies considerably. Typically the poorly resolved side chains are charged and extend out from the molecular surface. In the high resolution structure (inhibited molecule) the side chains of Tyr 27, 91, 93, 85, and 113 are very well resolved, that of Tyr 54 is only slightly less clear, but that of Tyr 115 is very diffuse and apparently highly exposed, lying above and to the rear of Tyr 113 (see Figs. 8d and 8e for the positions of Tyr 113 and 115). Although we do not yet have a high resolution map of the uninhibited nuclease, it is plausible to suggest that here, with no 3’-phosphate group to which a hydrogen bond can be formed, Tyr 85 may be similarly highly exposed, lying as it does a t the tip of an exposed loop on the extreme top right front of the nuclease. Thus, the high reactivity of T y r 85 and 115 may be ascribed to their being very highly exposed. This view agrees well with the spectral and titration behavior observed in the above-cited paper. It is easy to see why Tyr 85 is unreactive in the inhibited nuclease; it is tightly hydrogen bonded to the 3’-phosphate of the pdTp. I n harmony

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173

with this, it is found that NT8,Nuc has lost its enzymic activity and also binds 5’-AMP more weakly. Since 3-nitrotyrosine has a pK of 6.8 (58) and Tyr 85 can thus no longer form a good hydrogen bond to the 3‘phosphate a t p H values substantially above 6.8, the substrate can probably not be positioned in the active site correctly; consequently, the enzyme is no longer active ( 5 9 ) .I n ATg5Nucsome of the enzymic activity is restored which is consistent with a pK of the phenolic hydroxyl of 10 in 3-aminotyrosine (61). Incomplete restoration of activity in AT,,Nuc (and perhaps part of the loss of NT,,Nuc) can probably be ascribed to some sort of steric factor. The reduction of 5’-AMP binding is more difficult to explain, but since in NTlISNuc Tyr 85 is no longer easily nitratable, it is possible to postulate a hydrogen bond between T y r 115 and 85 when one or the other is nitrated, thus blocking access to the pocket. If our inference based on the 4-A maps of the Type I and Type I1 crystals (31) that in the uninhibited nuclease the side chains of T y r 113 and 115 are folded into the pocket is correct, it is possible to explain the resistance of T y r 115 to nitration. Indeed, if the peptide chain in this region pivots as we have surmised it does, then in the uninhibited nuclease the side chain of Tyr 115 should lie fairly deep within the pocket, conceivably in a position to bond to the carboxylate of Gln 80 or possibly that of Asp 83; Tyr 113 would be toward the front of the pocket and more accessible. NT,,,Nuc retains full activity toward DNA but loses about 50% of its activity toward RNA. The 2’-hydroxyl of ribonucleotides, if bound in the same stereochemistry (62) as pdTp, would be favorably situated for interaction with the side chain of T y r 115, such an interaction being enhanced by the low pK of a nitrotyrosyl 115 residue. In the high resolution structure, Tyr 91 and 93 are indeed “buried” deep in the hydrophobic area between the back wall of the pocket and the rear edge of the nuclease molecule. Tyrosine 27, which is more resistant to nitration in the inhibited nuclease, is probably hydrogen bonded to the carboxylate of Glu 10, partially shielded by the methylene chain of Lys 28, and, most importantly, at the rear upper right 58. J. F. Riordan, M. Sokolovsky, and B. L. Vallee, Biochemistry 6, 358 (1967).

59. The basic structural unit nerded for activity is R-pdT-R. R may be a free 3’-OH but a phosphate here markedly increases substrate binding. R may bc a p nitrophenyl group. The bond that is broken is the P-0 of the 5’-C-0-P ester linkage (60). 60. P. Cuatrecasas, M. Wilchek, and C. B. Anfinsen, Biochemistry 8, 2277 (1969). 61. M. Sokolovsky. J. F. Riordan, and B. L. Vallee, BBRC 27, 20 (1967). 62. Preliminary results with crystals containing 5’-AMP indicate that the binding positions are very similar.

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corner of the molecule far from the binding site. 1t7e are looking forward to finding out whether this reactivity change results from a conformational change transmitted from the pocket through a mass of protein or whether T y r 27 is close to one of the other binding subsites that the work of Cuatrecasas et al. indicated are present (63). As recently discussed by Cuatrecasas, some of the work with affinity labels, which are based on modification of pdTp, supports the subsite hypothesis ( 6 4 ) . The hydrolysis of dinucleotides by the staphylococcal iiuclease hab been recently studied in Laskowski's laboratory ( 1 2 ) .In 12 dinucleotides of the type d-NapNflp the susceptibility to hydrolysis varied about 100fold and appeared t o depend primarily on the type of base in the /3 position. In addition, the corresponding d-NpN dinucleosidemonophosphates were more resistant to hydrolysis, and the pNpN 5'-phosphodinucleotides were most resistant of all. This work would appear to be consistent with our view of the pdTp binding site of the nuclease a t high resolution.

V. Some Tentative Comments on Mechanism and Plans for Future Studies

A. A WARNING I n our high resolution structure of the nuclease-pdTp-CaZt complex, we find but a single calcium ion. The gel filtration studies of pdTp and CaZt binding and of calcium binding to the nuclease in the presence of nucleic acid indicate that there are probably a t least two sites of binding (28). These findings are not necessarily inconsistent. They may merely reflect the fact that the crystals prefer t o grow under conditions different from those used for the binding studies. Further studies on this point are in order.

B. MECHANISM It would be attractive to speculate that as part of the enzymic reaction T y r 113 donated a proton to the 5'-phosphate ester bond (59). Since the enzyme is active when T y r 113 is nitrated (33) and thus almost certainly present as a phenolate ion a t the optimum pH, this idea 63. P. Cuatrecasas, M. Wilchek, and C. B. Anfinsen, Science 162, 1491 (19668) 64. P. Cuatrecasas, JBC 245, 574 (1970).

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is doubtful. However, it is not unreasonable to think then in terms of a nuclcophilic attack or a stabilization of a transition state by this ionized tyrosine. Again, further discussion had best await further work.

C. PROPOSED FUTURE STUDIES Work is well under way on the high resolutioii structure of the uninhibited nuclease as are attempts to prepare crystals with other nioiio-, di-, and trinucleotidcs of various types so that more insight may be gained into the actual workings of the nucleasc. As described clscwhere in this volume ( 2 5 ) , some of the most exciting prospects for the future arc thc correlations to hc made t)ctn.cen the crystallographic studies of the threc-dimensional structure and the properties of the fragmented and synthetic versions of this nuclease.

Note: Viewer for color stereo figures can be found in back pocket. Green lens, right eye; red lens, left eye.

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Staphylococcal Nuclease, Chemical Properties and Catalysis CHRISTIAN B. ANFINSEN HIROSHI TANIUCHI

PEDRO CUATRECASAS

I. Introduction . . . . . . . . . . . 11. Isolation . . . . . . . . . . . . 111. Covalent Structure . . , . . . . . . IV. Behavior in Solution . . . . . . . . . V. Substrate Specificity and Catalytic Mechanisms . . . A. Polynucleotide Substrates . . . . . . B. Synthetic Substrates and Inhibitors . . . . C. Recapitulation-Size and Specificity of the Active Site VI. Stereochemical Probes of the Active Site . . . . VII. Complementation of Fragments . . . . . . VIII. Synthetic Analogs . . . . . , . .

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177 178 180 183 185 185 187 191 195 196 199

1. Introduction

Staphylococcal nuclease was discovered by Cunningham and his colleagues in cultures of pathogenic strains of Staphylococcus aureus (1,8). It was the first nuclease to be found that yielded 3’-nucleotides upon hydrolysis of polynucleotide chains (3-16). Historically, this feature of 1. 2. 3. 4. 5.

L. Cunningham, B. W. Catlin, and M. Privat de Garilhe, JACS 78, 4642 (1956). L. Cunningham, Ann. N . Y . Acad. Sci. 81, 788 (1959). P. Cuatrecasas, S.Fuchs, and C. B. Anfinsen, JBC 242, 1541 (1967). K . K . Reddi, Nature 187, 74 (1960). K . K . Reddi, Nature 182, 1308 (1958). 177

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C. B. ANFINSEN, P. CUATRECASAS, AND H. TANIUCHI

its catalytic action was of considerable use in the classic studies of Kornberg and his collaborators in the determination of nearest neighbor base frequencies in synthetic deoxyribonucleic acid (DNA) ( 1 7 ) . A radioactive phosphorus atom esterified to the 5‘-OH group of deoxyATP, for example, would be located, after staphylococcal nuclcase digestion of the newly synthesized DNA, on the 3‘-mononucleotide, dXp, derived from the sequence dXpdA-. The enzyme has also h e n used extensively in studies on the elucidation of sequences of polynucleotidc chains (18). In addition to interest in this enzyme because of its catalytic characteristics, a considerable body of information has accumulated on staphylococcal nuclease as a protein molecule. I t s relatively small size, the absence of covalent cross-linkages, and its behavior upon binding a variety of ligands have made it an ideal model substance for the study of various aspects of protein chemistry including X-ray crystallography. These investigations are reviewed in the present chapter and in Chapter 7 by Cotton and Hazcn, this volume, on the thrce-dimcnsional structure (19). II. Isolation

Early studies on the purification of staphylococcal nuclease led to conflicting reports on whether the enzyme could hydrolyze both DNA ~~~

~

~

6. M. Privat de Garilhe, G. Fassina, F. Pochon, and J. Pillet, Bull. Sac. Chim. Biol. 40, 1905 (1958). 7. F. Pochon and M. Privat de Garilhe, Bull. Sac. Chim. B i d . 42, 795 (1960). 8. K. K. Reddi, BBA 36, 132 (1959). 9. K. K. Reddi, BBA 47, 47 (1961). 10. A. Ohsaka, J.-I. Mukai, and M. Laskowski, Sr., JBC 239, 3498 (1964). 11. M. Alexander, L. A . Heppel, and J. Hurwits, JBC 236, 3014 (1961). 12. G. W. Rushizky, C. A. Knight, W. K . Roberts, and C. A. Dekker, BBRC 2, 153 (1960). 13. E. Sulkowski and M. Laskowski, Sr., JBC 237, 2620 (1962). 14. W. K . Roberts, C. A. Dekker, G. W. Rushizky, and C. A. Knight, BBA 55, 664 (1962). 15. G. W. Rushizky, C. A. Knight, W. K. Roberts, and C. A . Dekker, BBA 55, 674 (1962). 16. R. Hacha and E. Frederico, BBA 123, 493 (1966). 17. J. Josse, A. D. Kaiser, and A. Kornberg, JBC 236, 864 (1961). 18. G. von Ehrenstein, in “Aspects of Protein Biosynthesis” (C. B. Anfinsen, ed.), p. 139. Academic Press, New York, 1970. 19. F. A. Cotton and E. E. Hazen, Chapter 7, this volume.

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and ribonucleic acid (RNA). I t is now clear, however, that the cleavage of polynucleotides by nuclease is only quantitatively affected by the nature of the sugar moiety in the substrates ( 3 , 2 0 ) . The isolation of highly purified nuclease has also received special attention because of the desirability of having availahle, for polynucleotide sequence studies, enclonucleases free of phosphatase activity. When the enzyme is prepared from a crude ammonium sulfate fraetion by gradient elution from CM-cellulose columns, considerable levels of both acid and alkaline phosphatnse activities remain in the final product ( 2 1 ) . These activities may be removed by chromatography on phosphocellulose, on which the phosphatases are more highly retarded during elution with a salt and pH gradient ( 2 1 ) . The relative activities against RNA and DNA remain constant across the elution profile from both CM-cellulose and phosphocellulose, and this, together with other evidence described below, indicate that both activities result from a single species of enzyme molecule. Mikulski et 01. ( 2 2 ) have employed a combination of phosphocellulose chromatography and gel filtration in the final stages of an alternative preparation of phosphatase-free nuclease. The extremely high affinity of phosphocellulose for nuclease at pH values below 6.5 has formed the basis of a convenient method for the isolation of large amounts of enzyme from cultures of Staphylococcus aureus without the need for direct handling of this potentially dangerous pathogen ( 2 3 ) . The entire culture, in a 300-liter fermenter, is stirred with the cation exchanger a t pH 5.5-6.5 and, after the phosphocellulose with the attached nuclease has settled to the bottom and the supernatant fluid has been siphoned off, remaining bacterial cells may be removed by repeated suspension in distilled water. Subsequent purification to a state of homogeneity may then be carried out by gel filtration on Sephadex G-75 followed by phosphocellulose chromatography of an ammonium sulfate fraction of the total adsorbed protein, or on a smaller scale by the application of the principle of affinity chromatography using Sepharose-inhibitor columns ( 2 4 ) . 20. C. B. Anfinsen, M. K. Rumley, and H . Taniuchi, Acta Chem. Scand. 17, 270 (1963). 21. S. Fuchs, P. Cuatrecasas, and C. B. Anfinsen, JBC 242, 4768 (1967). 22. A . J. Mikulski, E. Sulkowski. L. Stasiuk, and M. Laskowski, Sr., JBC 244, 6559 (1969). 23. L. Morkvek, C. B. Anfinsen, J. L. Cone, and H. Taniuchi, JBC 249, 497 (1969). 24. P. Cuatrecasas, M. Wilchek. and C. B. Anfinsen, Proc. Nntl. Acad. S C ~u. . 61, 636 (1968).

s.

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C. B. ANFINSEN, P. CUATRECASAS, AND H. TANIUCHI

111. Covalent Structure

Nuclease is composed of 149 amino acid residues, and contains neither sulfhydryl groups nor disulfide bonds (20, 25, 267. Figure 1 shows the amino acid sequence of nuclease isolated from Staphylococcus aureus, strain V8 (20, 25-27). Physical measurements of the molecular weight of nuclease from strain V8 as well as from strain “Foggi” are consistent with this sequence (28). The determination of the amino acid sequence of nuclease was approached by preparing large fragments by BrCN cleavage (29) and arranging the fragments in linear order (25). Nuclease contains four methionine residues and essentially quantitative cleavage of the methionine bonds by BrCN yields five fragments; [l-261, [27-321, [33-651, [66-981, and [99-1491 (the numbers in the brackets indicate the residues contained in the fragment). The COOH-termini of the first four 1

10

10

FIG.1. The amino acid sequence of nuclease V8 (25-2’7). Residue 43 is revised as glutamic acid (H. Taniuchi and C. B. Anfinsen, unpublished results; see also text). 25. H. Taniuchi and C . B. Anfinsen, JBC 241, 4366 (1966). 26. H. Taniuchi, C. B. Anfinsen, and A. Sodja, JBC 242, 4736 and 4752 (1967). 27. H. Taniuchi, C. L. Cusumano, C. B. Anfinsen, and J. L. Cone, JBC 243, 4775 (1968).

28. F. A. Cotton, E. E. Hazen, Jr., and D. C. Richardson, JBC 241, 4389 (1966). 29. E. Gross and B. Witkop, JBC 237, 1856 (1962).

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peptides are, of course, homoserine or the lactone form of this amino acid. Separation of the BrCN fragments was best achieved by gel filtration on a long column of Sephadex G-50 (25) with further separation of [ 1-26] and [ 33-65] by continuous flow electrophoresis. Each cyanogen bromide fragment was digested with trypsin and the tryptic peptides were fractionated by ion exchange chromatography on a Dowex 50 column, using gradient elution with pyridinium acetate buffer (30). Further purification of these tryptic peptides was performed by paper chromatography, paper electrophoresis, or the combination of these two methods (two-dimensional peptide mapping) (31). Four tryptic peptides containing intact methionine residue were obtained from the tryptic digest of intact nuclease. Amino acid analysis (32) and NH?-terminus determination by the dinitrofluorobenzene method of the BrCN fragments, the tryptic peptides obtained from the BrCN fragments and the four from native nuclease provided the information required to deduce the linear arrangement of the BrCN fragments ( 2 5 ) . The tryptic peptides of the BrCN fragments were sequenced by Edman degradation using the subtractive method (33) together with identification of the phenyl thiohydantoins (PTH) of the amino acids (34, 3 5 ) . Amide groups were determined by examining the electrophoretic mobility, a t pH 6.5, of peptides on paper or by digesting peptides with leucine aminopeptidase followed by amino acid analysis. Ascending paper chromatography with 80% pyridine as solvent was sometimes used to separate glutamine or asparagine released by leucine aminopeptidase. The collidine-ninhydrin reagent stains glutamine and asparagine differentially (36). Arrangement in order of tryptic peptides derived from the BrCN fragments was based on the structures of overlapping chymotryptic peptides, which were separated and partially sequenced in a manner similar to that described above (26). Residues 30 and 31, glutamine and proline, respectively (27), were originally assigned in the reversed order on the basis of partial removal of residue (homoserine) from cyanogen bromide fragment [ 27-32] by carboxypeptidase A ( 2 6 ) . The action of the latter is generally as30. R. E. Canfield and C. B. Anfinsen, JBC 238, 2684 (1963). 31. A. Katz, W. J. Dreyer, and C. B. Anfinsen, JBC 234, 2897 (1959). 32. D. H. Spackman, S. Moore, and W. H. Stein, Anal. Chem. 30, 1190 (1958). 33. C. H. W. Hirs, S. Moore, and W. H. Stein, JBC 235, 633 (1960). 34. P. Edrnan and J. Sjoquist, Acta Chem. Scand. 10, 1507 (1956). 35. H. 0. Van Orden and F. H. Carpenter, BBRC 14, 399 (1964). 36. R. E. Canfield and C. B. Anfinsen, in “The Proteins” (H. Neurath. ed.), Vol. 1, p. 311. Academic Press, New York, 1963.

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C. B. A N F I N S E N , P. CUATRECASAS, AND H. T A N I U C H I

sumed to be effectively blocked by a proline residue, though the ahsumption is not always valid (37). Glutamic acid (see the legend to Fig. 1) was originally proposed as glutamine on the basis of apparent ncutrality during electrophoresis a t pH 6.5 of tryptic peptide 136-45 I (26). The unusual array of four hydrophobic residues (leucincs 36, 37, and 38 and valine 39) caused serious tailing of the spot on paper during paper electrophoresis, giving an ambiguous mobility. Removal of the four hydrophobic residues by leucine aminopeptidase from peptide [ 36-45 yielded peptide [40-45] with an unequivocal acidic mobility (38). The amino acid sequence of nuclease Foggi was also studied because this strain, Staphylococcus aui-eus, yields much more nuclease in culture (23, 3 9 ) . The close similarity of the covalent structures of nuclease V8 and Foggi was early indicated by immunological studies (%‘I), physicochemical characterization (40), amino acid analysis, and the two-dimensional peptide map of tryptic digests ( 4 1 ) . The availability of substantial quantity of nuclease Foggi was fundamental in the carcful reexamination of ambiguous points of amino acid sequcncc as described above and for the structural and functional studies of nuclease, including X-ray crystallography. Elucidation of the amino acid sequence of nuclcase Foggi was approached in the same manner as that of nuelease V8. Two fragmcnts (residues 6-48 and 49-149) obtained by the limited trypsiii digestion (see below) of nuclease were also used to make cyanogen bromide fragments. The sequential Edman-dansyl chloride method (42) was mainly employed for rapid sequencing of tryptic peptides (41) instead of the subtractive Edman method or PTH-amino acid identification. The amino acid sequences of nuclease V8 (Fig. 1) and Foggi (41, 43) arc the same with the exception of residue 124, which is leucine in V8 (26) and histidine in Foggi nuclease (41). Replacement of leucine 124 by a histidine residue appears to have no effect on the structure and function of the enzyme (27, 28, 40). A methionine auxotroph (M-) of Staphylococcus aureus also appears to yield nuclease with a covalent structure similar to that of nuclease 37. J. I. Harris and C. H. Li, JBC 213, 499 (1955). 38. H. Taniuchi and C. B. Anfinsen (see reference 4 3 ) . 39. J. N. Heins. H. Taniuchi, and C. B. Anfinsen, in “Methods in Nucleic Acid Rcsearch” (J. Cantoni and D. Davies, eds.), p. 79. Harper, New York, 1966. 40. J. N. Heins, J. R. Suriano, H . Taniuchi, and C. B. ilnfinsen, JBC‘ 242, 1016

(1967). 41. C. I,. Cusumano. H. Taniuchi, and C. R. Anfinsen, JBC 243, 4769 (1968). 42. W. R. Gray, “Methods in Enzymology,” Vol. 11, p. 139, 1967. 43. J. L. Cone, C. L. Cusumano, H. Taniuchi, and C. B. Anfinsen, JBC (it1 prcsd (1971).

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V8, as indicated by two-dimensional peptide maps of tryptic digests (44). It appears, therefore, that various strains of S. aureus produce a major extracellular nuclease with covalent structures extremely similar to that of nuclease V8. Staphylococcus aureus has been reported, however, to produce several minor extracellular nucleases with different isoionic points and heat stabilities ( 4 5 ) . The covalent structures of these materials have not been studied.

IV. Behavior in Solution

Nuclease behaves like a typical globular protein in aqueous solution when examined by classic hydrodynamic methods (40) or by measurements of rotational relaxation times for the dimethylaminonaphthalene sulfonyl derivative ( 4 6 ) . Its intrinsic viscosity, approximately 0.025 dl/g is also consistent with such a conformation. Measurements of its optical rotatory properties, either by estimation of the Moffitt parameter O,,,or the mean residue rotation at 233 nm, indicate that approximately 1518% of the polypeptide backbone is in the a-helical conformation (47, 4 8 ) . A similar value is calculated from circular dichroism measurements (48). These estimations agree very closely with the amount of helix actually observed in the electron density map of nuclease, which is discussed in Chapter 7 by Cotton and Hazen, this volume, and Arnone et al. ( 4 9 ) . One can state with some assurance, therefore, that the structure of the average molecule of nuclease in neutral, aqueous solution is a t least grossly similar to that in the crystalline state. As will be discussed below, this similarity extends to the unique sensitivity to tryptic digestion of a region of the sequence in the presence of ligands (47, 48), which can easily be seen in the solid state as a rather anomalous protrusion from the body of the molecule (19,4 9 ) . When the protein is exposed to pH values below 3.5-3.7, temperatures 44. C. B. Anfinsen and L. G . Corley, JBC 244, 5139 (1969). 45. T. Wadstrom. B B A 147, 441 (1967). 46. P. Cuatrecasas, H. Edelhoch. and C. 13. Anfinsen, Proc. N u l l . Acntl. Sca. U . S. 58, 2043 (1967). 47. H. Taniuchi, C. 13. Anfinsen. and A. Sodja, PTOC.&'all. Acod. Sci. c'. S. 58, 1235 (1967). 48. H. Taniuchi and C. B. Anfinsen, JBC 243, 4778 (1968). 49. A. Arnone, C. J. Bier, F. A . Cotton. E. E. Hmen. D. C. Richardson, nnd J . S. Richardson, Proc. Nntl. Acad. Sci. U . S.69, 420 (1969) ; see also Chapter 7, this volume.

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C. B. ANFINSEN, P. CUATRECASAS, AND H. TANIUCHI

in the range of 55”-65”, high concentrations of urea, guanidinium chloride, and certain organic solvents, it rapidly unfolds. This denaturation is completely reversible. Measurements of the rate of refolding of nuclease by stopflow measurements of the enhancement of tryptophan fluorescence after changing the pH from 3.4 to 6.5 indicate that most of the refolding process takes place with a half-time of about 45 msec, with a somewhat slower secondary “readjustment” process having a halftime of about 200 msec (50). Hydrodynamic, spectroscopic, and optical rotatory methods can “see” only the average molecule and a fraction of the population as large as 5 or 10% that differs significantly in shape might not be detected. On the other hand, measurements of hydrogen exchange, or of susceptibility to proteolysis of proteins in a relatively infrequent, transient state of partial denaturation, would detect such deviant molecules. Although native nuclease, like most proteins in solution, behaves, on the average, as though all the molecules were globular, this phenomenon of motility becomes apparent when the protein is examined by such “historical” methods. The exchangeable hydrogen atoms of nuclease can be shown to be completely susceptible a t room temperature to equilibrium with the hydrogen atoms of the aqueous solvent in one hour or less (51, 5 6 ) . Variations in the structure of the protein must therefore permit the rapid entrance and exit of bulk solvent during the extremely brief deviations from the rather tightly packed geometry indicated by the crystallographic studies. This interpretation is supported by the ligand-induced resistance to hydrogen exchange. When pdTp and calcium ions are present, approximately 35 hydrogen atoms become inaccessible to solvent for many hours in spite of the fact that only a slight and localized change in structure can be detected crystallographically (see Chapter 7 by Cotton and Hazen, this volume) in the nuclease-pdTp-Ca2+ complex. The results indicate that infrequent unfolding and refolding must be occurring in the native protein in solution and that only when the structure is “rigidified” by ligand addition is this motility suppressed. Similar conclusions may be reached from the study of the ligandinduced resistance to proteolysis (47, 5 3 ) , a phenomenon that has been employed in the production of large complementing fragments of nuclease (see below). 50. A. N. Schcchter, R. F. Chen, and C. B. Anfinsen, Science 167, 886 (1970). 51. A. N. Schechter, L. MorBvek, and C. B. Anfinsen, Proc. Natl. Acad. Sci. u. s. 61, 1478 (1968). 52. A. N. Srhechter, L. Morivek, and C. B. Anfinsen, JBC 244, 4981 (1969). 53. H. Taniuchi, L. Morivek, and C. B. Anfinsen, JBC 244, 4600 (1969).

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V. Substrate Specificity and Catalytic Mechanisms

A. POLYNUCLEOTIDE SUBSTRATES 1. Specificity

Staphylococcal nuclease is a phosphodiesterase which can cleave either DNA or RNA to produce 3’-phosphomononucleosides (3-16, 2 0 , 2 5 ) . The rates of hydrolysis of these substrates are dependent on the conformation of the substrate, Ca2+concentration, and the ionic strength and the pH of the buffer ( 3 , 5 4 ) . Denatured D N A is hydrolzed more rapidly than native DNA ( 3 , 12, 64-56), which reflects the important effect of substrate conformation on catalysis. I n native DNA the X p d T p and X p d A p bonds are preferentially attached (7, 12, 14, 15, 5 5 ) . With denatured D N A the order of cleavage appears to be nearly random (14, 15, 56, 57‘).The Xp-dCp and Xp-dGp linkages in the helical regions of DNA, which are more cxtensively stabilized, are more resistant to hydrolysis. The specific order of release of various mononucleotides from native compared to denatured DNA suggests that in the hydrolysis of D N A specificity toward the constitutent bases is less important than the substrate conformation (64-57). The mechanism of catalysis of macromolecular substrates is relatively complicated. The initial phases of digestion arc predominantly ( 4 , 8, 13, 58, 69),but not exclusively ( 1 1 , 58, 5 9 ) , endonucleolytic. These initial cleavages are probably followed by exonucleolytic cleavages of the same strand ( 5 9 ) . Subsequent scission of the exposed single-stranded region occurs, followed by unwinding of the region proximal to the scission. These interpretations are consistent with the marked changes in hyperchromicity, and the low di- to mononucleotide ratio, in the initial phases of digestion when the principal products are mononucleotides and long fragments (59). The cleavage pattern becomes progressively exonucleolytic as the reaction progresses, becoming principally exonucleolytic on the hexanucleotide level ( 1 1 , 13, 6 0 ) . Exhaustive digestion of deoxyribopolynucleotide substrates results in the release of 3’-mono54. E. Sulkowski and M. Laskowski, Sr., JBC 243, 4917 (1968). 55. M. I,. Dirksen and C. A. Dekker, BBRC 2, 147 (1960). 56. P. H. von Hippel and G . Felsenfeld, Biochemistry 3, 27 (1964). 57. L. Wingert and P. H. von Hippel, BBA 157, 114 (1968). 58. E. J. Williams, S. C. Sung, and M. Laskowski, Sr., JBC 236, 1130 (1961). 59. E. Sulkowski and M. Laskowski, Sr., JBC 244, 3818 (1969). 60. M. de Meuron-Landolt and M . Privat de Garilhe, BBA 91, 433 (1964).

186

C. B. ANFINSEN, P . CUATRECASAS, AND H . TANIUCHI

nucleotides, dinucleotides, and a 3,5’-nucleosidediphospliatc (the left tcrminus) (10, 11, 2 2 ) . Contrary to early reports (11, 13, 59),dinucleotides can be hydrolyzed by micrococcal nuclense (22, 61). No hydrolysis of 2’,3’-cyclic phosphate esters has been observed (11), and no evidence exist$ to suggest that the mechanism of hydrolysis proceeds through cyclic phosphate intermediates. Enzymic activity is inhibited by oligonucleotides or ~ i ~ o ~ i o ~ i ~ ~ c l e o t i c l e ~ bearing 5’-phosphate end groups (3, 11). The presence of a 3’-phosphate end group enhances hydrolysis of the adj accnt internucleotide bond in oligonucleotides (11, 22) and synthetic substrates (61). The enzyme shows preference for adenine and thymine over cytosine and guanine, as judged by (1) the rate of inononuclcotide release from I)NA, polynucleotidcs, and oligonucleotides (1, 10, 11, 13); ( 2 ) the pattern of hydrolytic susceptibility of oligo- and dinucleotides (11, 13, 22) ; and (3) the binding affinities of 5’-mononucleotides (61). 2. Kinetic Measurements The rate of hydrolysis of DNA, RNA, and polynucleotides can be measured by a sensitive spectrophotometric assay which is based on the hyperchromicity that occurs upon hydrolysis of these substrates (3). The enzyme has a 7-fold greater affinity for denatured D N A than for RNA (3).No inhibitory products accumulate during the course of the reaction. The p H optimum for RNase and DNase activities is between 9 and 10, depending on the Caz+concentration. At higher pH values less Ca2+is required. The inhibitory effect of high Ca2+observed consistently by many investigators is more pronounced a t higher pH values ( 3 ) . Enzymic activity appears to be completely dependent on the presence of Caz+ (1, 3, 4, 6, 11). There are no significant changes i n Ca‘+ requirements when the amounts of DNA or RNA are varied, suggesting that Ca2+is not primarily involved in producing conformational changes in the substrate. This is consistent with the observation that Ca2+ is required for the binding of nucleotide inhibitors to the enzyme active site. This has been confirmed by direct physical measurements (62) and by changes in the ultraviolet spectra (6S), fluorescence emission spectra (46), circular dichroic spectra ( 6 4 ) , and antigenic reactivity (6‘6) 61. P. Cuatrecasas, M . Wilchek, and C. B. Anfinsen, Biochemistry 8, 2277 (1969). 62. P. Cuatrecasas, S. Fuchs, and C. B. Anfinsen, JBC 242, 3063 (1967). 63. P. Cuatrecasas, S. Fuchs, and C. B. Anfinsen, JBC 242, 4759 (1967). 64. G. S. Omenn, P. Cuatrecasas, and C. B. Anfinsen, Proc. Natl. Acad. Sci. U. S. 64, 923 (1969). 65. S. Fuchs, P. Cuatrecasas, D. A. Ontjes, and C. B. Anfinsen, JBC 244, 943

(1969).

8.

STAPHYLOCOCCAL NUCLEASE

187

which occur upon binding of thesc inhibitors to the cnzyinically activr site. The location of a tightly bound Ca"+ (or Ba'+) i i i crystallinr nucleasr has also been detcrmined hy crystallographic methods ( 1 9 , 4 9 ) . Considerable DNase but no RNasc activity rcsults if CaS+is replaced by S f + , while Fe"+ and Cu"+ causc minimal activation ( 3 , 4 0 ) . A number of heavy metal cations inhibit DKase and RNasc activities competitivcly with Ca"; Hg", Zn2+,and Cd" are the most potent of thesc (3). Studies with synthetic substrates, to be discussed below, indicatc that Ca'+ is not only required for the proper lincliiig of substrates but also that it is required for thc sul)scqucnt independent hydrolytic process. Although several divalent cations can substitute for Ca2+ in the binding function, as evidenced hy their competitive inhibition of enzymic activity ( 3 ) and their ability to promote nuclcotitle binding ( 6 2 ) , the catalytic role of Ca'+ appears to be unique.

B. SYNTHETIC SUBSTRATES AND INHIBITORS 1. Specificity Studies of the cleavage patterns ant1 kinetic constants of a number of synthetic, low molecular weight substrates intlicatc that the basic structural element required for recognition a s substrate (R-pdN-R') (65a) need not be part of a polynucleotide chain ( T a b l e I ) (61). Cleavage of compounds of the general class R-pdTp-R' result in the release of R phosphate and dTp-R' (61). Hydrolysis occurs irrespective of the size of the substrate since it appears that the nature of the R group is unimportant as long as a dicster bond is present and since substitution of the 3'-OH is not required to achieve the maximal rate of catalysis (61). Cleavage occurs rapidly with compounds having as the R group a p aminophenyl, p-nitrophenyl, or methyl substituent. Although it is not known whether the base is an essential feature of a substrate, the nucleoside moiety is necessary since bis- and tris-pnitrophenyl phosphate esters are not hydrolyzed (61). The nature of the R' substitution on the 3'-OH clearly affects the affinity of the substrate or inhibitor for the enzyme, even though it does not affect the maximal catalytic rate constant (Table I). The importance of the 5'- and 3'phosphate groups in determining the affinity of inhibitors (3, 66) is consistent with the contribution of these groups to substrate affinity ( 6 1 ) . These effects result from the phosphoryl groups themselves rather than 65a. Here N stands for nuclcoside.

66. 1'. Cuatrecasas, H. Taniuchi, and C. B. Anfinsen, Brookhaven S y m p . B i d . 21, 172 (1969).

188

C. B . ANFINSEN, P . CUATRECASAS, AND H . TANIUCHI

TABLE I SYNTHKTIC SUI3STRATI~:S A N D INHInITORS O F STAPHYLOCOCCAL NUCLEASIP Kcat

Compound Ni tropheiiyl-pdl’ 3‘-0Acetyliii trophenyl-pd‘l’ Ni trophenyl-pdTp Ni trophenyl-pdTpnitrophenyl Nitrophenyl-pdTpdT Nitrophenyl-pdTpdTpnitrophenyl Ni trophenylpdTpmet hyl Aminopheny 1-pdT Met,hyl-pdTpni trophenyl Fluoro-pdT dTp-Nitrophenyl 5’-Sulfate-dTpni trophenyl 5'-O- Acetyl-dTpnitrophenyl 5’-Chloromethyl-pdTpnitrophenyl dTp-Fluoro PdT PdTP Sulfate-dT-sulfate pdTp-N itrophenyl pdTp-Aminophenyl PdTpdT

K,,,,, (M) (min-’) 2 . 2 x 10-3 2 . 1 x 10-3

9.1 9.2

1.0 x 4 . 2 x 10-3 6 . 4 x 10-3 1.7 X 9 . 6 X lo-‘ 4.6 x 5 . 9 x 10-5

10.6 10.5 0.1 9.3 10.2 0.03 10.9

Kib ( M )

Groiip released

Nitrophenyl phosphate Nitrophenyl phosphate Nitrophenyl Nitrophenyl Ni trophenol Nitrophenyl Nitropherryl Ni trophenol Nitrophenyl

phosphate phosphate phosphate phosphat,e phosphate

1.3 X

0.08

2 . 2 x 10-2 5.9 x 10-3

0.7 1.9

Aminophenyl phosphate Methyl phosphate Ni trophen ol Fluorophosphate Ni trophenol Nitrophenol

x

0.7

Ni trophenol

0.4

Nitrophenol

2.1

1 .7 X

10-3

10-

x 2.0 x 1.9

1.1 x 1.0 x 6.3 x

lo-‘ lo-’ 10-3 10-6 10-6 10-7

a All the values were determined in 0.05 M borate buffer (pH 8.8) and 10 mM Ca2+as described by Cuatrecasas et al. (61). Inhibition was competitive in all cases. Data from Cuatrecasas et al. (61).

from alteration or elimination of the free hydroxyl groups, since 0-acetyl substitution does not substantially affect the kinetic constants obtained with the substituted derivatives (Table I) (61). Apart from important similarities in the endo- and exonucleolytic properties of staphylococcal nuclease and other well-studied phosphodiesterases ( 6 7 ) ,those from snake venom and spleen, the basic structural substrate elements for these enzymes appear to be quite different 67. H. G. Khorana, “The Enzymes,” Vol. 5, p. 79, 1961.

8.

189

STAPHYLOCOCCAL NUCLEASE

(Fig.2). The staphylococcal enzyme may appear to be more akin in its mode of action to the spleen enzyme because they both hydrolyze DNA and RNA to 3’-nucleotides, whereas the venom enzyme releases 5’-nucleotides. However, their mode of action and specificity are quite different, and the structural requirements of the staphylococcal enzyme substrates are perhaps more nearly similar to those of the venom enzyme. The principal difference is that the staphylococcal enzyme cleaves the diester bond between the phosphate and the 5’-carbon of the sugar, whereas the venom enzyme cleaves on the other side of the phosphate, that is, between the phosphate and the nonspecific hydroxylic component of the diester bond. In contrast to both spleen and venom diesterases, the primary product released by staphylococcal nuclease hydrolysis is a derivative bearing a hydroxyl group (on the 5’ position) rather than a phosphoryl group. Therefore, the 3’-phosphoryl product formed from polynucleotide hydrolysis is a secondary consequence of such cleavage.

1

j

I

I R ’ O $ Y - O - c O

I I

I I

! 0I I I I

I OH

I

I I

I

i

1

0

I

o=~-o’

I I I I

.-.-.-<-.-.-.-.-. (C)

FIG.2. Proposed structural requirements for substrates of phosphodiesterases that hydrolyze DNA and RNA, those from (a) snake venom, (b) spleen. and (c) staphylococcus (R = thymine and R = p-nitrophenyl). The studies indicated for the venom and spleen enzymes are those suggested by Khorana (67) [data from Cuatrecasas et al. (61)l.

190

C. B. ANFINSEN, P. CUATRECASAS, AND H. TANIUCHI

2. Kinetic Measurements

Above pH 6 the release of p-nitrophenyl phosphate from synthetic substrates (Table I) results in a shift in the ultraviolet spectrum of the chromophore which may be followed spectrophotometrically (Fig. 3 ) . This forms the basis for a sensitive and convenient assay of this enzyme (61). Perhaps the substrate best suited for routine assay of enzymic activity is nitrophenyl-pdTp, since it has highly favorable kinetic constants. The dependence of the kinetic constants (Table I) upon the concentration of Ca2+,pH, and ionic strength is not the same for all the synthetic substrates shown on Table I. In general, however, with all these substrates the maximal velocities are achieved between pH 9 and 11, maximal affinity for substrate occurs between pH 7.5 and 8.5 (Fig. 4 ) , and inhibition of enzymic activity is observed with NaCl concentrations greater than 0.1 N . Similar dependence upon these parameters is seen when activities are measured with DNA and RNA (3). Various kinetic studies indicate that the K , values measured with the synthetic substrates are true dissociation constants Ks. Furthermore, the Ki constants of these substrates, obtained from their inhibition of DNase

240

260

280

300

320

340

360

380

400

X (mp)

FIG.3. Ultraviolet spectral changes resulting from hydrolysis of nitrophenylpdTp-nitrophenyl (32 p M ) by staphylococcal nuclease (14 p M ) . Thc spectrum a t zero time is that of the mixture of enzyme and substrate before addition of Ca". The reaction is started by addition of Ca'+ (10 mM) ; by 3 min the reaction is cornplete. Data from Cuatrecasas et al. (61).

8.

191

STAPHYLOCOCCAL NUCLEASE I

I

I

I

I

I

Molecular ionization Constants

5 -

2a 4 3 -

I

~ K E ,

7.0

PKE~

8.6

Km

I

1

I

I

I

I

6

7

8

9

10

II

PH

FIG.4. Dependenw of K , , , , and K,* on pH for the clcavagc of thr synthetic substrate nitrophenyl-pdTp by staphylococcal nuclcasc. Assays wcre prrforincd in 0.05 M borate buffer, 0.015 M C a C L

activity, are identical with the K,,,,,,, values derived from their hydrolysis. This suggests that these substrates are specifically bound to the same region of the enzyme which binds and hydrolyzes DNA. As with DNA and RNA (S), substrate affinity depends upon the Ca"+ concentration. The lowest li,,,, for nitrophenyl-pdTp-nitrophenyl occurs a t mM Ca'+, but for nitrophenyl-pdTp it is 2.5 iriM Ca2+.Apart directly (Fig. 4) ; maxitnal from its effect on K,", Ca2+increases the effects are observed with about 0.1 ill Ca"+. Again in analogy with studies using DNA and RNA (S), all the synthetic substrates show an apparent inhibition of activity when the conccntration of Ca'+ is raised above 0.05 M . This effect is solely the result of an increase in li, (Fig. 5 ) .

C. RECAPITULATION-SIZE ASD SPECIFICITY OF T H E ACTIVESITE The data available from studies of the actioii of nucleabe on synthetic substrates, oligonucleotidcs, and natural polynucleotidcs are consistent with the following generalizations and simplifications. I t appcars that

192

C. B. ANF INS E N, P. CUATRECASAS, AND H. TA N I U C H I

LOQ

[ca'']

I'Ic. 5 . Dcl)endrncc of kinetic constants on Ca'+ conccntratlon for t hr c1cnr:igi~ of p-nitroplienyl-pTpT by staphylococcal nuclrnse. Data from Cuatrccisas et ol.

(GI).

the basic structural clement of substrate specificity is R-pN, with relcasc of Rp upon hydrolysis. Thc nature of R is cssentially unimportant with regard to specificity. The affinity for substrates and inhibitors is affected by the size, charge, and nature of the substituent on the 3'-OH. Study TABLE I1 BINHING CONSTAHTS OF OI,IGON~:CI.~~OTII,IC-NUCLIEAS~~ COMPLEXES Dissociation constant"

AFob

(M1

(cal/mole)

1 . 9 x 10-4 6 . 3 x 10-7 9 . 3 x 10-8

-4,400 -8,500 -10,000 -9,500

Nucleotide

1.I 1.4 G.o 2.0 1.1 1.7

x x x x x x

10-7

10-7

-9,400

10-7

-8 ,,500

10-7

-9,000 -8,100 -6,500

10-6 10-5

K , obtaiiied by Dixon plots (Fig. 1 ) ; the value for N1'-pdTp-NP is Kn,(approximately), obtained from Liiicweaver-Burk plots under identical experimental conditions. AF" = ky' 111 K,. c p-Nitropheiiyl phosphate (NI') esters. Data from Cuatrecasas t l ul. (68). 68. P. Cuatrrcasns, M. Wilchek, and C. B. Anfinscn. Science 162, 1491 (1969)

8.

193

STAPHYLOCOCCAL NUCLEASE

.

of the dissociation constants of a progressively longer series of 5’phosphoryl oligothymidyl inhibitors (pdT) indicate that the optimal stability of the enzyme-nucleotide complex occurs with approximately a trinucleotide sequence (Table 11) (68).It can be postulated that three unequal phosphate binding “subsites,” principally electrostatic in nature, exist in the substrate binding region of the enzyme (Fig. 6) (68). The most important site is thought to coincide with the hydrolytic site, and the other two are presumed to exist to the “right” of this hydrolytic site. I n the case of inhibitors, the free 5’-phosphoryl group, owing to its strong anionic charge and properly placed adjacent nucleoside, is recognized by the hydrolytic region. I n this case [ (pdT).], nucleotide units other than that bearing the 5’-phosphoryl group are considered to be to the “right” of the hydrolytic site and would be expected to contribute principally to the affinity constant. The rationalization can then be made in the case of substrates so that R-pdTp-R’ derivatives, unlike the inhibitors, possess a cleavable phosphodiester bond on the 5’ position and will thus release Rp upon hydrolysis. The R’ group, as in the case of the 5’-phosphoryl group of inhibitors, should contribute solely in terms of affinity. As with the K , values of inhibitors, the K , values of substrates are very sensitive to the addition of ionic phosphate groups to the “right” of the hydrolyzable bond. Furthermore, in contrast to the K,,, constants, the Kcatconstants of the various deoxythymidyl 5’-phosphate ester substrates are essentially identical. These considerations are consistent with the observation that 5’phosphoryl oligo- and mononucleotides are inhibitory (3, 1 1 ) , that the presence of a 3’-phosphate end group enhances cleavage of the adjacent bond of dinucleotides and oligonucleotides (11, 2 2 ) , and that endonucleolytic cleavage will readily occur in relatively random fashion in large oligonucleotides and polynucleotides. Although the contribution of the purine and pyrimidine bases is considered to be relatively minor compared to the electrostatic effects of the phosphates, the enzymic subsites must clearly be able to preferentially distinguish and discriminate the various bases. For example, Mikulski et al. have shown that the nature of the base in the /3 position exerts a dominant role in the susceptibility of hydrolysis of dinucleotides; a clear preference is demonstrated for A and T ( 2 2 ) . Also consistent with this are the observations that 5‘-mononucleotide binding shows a clear preference for A and T (3,6 S ) , that poly A is more rapidly hydrolyzed than poly C or poly U (S), and that T p are preferentially released during the early phases of RNA and DNA hydrolysis. For simplification, the scheme shown in Fig. 5 does not take into account the essential role of Ca2+in binding and hydrolysis of substrates and inhibitors. Complex formation with possibly several side chain car-

194

C. B. ANFINSEN, P . CUATRECASAS, AND H. TANIUCHI

35%

4

-20

r--

0

-

Nuclease 30 A x 3 0 i x 4 0 i

T

B

*

A

T

T

B

(C)

FIG.6. Schematic: representation of (a) complex formation brtwrrn stalh,~.loc~occal nucleasc and polynucleotide substrates ; ( b ) inhibitory oligonuclrotid(ss h i r i n g 5’-phosphoryl termini ; and (c) a slowly hydrolyzrd 3 ’ - ~ ~ h o s ~ 1 l i o r ~ l o l i g o n ~ i ~ ~ l ~ ~ o t i d r . XpYpZp. The major substrate binding rrgions of the enzyme, or “phosphatr subsites,” are indicated as P1, P2,and P3. The hydrolytic site (H) consists of a rcgion closely related to P1 which recognizes the phospliodirster bond ( A ) :inti a rrgion which recognizes the sugar-base moiety whose 5’-OH is linked to the phosphatc group (B).

8.

STAPHYLOCOCCAL NUCLEASE

195

boxyl groups, and possibly phosphate groups, are strongly suggested by the three-dimensional structure (see section below on synthetic analogs).

VI. Stereochemical Probes of the Active Site

Some information concerning the stereochemical orientation of active site residues in the solution state has been obtained by affinity labeling studies using broinoacetarnidophenyl ( 6 9 ) and diazoniuin (70) derivainhibitors. tives of p-aniiiio~~liciiylnuclcotide The tyrosyl residue a t position 115, which can be specifically modified with tetranitromethane in the presence of pdTp and Ca'+ ( 7 l ) ,is labeled selectively with affinity labeling reagents derived from pdTp-aminophenyl (6'9). The bromoacetyl reagent derived from this inhibitor also reacts specifically with lysines 48 and 49 and with histidine 46. These results suggest that there may be some flexibility in the 46-49 region of the protein, and that this region as well as that of tyrosine 115 must be near the active site. The selective cleavage of the 4%49 region by trypsin in the presence of pdTp and Ca2+ (48) is also consistent with these observations. X-Ray crystallographic analysis of this protein indicates that the three-dimensional structure of the active site permits precise attachment of the alkylating and diazonium reagents to the hydroxyl and ortho carbons, respectively, of tyrosine 115 ( 4 9 ) .Furthermore, a peptide loop containing residues 46 through 49 lies near the binding site. The catalytically essential nature of tyrosine 85 and its proximity to the substrate binding site and to tyrosine 115 were demonstrated from studies of modification with tetranitromethane (7f) and from studies of intramolecular cross-linking of aminotyrosyl residues ( 7 2 ) . The bromoacetamidophenyl (69) and diazonium (70) reagents obtained from aminophenyl-pdT both react selectively and exclusively with tyrosine 85. This residue is situated, stereochemically, such that its hydroxyl group can interact with the 3'-phosphate of pdTp. The regions of residues 46-49 and of tyrosine 115 must be located in the section of the binding site which intcracts with that part of the substrates or inhibitors extending to the 3'-carbon side of the basic structural element, R-dpT. This region is probably of importance in the binding function of the enzyme. More exact correlation of affinity labeling 69. 70. 71. 72.

P. Cuatrerasas, P. Cuatrecasas, P. Cuatrecasas, P. Cuatrecasas,

M . Wilchek, and C. B. Anfinsen, JBC 244, 4316 (1969) JBC 245, 574 (1970). S.Fuchs, and C. B. Anfinsen, JBC 243, 4787 (1968). S. Fuchs, and C. B. Anfinsen, JBC 244, 406 (1969).

196

C. B. ANF INS E N, P. CUATRECASAS, AND H. TA N I U C H I

studies and catalytic mechanism with structure must await the further refinement of the crystallographic data that is now in progress.

VII. Complementation of Fragments

One approach to the understanding of the relationship between the amino acid sequence of a protein and its three-dimensional structure consists of preparing fragments which reconstitute a functional nativelike structure by noncovalent association. Richards first demonstrated that the two fragments of hovine pancreatic ribonuclease, RNase-S-peptide (residues 1-20) and RNase-S-protein (residues 21-124), the latter with four intact disulfide bonds, bind noncovalently t o form the original functional structure, RNase-S’ (73, 7 4 ) . The elucidation of the three-dimensional structure of RNase-S by X-ray crystallographic study confirmed these observations (‘75).The RNase-S-protein-RNase-S-peptide system also provided a way by which chemically synthesized fragments could be used to test the role of individual residues in the formation of the functional structure of the protein (76-79). Nuclease is a second case in which fragments could be obtained which yield productive complementation (47, 48, 80, 81) (Fig. 7). Observations on such complementations have contributed to the understanding of protein folding and, as with ribonuclease, have provided convenient systems for the synthetic approach t o the study of the structure-function relationship in nuclease. Binding of thymidine-3’,5’-diphosphate (pdTp) and Ca’+ to nuclease induces resistance t o proteolysis by a number of proteases (47, 5 3 ) . Trypsin rapidly cleaves the peptide bond between residues 5 and 6 of the liganded nuclease. The fragment thus formed, nuclease- (6-149), possesses enzymic activity and structure similar t o nuclease ( 4 8 ) . Further cleavage of nuclease-(&149) in the presence of the ligands takes place 73. F. M. Richards. Proc. Natl. Acod. Sci. U.S. 44, 162 (1958). 74. F. M. Richards and P. J . Vithayathil, JBC 234, 1459 (1959). 75. H. W. Wyckoff, K. D. Hardman, N. M. Allewell. T. Inagarni, L. N. Johnson, and F. M. Richards, JBC 242, 3984 (1967). 76. F. M. Finn and K. Hofmann, JACS 87, 645 (1965). 77. B. Gutte and It. B. Merrifield, JACS 91, 501 (1969). 78. R. Hirchmann, R. F. Nutt, D. F. Vcber, R. A. Vitali, S. L. Valgn, T. A . Jacob, F. W. Holly, and R. A . Denkewalter, JACS 91, 508 (1969). 79. I. Kato and C. B. Anfinsen, JBC 244, 5849 (1969). 80. H. Taniuchi and C. B. Anfinsen, JBC 244, 3864 (1969). 81. H. Taniuchi, Federation Proc. 29, No. 2, 335 (1970).

8. STAPHYLOCOCCAL NUCLEASE I49

I

Nuclease

I49

6

Nuclease- (6-149) Nuclease-T-(6-48)

197

48

I -?

Nuclease-T-(49- 149)

49

149

Nuclease-T-(50-149)

50

149

Nuclease-(l-126)

!

126

Nuclease-(99-149)

99

149

,

149

ILI

Nwlease-(Ill-I49)

1 2 7 4 9

Nuclease-(127-149)

FIG.7. Fragments of nuclease. The residues of the fragments are indicated by the numbers in the parentheses. Unfilled lines represent inactive and disordered fragments.

a t the bonds between residues 48 and 49 and between residues 49 and 50 a t approximately equal rate. The two derivatives formed each consist of two noncovalently bonded fragments: [nuclease-T- (6-48) ; nucleaseT- (49-149) ] and [ nuc1ease-T- (6-49) ; nuc1ease-T- (50-149) 1. Trypsin removes lysine 49 from the latter. The final product, “nuclease-T,” is a mixture of the two complexes [nuclease-T- (6-48) nuc1ease-T- (49149) ] and [nuclease-T- ( 6 4 8 ) nuc1ease-T- (50-149) ] (48).Nuclease-T has an enzymic activity approximately 8% that of nuclease and appears to have ordered structure similar to nuclease (47, 48, 5 3 ) . Both nuc1ease-T- (4S149) and nuc1ease-T- (50-149) are enzymically inactive and disordered but bind to the inactive fragment nuc1ease-T- (6-48) to form nuclease-T‘ which is functionally and structurally like nuclease-T (48, 53). Thus nuclease-T (or T’) represents complementation involving a discontinuity of the polypeptide chain in the region of residues 48-50. Nuclease-T (or T’) also exhibits the ligand-induced resistance to proteolysis observed with native nuclease ( 5 3 ) . Another large fragment of 126 residues, including the NH,-terminus nuclease-(1-126) can be obtained (80) from nuclease that has been trifluoroacetylated (82). Trypsin cleaves the bond involving the carboxyl group of arginine 126 in this trifluoroacetylated derivative more rapidly than those of the other arginine residues. Limited incubation of trifluoroacetylated nuclease with trypsin produces the fragment containing

+

82.

R. F. Goldberger and C . B. Anfinsen, Biochemistry

+

1, 401 (1962).

198

C. B. ANFINSEN, P. CUATRECASAS, AND H . TANIUCHI

residues 1-126, together with the residual sequence, residues 127-149, in fairly good yields. Nuclease- (1-126) and nuclease- (127-149) were isolated from the incubation mixture after removal of the trifluoroacetyl groups. Nuclease-(1-126) is inactive and appears to be disordered as judged by measurements of optical rotation, circular dichroism, and intrinsic viscosity (80). Nuclease- (127-149) is also inactive and does not bind to nuclease-(l-126) (80). Thus, the cleavage of the peptide bond between residues 126 and 127 destroys the functional structure of nuclease in contrast to the cleavage of bonds in the region of residues 48-50, which leaves the structure of nuclease essentially unchanged. However, nuclease- (1-126) complements with a BrCN fragment nuclease-(99-149) (see above) to form an enzymically active (at the level of nuclease-T) and ordered complex (81). The structure of this complex is less resistant to trypsin digestion in the presence of pdTp and Caz+than that of nuclease-T although such resistance is still observed (47, 81). Trypsin very rapidly removes residues 99-110 from the complex in the presence of the ligands. The derived complex, composed of nuclease- (1-126) and nuclease- (111-149), possesses an enzymic activity similar to that of the original complex. Isolated nuclease- (1-126) and nuclease- (111-149) combine to regenerate the enzymic activity (81). Accordingly, residues 99-110 of nuclease- (99-149) appear not to participate in the ordered structure formed by nuclease- (1-126) and nuclease- (99-149) and probably extends flexibly from the ordered structure. Further removal of residues 111-113 and 125-126 with exoproteases also appears not to affect the complementation. When nuclease- (1-126) is mixed with nuc1ease-T- (49,50-149) [the mixture of nuc1ease-T- (49-149), and nuclease-T- (50-149) 1 , enzymic activity is generated a t the level of nuclease-T and the formation of ordered structure is also indicated (80). After incubation of the enzymically active mixture in the presence of pdTp and Ca'+ with trypsin for a fairly long time, approximately 50% of the original enzymic activity remains. The enzymically active complex thus formed is the same as nuclcase-T' and is composed of the two fragments nuclease-T-(6-48) and nucleaseT- (49-149) [or nuc1ease-T- (50-149) 3 (80). Therefore, nuc1ease-T- (49, 50-149) binds to the nuc1ease-T- (6-48) portion of nuclease- (1-126) forming an enzymically active structure like nuclease-T', and the redundant residues 49-126 of nuclease- (1-126) extend in a disordered manner from the ordered structure. When the mixture of nuclease- (1-126) and nuclease-T- (49, 50-149) are incubated in the presence of the ligarids with trypsin for a short time, the fragments composing the undigested cornplexes are nuclease-T- (49,

8.

199

STAPHYLOCOCCAL NUCLEASE

n

II FIG.8. Sdwmatic illustration of the simultnnvous formation of two types of complrmentations from nuclease-(l-l26) nnd nuclease-T-(49-149). The polypeptide chains hypotlirtically participating in the ordercd structtires are circlrd.

50-149), nuclease- (1-48), nuclease- (1-126), and nuclcase- ( 1 11-149) (81). Therefore the complex composed of the nuclease- (1-126) and nuclease(111-149) as well as nucleasc-T were formed in the mixture (Fig. 8 ) . The latter type of complex appears to predominate (81). These observations on the co~nplementationof fragments of nuclcasc are consistent with the concclpt that almost the entire arnino acid sequence of nucleasc is required for the minimum information necessary to determine a stable, functional nuclcasc structure. Furthermore, the native conformation of nuclease cannot be forrned during assembly from the NH,-terminus until the polypeptide chain has been extended beyond residue 126 (80). Finally, i t appears that stable and functioiial structures can he formed in several ways when the minimum requirement for information is fulfilled (81).

VIII. Synthetic Analogs

T h e solid phase method (83)for the synthesis of peptides and proteins has been used in the synthesis of a large number of analogs of nuclease. 83. R . B. Mcrrifirld. JACS 85, 2148 (1863) ; Scieme 150, 178 (1965).

200

C. B. ANFINSEN, P. CUATRECASAS, AND H. TANIUCHI

The solid phase method possess certain inherent sources of inaccuracy resulting for the most part from the slight incompleteness of coupling that may occur a t each step of amino acid addition to the growing chain and from certain side reactions during the final deprotection steps. However, careful purification of synthetic products has been carried out (84, 85) , and synthetic variations in sequence have been restricted to those that yield essentially all-or-none answers about questions of enzymic function and structure. The synthetic studies have been made possible by the availability of complementing fragments of nuclease that contain approximately 50 amino acid residues or less (48, 81, 86). These fragments, nuclease-T- (& 48) or nuclease-T-(&47) and nuclease-(99-149) (48, 81, 86), have been discussed above in connection with the limited proteolysis of nuclease and the study of principles involved in the folding of the chain. Following synthesis and purification, the fragments can be assayed for their capacity to generate enzymic activity when combined, noncovalently in solution, with the proper complementing peptide. The ability of the various analog peptides to yield nativelike conformations upon such combination can also be estimated by spectroscopic, fluorimetric, and other methods. For example, the measurement of the intensification of tryptophan fluorescence, and the blue shift in the wavelength of maximum emission, have been of particular convenience in such studies (80, 85, 87) (see Fig. 9 ) . Several classes of analogs of the two sequence regions 6-47 and 99-149 have now been prepared, including analogs varying in the extent of deletion at the amino terminus of the protein (87, 8 8 ) , those concerned with changes in the nature of the amino acids associated (from a study of the three-dimensional crystallographic data) with the active site (87-89) and analogs that were designed t o give information about the side chain requirements for binding of the carboxyl-terminal helical region to the rest of the molecule to yield an cnzymically active recombinant (90). I n the case of the fragments containing residues 6 4 7 , which can combine with nuclease-T- (49, 50-149) to yield an active complex (see above), omission of residues 6 , 7, and 8 yields a derivative which still binds to the larger fragment and shows undiminished activity (87). Deletion of residue 9, however, reduces activity to about 2070 of the normal level, and further deletion of residue 10 produces an entirely inactive semisyn84. D. A. Ontjrs and C . B. Anfinsen, Proc. Nntl. Acnd. Sci. U . S. 64, 428 (1969). 85. I. M. Chaikrn and C. B. Anfinsen. JBC 245, 2337 (1970). 86. H . Taniuchi and C . B. Anfinsen. JBC (in prrss) (1971). 87. D. A. Ontjcs and C. B. Anfinscn. JBC 244, 6316 (1969). 88. I. M . Cliaiken and C. B. Anfinscn, JBC 245, 4718 (1970). 89. I. M. Chaiken a n d C . n. Anfinsrn, JBC (in press) (1971). 90. I. Parikh and C. B. Anfinsen, manuscript in preparation.

8.

201

STAPHYLOCOCCAL NUCLEASE

60

50

-2 -

A

2,

._

40

0

5 0

c

30

e 3 -

LL

20

10

Emission wavelength, nm

FIG.9. Fluorescence emission spectra of mixtures of nuclease-T-(49,50-149) with nuclease-T-(6-48) or synthetic pcptides upon excitation a t 295 nrn. Samples containing 2 X 10.' M nuclease-T-(49,50-149) and 4 X 10' M nuclease-T-(648) or synthetic peptide were in 0.05M tris, pH 8, with 0.01 M CaCL and O.OOO1 M pdTp. The spectrum for nuclease-T-(49,50-149) alone was corrected for the small measured emission exhibited by the buffer solution containing pdTp and Ca*+. The spectra for mixtures of synthetic peptide or nuclease-T-(6-48) with nuclease-T(49,50-149) were corrected for the meoswed emission exhibited by the appropriate synthetic peptide [or nuclease-T-(6-48) 1 alone in buffer containing pdTp and Ca'+. ( 0 )Nuclcase-T-(&48), (0) synthetic ( 6 4 7 ) , (A)synthetic (%47), ( A ) synthetic (18-47), (0) synthetic ( 3347) , and (m) nuclease-T-(49, 50-149) alone. Taken from Ontjes and Anfinsen (87).

thetic complex (88).These effects may be rationalized on the basis of the three-dimensional structure in which lysine (9) and glutamic acid 10 appear to contribute to the stabilization of a portion of the structure containing a number of hydrophobic amino acid side chains (19). Even shorter fragments retain some capacity for combination with nuc1ease-T(49, 50-149), as evidenced by their ability both to enhance the binding of 4251-[nuclease-T-(49,50-149) ] to antinuclease (87, 91) and to bind to Sepharose t o which nuclease-T- (49-149) is covalently bound (87). 91. G . S. Ornenn, D. A. Ontjcs, and C. B. Anfinsen, Biochemistry 9, 313 (1970).

202

C. B. ANFINSEN, P. CUATRECASAS, AND H. TANIUCHI

A number of residues may be replaced with other amino acids without apparent change in enzymic activity. These replacements include norleucine for methionine a t positions 26 and 32 (87), phenylalanine for tyrosine 27 ( 9 2 ) , and glycine for histitline 46 (85).As mentioned above, deletion of histidine 8 is without deleterious effect. The two remaining residues of histidine at positions 121 and 124 may also be ruled out as components of the active site since the former is, stereochemically, far on the other side of the molecule and the latter is replaced by leucine in nuclease from the Foggi strain of Staphylococcus aureus. Changes in residues that are implicated in the active center of the

I GLU

A S N 8, A

f

FIG.10. Correlation of active site synthetic analog results for nuclease-T (takrn from Table 111) with the X-ray crystallographic model for the binding site region for nuclease [taken from Cotton and Hazen (IS)]. B indirates binding of a synthetic ( 6 4 7 ) analog, with noted change, to nucleuse-T-(49,50-149) ; NB, ineffectivc binding; A , the ability of the synthetic ( 6 4 7 ) analog to generate a t least partial enzymic activity upon addition to nuclease-T-(49, 50-149) ; I, little or no ability t o generate such activity. 92. C. B. Anfinsen, D. A . Ontjes, and I. M. Chaiken, Proc. 10th Europecin Peptide Symp., 1969 (in press).

8.

203

STAPHYLOCOCCAL NUCLEASE

enzyme lead to results that are essentially as one would predict from the structure (19). Thus, as shown schcrnatically in Fig. 10 and summarized in Table 111, glutamic acid 43, aspartic acid residues 21 and 40, and arginine residue 35 play important and, indeed, except for aspartic acid 40, essential roles in the binding of the calcium atom and the inhibitor molecule pdTp. The presence of a charged carboxylate group on aspartic acid 40 is apparently not obligatory for activity (asparagine may be introduced for aspartic acid) although replacement with the bulkier glutamic acid leads to an inactive, semisynthetic nuc1ease-T complex. Both the capacity to bind to, and to induce activity with, nuc1ease-T(49, 50-149) are lost when arginine 35 is replaced with the neutral residue citrulline. The same occurs upon replacement of aspartic acid 21 with

PROPERTIES

Residue position

OF

TABLE I11 COMPLIGMENTATION OF SYNTHETIC ( 6 4 7 ) ACTIVESITE ANALOG PEPTIDIB WITH NUCLEASE-T-(4Q,50-149)'

Binding to Normal Replacement nuclease T- (49, 50-149)

21

ASP

35

-k%

40

ASP

43

Glu

(I

Glu Asn LYS Cit Glu Asn ASP Gln

+++ + + +

Enzymic activity upon addition to nuclease-T-(49, 50-149) 0 0

0 0 0

Partial 0 0

From Chaiken and Anfinsen (89).

asparagine. Both the stereochemistry and charge of the side chain on glutamic acid 43 appear to be critical for activity since replacement with either glutamine or aspartic acid causes complete inactivation, although both analogs form complexes with nuclease-T- (49, 5C149) as indicated by fluorescence measurements. As discussed earlier in relation to the complementation of fragments of nuclease to yield enzymically active, structured complexes, a fragment consisting of the first 126 residues is able to interact through noncovalent bonds with a fragment containing residues 99-149 which results from BrCN cleavage of the native protein (81, 86). This latter polypeptide, itself structureless in solution as is the 126 residue fragment, has been synthesized in several analog forms (go), and these derivatives have been tested for their ability to yield active complexes. The fragment

204

C. B. ANFINSEN, P. CUATRECASAS, AND H. TANIUCHI

may be shortened a t both ends (deleting residues 99-114 and 142-149), and the tryptophan residue a t position 140 may be replaced with phenylalanine, apparently without significant loss of activity and structureproducing capacity.

Microbial Ribonucleases with Special ReJerence to RNases TI. T2.JV;. and U2 TSUNEKO UCHIDA

F U J I 0 EGAMI

I . Introduction . . . . . . . . . . . . . I1. Fungal RNases TI. T.. N.. U.. and Ug . . . . . . . . A . General Survey . . . . . . . . . . . B . RNase TI . . . . . . . . . . . . C . RNase T. . . . . . . . . . . . . D . RNase N. . . . . . . . . . . . . E . RNase U. . . . . . . . . . . . . I11. Other Microbial RNases of Special Interest . . . . . . A . Extracellular RNases of B . subtilis Strain H . . . . . B . Intracellular RNase of B . subtilis . . . . . . . C . RNase PP, of Physarumpolycephalum . . . . . D. RNases 11. 111, IV. and V of E . coli . . . . . . IV . List of Microbial RNases . . . . . . . . . .

205

208 208 212 223 230 234 239 239 240 241 241 243

.

1 Introduction

Ribonucleases (RNases) may be defined as phosphodiesterases that attack the internucleotide bonds in ribonucleic acid (RNA) and its products but not those in deoxyribonucleic acid (DNA) or simple phosphodiesters such as bis-p-nitrophenyl phosphate . Ribonucleases may be classified conventionally by their mode of 205

206

T. UCHIDA A N D F. EGAMI

TABLE I CLASSIFICATION O F RIBONUCLEASES A. Classification by the mode of action 1. Degradation by nucleotidyl transfer 2. Degradation by hydrolysis 1. Endonucleolytic 2. Exonucleolytic (a) From 3‘ terminal (b) From 5’ terminal B. Classification by the reaction products 1. Enzymes producing 5’-nucleotides 2. Enzymes producing nucleoside 2’,3’-cyclic phosphates (a) Enzymes producing nucleoside 2’,3’-cyclic phosphates as final products (b) Enzymes producing 3’-nucleotides as final products 3. Enzymes directly producing 3‘-nucleotides C. Classification by the substrate specificity 1. Classification by the base specificity (a) Nonbase specific (b) Pyrimidine specific (c) Guanine specific (d) Purine specific 2. Classification by the conformation specificity (a) Preferential to single-stranded RNA (b) Practically inactive to double-stranded RNA (c) Specific for double-stranded RNA

action, by their reaction products, and by their substrate specificity as shown in Table I. Most typical RNases such as pancreatic RNase (RNase A ) , described in the chapter by Richards and Wyckoff, and RNases TI, T,, N,, and U,, which will be described in this chapter, are characterized as follows: Degradation by nucleotidyl transfer Endonucleolytic Producing nucleoside 2’,3‘-cyclic phosphates which are more or less slowly hydrolyzed to produce 3’-nucleotides Practically inactive to double-stranded RNA Fairly heat stable

As for the base specificity, it is generally recognized that RNase T, is nonbase-specific, but RNases A, T, , and U, are base-specific for pyrimidine, guanine, and purine, respectively. However, generally speaking, the base specificity is not absolute but relative. For example, RNase A splits poly A a t a high concentration of the enzyme.

9.

207

MICROBIAL RIBONUCLEASES

This chapter deals with microbial RNases ( 1 ) of interest because of their enzymology or their use in nucleic acid research, with special reference to fungal RNases. Details of experimental methods are not included. Original papers or appropriate books (2, 3) should be consulted for methods of enzyme preparation and for assays of enzymic activity. For earlier surveys the chapters by G. Schmidt and M. Laskowski, by H. G. Khorana, and by C. F. Anfinsen and T. H. White, *Jr., in the second edition of “The Enzymes,” Vol. 5 , and specialized monographs (1, 4 ) should be consulted. I

I

O=P-O-

I

o=p-o-

0

0

I

0

I

o=p-o-

I

I

I

?

o=p-oI

OH

FIG. 1. Reaction catalyzed by typical RNases. 1. F. Egami and K. Nakamura, “Microbial Ribonucleases.” Springer, Berlin, 1969. 2. G . L. Cantoni and D. R. Davies, cds., “Procedures in Nucleic Acid Research.”

Harper, New York, 1966. 3. 6 . P. Colowick and S . 0. Knplan. ds.. “Methotis in Enzymology.” Vol. 12. Part A , 1967. 4. M. Privat de Garilhe, “Enzymes in Nuclric Acid Research.” Hermmn. Paris. 1967.

208

T. UCHIDA AND F. EGAMI

II. Fungal RNases, T,, T,, N,, U,, and U,

A. GENERAL SURVEY All fungal RNases (TI, T,, N,, U,, and U,) treated in this section catalyze the reaction shown in Fig. 1. The first step (phosphate transfer) is the cleavage of the phosphodiester bond between the 3' and 5' positions of the ribose moities in the RNA chain with the formation of nucleoside 2',3'-cyclic phosphates and oligonucleotides with 2',3'-cyclic phosphate a t 3' terminal. The nature of the phosphodiester bonds to be cleaved depends on the base specificity of the enzyme. This phosphoryl transfer step is reversible. In the second step (hydrolysis), these terminal cyclic phosphate groups are hydrolyzed with the formation of corresponding %-phosphates. Because the first-step is usually faster than the second step, more or less accumulation of the cyclic phosphate may be observed. TABLE I1 MAIN PROPERTIES OF FUNGAL RNasesovb RNases

Source pH optimum for RNA digestion Base specificity Specific activity (units/mg) for RNA digestion Molecular weight (amino acid analysis) Absorption max (nm) Absorption min (nm) 0Dmx/ODm1.

Ul (Ref. 7)

uz

TI

(Ref. 7)

(Ref. 6)

Ti (Ref. 6)

NI (Ref. 6 )

Aspergillus oryzae

Neurospora crassa

7.5

7.0

8.0

4.5

4.5

Guanine

Guanine

Guanine

Purine

1.6 X lo"

4.0 X lo4

1.7 X 10'

-

Nonbasespecific 1.4 x 103

11,085

11,000

11,000

10,000

36,200

278

279

278.5

277.5

281

251.5

252

251.5

251

252

3.68

3.15

Ustilago Ustilago sphaerogena sphaerogena

4.16

2.6

Aspergillus oryzae

2.52

One unit; the amounts of enzyme causing an increase in absorbance a t 260 nm of 1.0 by the assay described by Uchida and Egami (3, see p. 228) is defined as one unit of enzyme. * Some unpublished data are included.

9.

209

MICROBIAL RIBONUCLEASES

Main properties of these enzymes are summarized in Table I1 (5-7). The amino acid compositions of these enzymes are shown in Table I11 in comparison with each other and with RNase A. Base-specific RNases TI, N,, U,, and U, are of similar molecular size, unlike nonbase-specific RNase T,, which is much larger. The amino acid composition of RNases T,, N,, and U,, which are classified as guanyloribonucleases, are similar to each other, as shown in Table 111. It is remarkable that all three RNases have four residues of half-cystine. From this finding, it may be expected that these three enTABLE I11 AMINOACID COMPOSITION OF RNases Amino acid

Ti (Ref. 6)

Tz Nia

UP

A

(Ref. 6)

3 1 15 6 15 9 4 12 7 4 8 0 2 3 9 4 1

3 3 3 14 4 14 4 5 13 10 4 4 2 5 4 9 5 1

3 2 2 16 8 13 6 4 15 5 4 6 0 2 1 12

0

10 4 4 15 10 15 12 4 3 12 8 9 4 3 2 6 3 0

23 6 4 39 25 32 39 23 28 19 11 7 1 19 18 14 10 7

Total

104

107

103

124

325

N-terminal

Ala

Ala

-

Lys

C-terminal

Thr

-

TYr

Val

Glu (or Gln) -

LYS His -4% ASP Thr Ser Glu Pro GlY Ala +cys Val Met Ile Leu TYr Phe TrP

a

1

4

Unpublished data.

5. F. Egami, K. Takahashi, and T. Uchida, Prog. Nucleic Acid Res. Mole. Biol. 3, 59 (1964). 6. N. Takai, T. Uchida, and F. Egami, BBA 128, 218 (1966). 7. T. Arima, T. Uchida, and F. Egami, B J 106, 601 (1968).

210

T. UCHIDA AND F. EGAMI

zymes may have a similar architecture maintained by two disulfide bonds. Two or three residues of histidine, which has been regarded to be essential for active site in RNase A and also RNase T,, are found in each RNase. Two methionine residues are found only in RNase N,, and no tryptophan residue is found in RNase U,. Thus, neither absence of methionine nor presence of tryptophan is considered to be characteristic in guanyloribonucleases. At least the presence of tryptophan is not essential for the activity of guanyloribonuclease. The contents of aromatic amino acid residues are also similar to each other, as indicated by the similarities of the UV absorption spectrum shown in Tahle 11. However, the UV absorption spectrum of RNase U, is different from the others in that it has no shoulder a t 285 nm. This may reflect the difference in contents of tyrosine and tryptophan. The content of acidic amino acid residues increases in an order of N, < U, < T,, whereas that of basic amino acid residues decreases in the same order. This shows a good agreement with the order of the acidities of these enzyme proteins as observed in their behaviors on DEAE-cellulose column chromatography. The effects of inhibitors and activators on RNases T,, N,, U,, U,, and T, are summarized in Table IV (5, 7, 8 ) , showing the characteristics of each RNase. Ribonuclease T, is strongly inhibited by 10-3M Zn2+; RNases U, and T, are inhibited about half; but RNases N, and U2 are not inhibited. Ribonuclease T, is also strongly inhibited by 10-3M Ag+, while all other RNases are inhibited about half. But M Cu2+strongly inhibits RNase T,and only about half inhibits the other RNases. All five RNases are not inhibited by IO-, M ethylenediaminetetraacetate (EDTA) M ICH,COOH. No metallic cofactor is required for their action. and An immunochemical study on RNase T, with its immunological relation to other RNases has been carried out by Uchida (9). Antisera to RNase T, can be prepared in rabbits. The content of antibody in the antisera thus obtained is about one-tenth that in the antisera to RNase A prepared by the same method, suggesting that the immunogenicity of RNase T, may be very weak. Enzymic activity of RNase T, assayed with RNA is completely inhibited by binding with the antibody in the region of antibody excess, However, about 20% of the enzymic activity of the RNase T, used is found in the washed immune precipitate when assayed with guanosine 2',3'-cyclic phosphate. Therefore, the inhibition of the enzymic activity by binding with the antibody is considered t o result not from the specific masking of the active site but from the 8. N. Takai, T. Uchida, and F. Egami, Seikagaku 39, 473 (1967). 9. T. Uchida, J. Biochem. ( T o k y o ) 68, 255 (1970).

9.

21 1

MICROBIAL RIBONUCLEASES

TABLE I V ~ ~ : ~ ' F I ~ ;OcYT sINHIIIITOBS ANI)

ACTIVATORS O N FUNGAL IlNasea

Activity remaining (7;) Final COIlC.

Ileagents NaCl NaF NaN3 Na2S MgC1,

CaC12 AgNOa HgCh MIISO~ ZnSO,

cuso, FeSOl ICH&OOH BrCHICOOH DFP His

EDTA PCMB (I

-log ill

TI

(Ref. 6 )

N, (Ilef. 8)

U, (Ilef. 7)

9 66 81 100 100

-

-

-

-

33a 96O 56 60

79" 1100 94 60

0 1 3 1 2 2 1 2

75 100-1 15

-

19 67

3 2 3 3 2 3 3 3 2 3 2 3 4 6 2 3 2 4

-

-

70-75 0 10 45

114 51 20 103 62 108 47 245 160 190 115

Y3-100 95-100 10 60

-

0 20-50 20

-

100

-

-

100 100 82 150 120-150

10b

-

-

110

83

-

-

lJ2 (Ref. 7) 71

-

-

24b 104b 41" 61

72b 15gb 102c 53

109 117 108

112 55 103

-

-

115 101

91 95

-

-

-

T p (Ref. 5 ) 40-50 90

104 92 36 78

-

90 50 40 66

50 0 58

-

120

-

100 100 100 100 110

-

MgSO, is used instead of MgCL MnClz is used instead of MnSO1. ZnAc is used instead of ZilSO,.

steric hindrance of accessibility of substrate molecule to the enzymic site by binding with the large antibody molecules. Both RNases T, and U,, with quite different base specificities from that of RNase T,, are immunologically unrelated to RNase T,. Among the guanyloribonucleases, RNase N, has no immunological relation to RNase T, but RNase U, is found to be immunologically somewhat correlated to RNase T,. It remains to be determined if this observation is related to the difference of their enzymic activity: RNase N, is distinguishable from RNases T, and U, by its lower hydrolase activity.

T. UCHIDA AND F. EGAMl

B. RNase T, In 1957, RNasc T, ( 5 , 10, 11) was isolated from Takadiastase, a commercial product of Aspergillus oryzae ( 1 2 ) .It is now a familiar enzyme used as an essential tool for the structural analysis of RNA. 1. Preparation

Since the partial purification of RNase T, was carried out by Sat0 and Egami ( l a ) , several purification procedures have been reported. Takahashi (13) first obtained the purified RlSase T, as a homogeneous protein in the form of fine spherular crystals by the following procedure: heat treatment in acidic pH of crude extract, fractionation by ammonium sulfate (0.61.0 saturation), absorption of impurities by acid clay, two separations on DEAE-cellulose, reabsorption of impurities by acid clay, and crystallization by dialysis against the solution of 0.6 ammonium sulfate saturation. Soon after, Rushizky and Sober (14) also succeeded in the purification of this enzyme by a different procedure: After the simplified extraction in pH 2.6 and then concentration by 66% acetone, they mainly used four systems of DEAE-cellulose column chromatography. Judging from the ratio of the absorbance a t 280nm to that a t 260nm, the resulting preparation of RNase T, appears to be somewhat less pure than that prepared by Takahashi's procedure. Uchida (16) simplified a purification procedure with increase in both yield and purity. First an RNase T, fraction was obtained from the crude extract of Takadiastase Y powder by a batchwise treatment with DEAEcellulose, and then other enzymic activities in the RNasc T, fraction were inactivated by heating in pH 1.5 at €302 2" for 2 min. Ribonuclease T, in the heat-treated fraction was absorbed on the acid clay with other proteins a t acidic pH and specifically eluted at neutrality. This step serves effectively for the isolation and concentration of the enzyme preparation. After the eluate was fractionated by ammonium sulfate (70-100% saturation), the dialyzed RNase T, was subjected to DEAEcellulose column chromatography, in which RNase T, of specific ac10. F. Egami, J. Sn'. Ind. 12e.s. (India) 25, 1 (1966). 11. K. Takahashi, F. Egami. and T. Uchida, Advan. Biophys. (Japan) 1, 53 (1970). 12. K. Sat0 and F. Egami, J . Biochem. ( T o k y o ) 44, 753 (1957). 13. K. Takahashi, J. Biochem. (Tokuo) 51, 95 (1962). 14. G. W. Rushizky and H. A. Sober, JBC 237, 834 (1962). 15. T. Uchida, J. Biochem. ( T o k y o ) 57, 547 (1965).

9.

MICROBIAL RIBONUCLEASES

213

tivity approximately 500-800 could be prepared in a yield of approximately 29487%.Whenever further purification is desired, the crystallization as described in Takahashi's method for the removal of color contaminants, or gel filtration by Sephadex G-75 for the exhaustive removal of RNase T,, may be added to the above procedure. Minato e t al. of Sankyo Co. Ltd. (Tokyo, Japan) (16, i7) developed independently a large-scale purification procedure for the commercial preparation of RNase T1. Recently, R. Fields, H. B. F. Dixon, and C. Yui (unpublished) have developed a chromatographic system for the isolation of RNase TI. This procedure uses only a column of DEAE-cellulose equilibrated with a buffer of 0.0844 NaH2P0, and 0.1244 Na,HPO, after the water extraction of Takadiastase Y powder. 2. Properties Most of the properties of RNase TIare summarized in Tables I1 and IV. It is a very acidic protein, active between pH 4 and 8.5; it is most active at pH 7.5 for RNA digestion (12) and a t pH 7.2 for the hydrolysis of guanosine 2',3'-cyclic phosphate (18). The purified enzyme possesses a specific activity of about 1.6 x lo4 units/mg of protein. The molecular activity (standard units/pmole enzyme) has not been determined for the cleavage of a definite dinucleoside monophosphate such as GpC or for the hydrolysis of guanosine 2',3'-cyclic phosphate. Ribonuclease T1 is a stable enzyme. It is fairly resistant to heat (100' for 10 min a t pH 6) and acid but somewhat unstable in alkaline solution ( > 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

T. UCHIDA AND F. EGAMI

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

MICROBIAL RIBONUCLEASES

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

T. UCHIDA AND F. EGAMI

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 <;In L e u 111s (;lu A l p G l y GluwThr

I

Asp

I

Asn

I Ser

lyr I

45 A * p Phv l i l y (;lu T y r A l n A m Tyr I.y\

50 Sl‘r Scr Val Scr I’hc I

b I Thr ( ‘ y , C l y II HO l h r C y i

411 11th

Pro

35

VJI

I

r y r Srr A w Srr (;ly

P w S5 I

l y r Tyr (;lu

I

(;I” I 100 VJI Phc h m A m (;ly

70 75 l y r Srr l i l y Pro <;ly Srr Gly A l a A t p I A rg 90 RS 80 I h r llir Thr l k Val (;ly A h Leu (;In A w A * n G l u A m Phe Val V a l

1311 hS T q l 1’ro I I U Lru Sl‘r Srr (;ly A l p V r l

95 Sl‘r Ala G l y

FIG.2. Thc :imino iicid srqucncc of RNasc TI.

air-oxidation regenerates 110th the activity and the native structure (52, 53). Analyzing the data of UV absorption, circular dichroism (CD), and optical rotatory dispersion (ORD), Yamamoto and Tanaka suggested that RNase TIhad ahout 33% of a-helix structure, 24% of p structure, and 43% of random coil (54).Spectrochemical analysis suggests that most of the tyrosine residues and the only tryptophan residue in the enzyme are embedded in the hydrophobic region in the interior of the three-dimensional structure. This is consistent with the results on the potentiometric titration of the enzyme (19). From the data, pK values of dissociable groups were calculated and tentatively assigned to the dissociable groups of the molecule as shown in Table VIII. Most of the carboxyl groups TABLE VIII pK

VALUES OF

IhssocImLE GROUPSI N RNase Tp

Intrinsic PKb values

No. of groups/ molecule

Tentative assignment to dissociable groups

3.85 5.0 6.6 7.2 7.4 8.6 9.7

10 3 2 1 1 1 2

Carboxyl Carboxyl Imidazole Imidazole Cy-Amino rAmino Phenolic

From Iida and Ooi (19). Reprinted from Biochemistry 8 , 3897 (1969). Copyright 1969 by the American Chemical Society. Iieprinted by permission of the copyright owner. Potentiometric titration was performed i l l 0.1 M NaCl a t 25”. Seven of the phenolic groups and a guanidyl group were not titrated. 52. 8. Yamagata. K. Taknhashi, and I;. Eganii, J. Biochem. (Tokyo) 52, 272 (l(362). 53. K. Kasai. J. Biochem. (Tokyo) 57, 273 (1965). 54. Y. Yamamoto and J . Tnnaku. BBA 207, 522 (1970).

220

T. UCHIDA A N D F. EGAMI

are not in direct contact with the solvent but interact with other groups in the molecule. Riiterjans et al. (55) carried out proton magnetic resonance studies of RNase T, and, based upon the chemical shifts of the C-2 proton of the histidine residues of the enzyme, suggested that histidine residues interact with carboxylate anions of amino acid residues. By gel filtration experiments and by the difference spectrum of the mixture of RNase T, and 2’-guanylate, a strong competitive inhibitor, it was found that one molecule of RNase T, binds one molecule of the inhibitor. Thus, the enzyme molecule may be regarded to have one active center for the specific binding with the substrate ( 5 6 ) . This was confirmed by the analysis of the influence of pH and substrate analogs on the fluorescence of the enzyme (57). From the pH dependence of kinetic parameters (K,,, for guanylyl nucleosides and K , for 2’-gu:inylatesJ, Iric concluded ( 4 1 , 58) that RNase T, had at least three dissociable groups with pK, values 3.5, 5.7, and 7.5 a t its active center. The 3.5 group is very likely the carboxyl group of Glu 58, essential for the enzymic activity. At least one of others may be assigned to an imidazole group of histidine residues, which has been considered an active site of the enzyme. Riiterjans et al. also reported the presence of histidines i n the active center of the enzyme from the changes of the C-2 proton magnetic resonance caused by the addition of guanosine 2’-phosphate to RNase T1( 5 5 ) . The search for active sites of the enzyme, by chemical and enzymic modifications has been extensively carried out since 1959 and was recently reviewed in detail (11).Only a short summary will be presented here. Inactivated by carboxymethylatiori with iodoacetatc ( 5 6 ) , Glu 58 is the carboxymethylated residue and is an essential residue for the catalytic activity ( 5 9 ) . Inactivated by photooxidation in the presence of methylene blue ( G O ) , riboflavine ( G O ) , or Rose Bengal (61).Together with the inactivation the concomitant loss of histidine residues is observed. Onc (probably His 92) or more histidine residues might participate in the enzyme activity. 55. H. Ruterjana, H. Witzrl, and 0. Pongs, BBRC 37, 247 (1969). 56. S. Sato and F. Egami, Biochem. 2.342, 437 (1965). 57. 0. Pongs, B i o c h e k t r y 9, 2316 (1970). 58. M. Iric, J. Biochem. ( T o k y o ) 61, 550 (1967). 59. K. Takahashi. W. H. Stein, and S. Moor, JBC 242, 4682 (1968). 60. S. Yamagata, K. Takahashi, and F. Egarni. J . Biochem. ( T o k y o ) 52, 261 (1962). 61. K. Takahashi, J. Biochem. ( T o k y o ) 67, 833 (1970).

9.

MICROBIAL RIBONUCLEASES

221

Deamination (62) or trinitrophenylation (63) of two amino groups (a-amino group of N-terminal alanine and c-amino group of a lysine rcsidue) retains the activity. Amino groups have nothing to do with the activity. Modification with phenylglyoxal and glyoxal, arginine-modifying reagents, causes inactivation and concomitant loss of the only arginine residue. The single arginine residue might be a part of the active center (64)*

The single trytophan residue in the native RNase T, seems to be embedded in the interior of the molecule and cannot be modified by specific reagents. The modification of the residue in 8 M urea was carried out with 2-hydroxyl-5-nitrobenzyl bromide by Takahashi (64a) and with N-bromosuccinimide by Kawashima and Ando ( 6 5 ) . In both cases complete inactivation occurred when the tryptophan residue was completely modified. However, RNase TI modified with 2-hydroxyl-5-nitrobenzyl bromide in 4 M urea retained about 30% of the original activity (66). Thus, it may be concluded that the tryptophan residue does not participate directly in the catalytic process, since i t is situated quite close to the essential Glu 58, it may indirectly participate in the building up of the active conformation (11). Various experimental evidence suggests that only 2 or 3 of the 9 tyrosine residues are on the surface of the enzyme (19, 5 6 ) . Indeed only a part of the tyrosine residues can be easily modified by acetylimidazole at pH 7.5 or by tetranitromethane at pH 8.0 (H. Kasai, K. Takahashi, and T . Ando, unpublished). As enzymes thus modified have catalytic activity, the tyrosine residues that are probably located at the surface of the enzyme do not seem to be essential for activity. Consistent results were also obtained from the modification by fluorodinitrobenzene or by diaao1H-tetrazole (H. Kasai, K. Takahashi, and T. Ando, unpublished). Especially noteworthy is the derivative, in which one to two tyrosine residues, amino terminal alaninc, and one lysine residue were modified with diazo-lH-tetrazole. The derivative was deprived of most of its activity toward RNA but retained about 5Q% of its activity toward guanosine 2’,3’-cyclic phosphate. This may be explained by some steric hindrance owing to the modification of a tyrosine residue near the active center. 62. Y. Shiobara, K. Takahashi, and F. Egarni, J . Biochem. (Tokyo) 52, 267 (1962). 63. H. Kasai, K. Takahashi, and T. Ando, J . Biochem. (Tokyo) 56, 591 (1969). 64. K. Takahashi, J . Biochem. (Tokyo) 67, 541 (1970). 64a. K. Takahashi. J . Biochem. (Tokyo) 68, 659 (1970). 65. H. Kawashima and T. Ando, Intern. J . Protein Res. 1, 185 (1969). 66. T . Terao and T. Ukita, BBA 181, 347 (1969).

222

T. UCHIDA A N D F. EGAMI

Ribonuclease TI is fairly resistant to proteases. The threonine residue a t the carboxyl terminal of the enzyme can be removed by carboxypeptidase A without loss of activity (67). Leucine aminopeptidase does not release amino acids from the amino terminal (68). Ribonuclease T, is not inactivated by trypsin or chymotrypsin in the presence of 0.2 M phosphate (69), which probably binds the enzyme and protects it from inactivation (67). Treatment of the enzyme with trypsin in the absence of phosphate inactivates it (67). Ribonuclease T, is hydrolyzed by pepsin with progressive loss of activity (6‘9). It is not yet possible to coordinate all the information about RNase T, to elucidate the nature of the active center of the enzyme and to explain the mechanism of specific reactions catalyzed by the enzyme. Further studies, including X-ray crystallographic analysis are required. However, if it is assumed that there is a base-specific binding site and a catalytic site in the active center, the former may be essentially different from that of RNase A and the latter may somewhat resemble that of RNase A. It is believed that the residues Glu 58, Arg 77, and His are probably essential to the activity, and that at least one of the histidine residues participates in the catalytic site.

5. Applications Soon after the discovery of RNase T,, it was suggested (70, 71) that it would become an important tool for the elucidation of nucleotide sequence in RNA. Indeed, since 1962 several workers have tried to use the enzyme for the nucleotide sequence analysis of RNA, especially in highly purified specific tRNA’s. Finally, the brilliant research of Holley and his associates in 1965 resulted in the first elucidation of the complete nucleotide sequence of an RNA, alanine specific yeast tRNA, using RNase T, as a main tool (29). Since then many successful elucidations of nucleotide sequence of various RNA’s, using RNase T, as a main tool, followed, and now the enzyme is well-known as an essential tool for the structural analysis of RNA. Ribonuclease TI has been used by several workers to synthesize guanylylnucleosides, oligoguanylate, and other guanosine-containing oligonucleotides with (3’-5’)-phosphodiester bonds (72-79). 67. 68. 69. 70. 71. 72.

C. Shiozawa, J. Biochem. (Tokyo) 66, 733 (1969). K. Takahashi, J. Biochem. (Tokyo) 52, 72 (1962). K . Takahashi, J. Bwchem. (Tokyo) 60, 239 (1966). F. Egami, Kagaku No Ryoiki 12, 9 (1958). K. Sato-Asano and F. Egami. Nature 185, 462 (1960). K . Sato-Asano and F. Egami, BBA 29, 655 (1958).

9.

MICROBIAL RIBONUCLEASES

223

Since recently isolated RNase N, has been shown to be a more suitable enzyme for the same synthetic reactions, the synthetic reactions will be described in the section on RNase N,. However, it should be pointed out that Podder and Tinoco recently demonstrated that the synthetic reactions by RNase TI can lead to the formation of unnatural (2’-5’)phosphodiester bonds ( 4 8 ) . This curious phenomenon occurs when the enzyme and the products are kept a t a higher temperature (100’) (K. Satoh, Y. Inoue, and F. Egami, unpublished). A t normal temperature, (3’4’) -phosphodiester bonds are formed exclusively by the enzyme (79)*

C. RNase Tz Ribonuclease T, ( 5 ) was found in 1957 by Sat0 and Egami in Takadiastase (12). Since the partially purified RNase T, was found to preferentially attack phosphodiester bonds of adenosine-3’-phosphate in RNA ( 8 0 ) ,this enzyme had long been expected to be specific for adenylic acid phosphodiester bond if it could be completely purified. However, sufficiently purified RNase T, showed no absolute base specificity (81, 82) and was found rather to split all phosphodiester bonds in RNA with a preference for adenylic acid bonds. Thus, RNase T, is effective in the analysis of the base composition of RNA. 1. Preparation

The partial purification of RNase T, was carried out by Naoi-Tada et al. (80). Uchida and E’gami (81, 83),and Rushizky and Sober (82) 73. K. Sato-Asano, J . Biochem. (Tokyo) 48, 284 (1960). 74. H. Hayashi and F. Egami, J. Biochem. (Tokyo) 53, 176 (1963). 75. K. H. Sheit and F. Cramer, Tetmhedron Letters 11. 2765 (1964). 76. T. Sekiya, Y. Furuichi, N. Yoshida, and T . Ukita, J . Biochem. (Tokyo) 63, 514 (1968). 77. D. Griinberger, A. Holj., and F. Sorm, Collection Czech. Chem. Commun. 33, 286 (1968). 78. S. C. Mohr and R. T. Thach, JBC 244, 6566 (1969). 79. H. J. Rowe and M. A. Smith, BBRC 38, 393 (1970). 80. M. Naoi-Tada, K. Sato-Asano, and F. Egami, J . Biochem. (Tokyo) 46, 757 (1959). 81. T. Uchida, J . Biochem. (Tokyo) 60, 115 (1966). 82. G. W. Rushizky and H. A. Sober, JBC 238, 371 (1963). 83. T. Uchida and F. Egami, Procedures Nzicleic Acid Res. p. 46 (1966).

224

T. UCHIDA AND F. EGAMI

independently obtained the purified RNase T, as a homogeneous protein. Ribonuclease T, is more stable in acidic pH and heating than other proteins in Takadiastase and has higher isoelectric point and molecular weight than RNase TI.The purification of RNase T, was based upon these properties: Heat treatment in acidic p H or acid extraction was effective for the inactivation and removal of other enzymic activities in Takadiastase, batchwise treatment by DEAE-cellulose or fractionation with ammonium sulfate (40-100% saturation), ethanol (4052%), or acetone (<66%) for the removal of other proteins and concentration of RNase T, activity, and column chromatography by DEAEcellulose or CM-cellulose or gel filtration by Sephadex G-75 for the separation from RNase TI. Ribonuclease T, was purified (900-1 100-fold with a yield of about 10%) from the water extract of Takadiastase A powder by the suitable combinations of several methods described above (81). By the process of DEAE-cellulose column chromatography, RNase T, was separated into two fractions, RNase T2-A and T,-B. However, the only difference is in the nature of their sugar components. 2. Properties Most of the basic properties of RNase T, are summarized in Tables I1 and IV. It is a neutral protein. It is most active at pH 4.5 for RNA or homopolymer digestion and a t pH 6.0-6.3 for the hydrolysis of various nucleoside 2’,3’-cyclic phosphates and for the degradation of 3’-nucleotide beneyl ester (84). Considering that the value was obtained for the first step of RNase T2 action, the degradation of the benzyl esters, the difference of optimum pH may result not from the difference of transphosphorylation and hydrolysis but probably from the difference of molecular size. Indeed this consideration is also confirmed by the finding that the curve of pH dependence of the RNase T, action on commercial RNA does not strictly coincide with the curve of that on the high molecular weight yeast RNA but expands larger than the latter in the region of pH 6-5 (81). The purified enzyme possesses a specific activity of about 1.4 x 103 units/mg of protein. The molecular activity determined for the cleavage of uridine ‘2‘,3’-cyclic phosphate is 15 x lo3 standard units per micromole of protein (83).Onc unit, determined using RNA as substrate, corresponds to about 0.30 standard unit (83). Ribonuclease T2 is most stable a t pH 6.0, 80” for 5 min. At room temperature it is more stalde in alkaline medium and less stable in acidic medium than RNase TI. Ribonuclease T, contains 12-15% carbohydrates, in addition to amino 84. S. Sato,

T.Uchidn, and I?. Egami, A B B 115,

48 (1966).

9.

MICROBIAL RIBONUCLEASES

225

acids. Ribonuclease T,-A can be distinguished from RNase T,-B by the nature of sugar components: RNase T,-A contains mannose and glucose and RNase T,-B contains mannose and galactose as the main sugar components (81). 3. Specificity and Mode of Action Ribonuclease T, is regarded as a nonspecific endoribonuclease [EC 2.7.7.17, ribonucleate nucleotido-2’-transferase (cyclizing) 1. It preferentially splits the internucleotide bonds between the 3’-adenylic acid group and the 5’-hydroxyl group of adjacent nucleotides in RNA, with the intermediary formation of adenosine 2’,3’-cyclic phosphate and splits consequently all secondary phosphate ester bonds of other nucleotides in RNA via the nucleotides 2’,3’-cyclic phopshates. The mode of action on various RNA’s and synthetic homopolymers has been investigated by the 3’-terminal analysis of digestion products (85).I n the exhaustive digestion with RNase T,, RNA is quantitatively digested to produce all nucleoside-3’-phosphates. I n the partial digestion of RNA or tRNA mixture, the mode of action of RNase T, is as follows: (1) Some adenylyl bonds (about 20%) in RNA are preferentially split to produce the large fragments terminating with adenylyl residue in the very early stage of digestion, in which no increase in the optical density of acid-soluble products is observed. (2) Whereas adenylic acid appears rapidly in the initial stage, other mononucleotides appear after a short lag time. The rates of appearance of mononucleotide are in the order of A >> U > 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

T. UCHIDA AND F. EGAMI

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.

MICROBIAL RIBONUCLEASES

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

T. UCHIDA AND F. EGAMI

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.

<

4. Applications

Rushiaky and Sober ( 8 2 ) first described the use of RNase T, for the base analysis of RNA as it hydrolyzes RNA practically completely to 3'-nucleotides. Ribonuclease T, digestion is certainly more advantageous for the base analysis of RNA than alkaline hydrolysis which gives TABLE X

K,

AND

Substrates

V,

VALUESOF RNase Tf

K , (MI

(pmole/min/ pmole protein)

~

UPU UPA GPU GPA CPU CPA APU APG APC APA 4

1.2 x 0.47 X 1.1 x 0.63 x 0.72 X 0.41 x 0.82 X 0.75 x 0.67 x 0.39 x

10-4 lO-' 10-4 10-4 lW4 10-4 lO-' 10-4 10-4 10-4

26 X 23 X 12.0 x 12.7 X 20 x 11.3 X 10.3 X 7.2 x 15.3 X 3.6 X

lo8 lo" 10s 10s 10s 10s 10s 103 lo"

lo"

From Imaaaws et al. (97).

97. M. Imazawa, M . hie, and T. Ukita, J. Biochem. ( T o k y o ) 84, 595 (1968).

230

T. UCHIDA AND F. EGAMI

rise to the mixture of 2’- and 3‘-nucleotides to complicate the analyzing procedure, and which may be accompanied by a slight degradation of nucleotide bases, especially of certain minor components unstable in the alkaline region. Nirenberg et al. (98) applied it to the analysis of certain oligoribonucleotides. A map of the complete digestion products of tRNA showed that all the products were localized in the area of mononucleotides and no oligonucleotide was detected (85). Recently, RNase T, has also played an important role in the discovery and identification of new minor constituents in tRNA: 5,6 dihydrouridylic acid (93), uridine-5-oxyacetic acid (Vp) (86, 94), “;-methyl adenosine (86), and N-9- (P-D-ribofuranosyl) purin-6-ylcarbamoyl threonine (89, 90). Two-dimensional paper chromatography of RNase T, digest was presented by Saneyoshi et al. (86) as the most effective and general method for detection of minor nucleotides in tRNA. Besides these applications, RNase T, has been used for the syntheses of certain nucleoside-3’-phosphates such as coenzyme A (99).

D. RNase N, Although Neurospora crassa had been extensively studied from a genetic point of view, the RNases received little attention until recently. A short report by Suskind and Bonner (100) was the only one before our finding of RNase N, (6). Soon after our finding, genetic studies were reported by Ishikawa et al. (101). Takai et al. (102) studied the formation of ribonuclease, phosphodiesterase, and phosphomonoesterase in cultures of three different strains of N . cl’assa, wild strain 74A and two adenine-requiring strains, in various culture media. They found an extracellular RNase, designated RNase N,, in the culture broth of all strains tested, during the stationary phase. Ribonuclease N, has properties similar to those of RNase TI in regard to base specificity and molecular size (6). However, it has a higher phosphotransferase activity and far less hydrolase activity than RNase T,. As suggested from this finding, RNase N, is more advantageous than RNase T, for the synthesis of oligonucleotides with 5’-terminal 98. M. Nirenberg, P. Leder, J. Trupin, F. Rottman, and C. O’Neal, Proc. Nall. Acad. Sci. U . S . 53, 1161 (1965). 99. A. M. Michelson, BBA 93, 71 (1964). 100. S. R. Suskind and D. M. Bonner, BBA 43, 173 (1960). 101. T. Ishikawa, A. Toh-e, I. Uno, and K.Hasunuma, Genetics 63, 75 (1969). 102. N. Takai, T. Uchida, and F. Egami, Seikagaku 39, 285 (1967).

9.

MICROBIAL RIBONUCLEASES

231

guanylate and those with a definite nucleotide sequence containing guanylyl or inosinyl residue (103, 104). 1 . Preparation The partial purification of RNase N, was carried out by Takai et al. (6,8)from the culture broth (5.5 liters) of Neurospora crassa, adenine-requiring strain 74A-T32-M12. The specific activity of their enzyme preparation was increased to about 5000 times that of the culture filtrate, but their preparation was not yet homogeneous as a protein. It is remarkable that although it was only partially purified, RNase N, had a higher activity than homogenous RNase T,. Kasai et aZ. (105) succeeded in the purification and the crystallization of RNase N,. The culture filtrate of N . crassa was first concentrated by lyophilization. The solution of the lyophilieed powder was fractionated with ethanol (5066%) followed by precipitation with 90% ammonium sulfate saturation. After the resulting precipitate was dissolved in buffer and then dialyzed against 0.005 M Na,HP04, the dialyzed solution was subjected successively to DEAK-cellulose column chromatography, gel filtration by Sephadex G-75, and CM-cellulose column chromatography. This procedure was further simplified a t the Research Institute, Seikagaku-kogyo Co. Their procedure consists of only three steps: the salting-out with ammonium sulfate, DEAE-cellulose column chromatography, and gel filtration by Sephadex G-75. For crystallization, the RNase N, fraction eluted from the CM-cellulose column or Sephadex G-75 column was concentrated by lyophilization and then dialyzed against distilled water at 4". During dialysis, fine needle-shaped crystals were formed. The specific activity of the crystals was about 2200. The solubility of the crystals is small around neutrality and increases below pH 4.0. Thus, the crystals are usually dissolved in dilute acetic acid. 2. Properties

Most of the basic properties of RNase N, (6, 8) are summarized in Tables I1 and IV. It is a neutral protein. It is most active a t pH 7.0 for 103. F.Egami, T.Uchida, T. Arima, and T. Koike, 6th Intern. Symp. Chem. N a t . Prod., London, 1968 Abstr. E-20, p. 266. International Union of Pure and Applied Chemistry, Oxford, England. 104. T. Koike, T. Uchida, and F. Egami, BBA 190, 257 (1969). 105. K. Kasai, T. Uchida, F. Egami, K. Yoshida, and M. Nomoto, J . Biochem. (Tokyo) 68, 389 (1969).

232

T. UCHIDA AND F. EGAMI

RNA and the activity decreases steeply in alkaline medium. The pH dependence of hydrolase activity for guanosine 2’,3’-cyclic phosphate is not yet determined. The crystals of RNase N, possess a specific activity of about 4.0 x lo4 units/mg of protein, about 2.5 times that of purified RNase T,. Ribonuclease N, is most stable in neutral or acid media at 37”. It is resistant even to heating a t 80” for 2 min in acidic media of pH 2 4 , and is fairly unstable in alkaline medium like RNase T,. It is not as stable as RNase T, in higher salt concentrations and exhaustive dilution with water makes the enzyme unstable. This enzyme needs a protective protein such as gelatin or bovine serum albumin for assay. 3. Specificity

Ribonuclease N, is regarded as a guanyloribonuclease [ EC 2.7.7.26, ribonucleate guaninenucleotide-2’-transferase (cyclizing) , Neurospora crassa]. Like RNase T,, it splits specifically the internucleotide bonds between 3’-guanylic acid group and the 5’-hydroxyl groups of adjacent nucleotides in RNA, with the intermediate formation of guanosine 2’,3‘cyclic phosphate The base specificity has been investigated by the 3’-terminal analysis of digestion products of RNA. In the partial digestion of high molecular weight RNA, RNase N, produced almost exclusively guanosine 2’,3’cyclic phosphate and oligonucleotides with guanylyl residues as 3’ terminus (6, 8). Since 3’-guanylic acid was scarcely detected in the RNA digestion products, the hydrolase activity of RNase N, for guanosine 2’,3’-cyclic phosphate was much lower than that of RNase T,. Ten units of RNase T, hydrolyzed 45% of the guanosine 2’,3‘cyclic phosphate (1.4 pmoles/O.5 ml) to 3’-guanylic acid in 6 hr a t pH 7.5, while comparable hydrolysis by RNase N, was produced only with about 150 units of the enzyme in 6 hr a t pH 7.0.

4. Applications Recently, Koike et al. (104) have proved that RNase N, is a more useful tool than RNase TI for the synthesis of oligonucleotides such as guanylyl- (3’,5’)-nucleoside, oligoguanylic acid, and the oligonucleotides of defined sequence containing guanylyl or inosinylyl residue. When guanosine 2’,3’-cyclic phosphate is incubated with about %fold nucleoside a t a low temperature in the presence of RNase N,, guanylyl(3’,5’)-nucleoside can be obtained as a synthetic product. For example, using uridine as a phosphate acceptor, GpU was obtained with a high yield of 27% calculated upon guanosine 2’,3‘-cyclic phosphate added. This

9.

MICROBIAL RIBONUCLEASES

233

yield of GpU synthesized by RNase N, is about twice as much as that of GpU by RNase T, under similar conditions. Furthermore, more than 50% of guanosine 2’,3’-cyclic phosphate remained unchanged, unlike the synthesis by RNase T,, showing the poor hydrolase activity of RNase N, a t lower temperature. The high yield of synthetic product and the possibility of recovering reactants might thus be more advantageous for the practical use of the enzymic synthesis by RNase N,. Ribonuclease N, can also synthesize inosinyl- (3’-5’) -nucleoside from inosine 2’,3’-cyclic phosphate and the nucleoside as phosphate acceptor a t low temperature, like the synthesis of guanylyl- (3’-5’) -nucleoside. Generally, the yield of synthetic products by ribonuclease depends on temperature, pH, incubation time, enzyme concentration, substrate concentration, the ratio of acceptor to phosphate donor, and the nature of acceptor (73, 78, 103, 104). Among these variables, temperature is most important. A low temperature gives a larger amount of the product because it reduces the competing hydrolytic activity; both the rate and extent of synthetic reaction increase when the initial concentrations of the phosphate donor and the ratio of acceptor to phosphate donor are increased (73, 78). Especially when the ratio of acceptor to phosphate donor is large, the production of by-products such as GpGp or GpGpU in the case of GpU synthesis is greatly reduced (104). The optimum p H of synthetic reaction is usually near that of RNA digestion by the ribonuclease (pH 7.0 in the case of RNase Nl). As expected, the rate of the synthetic reaction parallels the enzyme concentration, but the latter does not affect the extent of the reaction (78, 104). The effect of the nature of the nucleoside as a phosphate acceptor on the yield of dinucleoside monophosphates is in the following order: cytosine > uridine > inosine > adenosine. This order has been shown from the results with RNase TI, N,, and U, to be common to all the ribonuclease tested and suggests that the affinity of the dinucleoside monophosphate to enzyme has no relation to the yield of synthetic products in the reaction. The 2’ (3’)-mononucleotides are less effective as acceptors than the nucleosides because of the inhibitory effect of phosphate (78, 104). When RNase N, was incubated only with guanosine 2’,3’-cyclic phosphate (0.1 mmole/O.l ml) in the absence of phosphate acceptor, about 57% of polymerized products consisting of 2-6 guanylyl residues such as GpG cyclic-p (9.1%), GpGp (13.3%), GpGpG cyclic-p (10.20/0), GpGpGp (7.4%), and tetramer (6.5%) were obtained (104). It is characteristic of RNase N, that the oligonucleotides terminating with 2’,3’-cyclic phosphate is produced in higher yield than those with 3’phosphate, unlike RNase T,. The synthetic reaction can be extended to acceptors containing

234

T. UCHIDA AND F. EGAMl

more than one nucleoside residue (T. Koike, T. Uchida, and F. Egami, unpublished), for example G cyclic-p I cyclic-p

-

+ ApG RNaae NI GpApG (7.5% for 46 hr)

+ ApCpC RNaae NI IpApCpC (7.6% for 20 hr)

Furthermore, when ApG cyclic-p was incubated with about 14-fold cytidine at a low temperature in the presence of RNase N,, trinucleoside diphosphate, ApGpC could be obtained with the yield of about 38% after 1 hr of incubation. When ApG cyclic-p alone was incubated with RNase N,, the polymerized products with the definite repeating sequence (ApGp), could be produced. The yield of tetramer (n = 2 ) , ApGpApG cyclic-p ApGpApGp, amounted to 34% and that of hexamer (n = 3), to 7%. But larger polymers were scarcely detected (T. Koike, T. Uchida, and F. Egami, unpublished).

+

E. RNase U, Ribonuclease U, is a novel enzyme found in the culture broth of Ustilago sphaerogena (7, 106). Ribonuclease U, splits, practically specifically, the phosphodiester bonds of purine nucleotides in RNA with the intermediary formation of purine nucleoside 2',3'-cyclic phosphates, indicating the specificity is complementary to that of pancreatic RNase A (106). Like RNase N,, RNase U, very slowly hydrolyzes the intermediate, nucleoside 2',3'-cyclic phosphate, to 3'-nucleotides (SO, 1 0 6 ) . Thus, RNase U, is a useful tool, not only for the analysis of nucleotide sequences of RNA (90,92, 10'7, 108) but also for the synthesis of various oligonucleotides containing adenylyl or guanylyl residue (30) (T. Koike, T. Uchida, and F. Egami, unpublished). 1. Preparation

Arima et a2. (7) have partially purified RNase U, with RNases U, and U, from the culture broth (25 liters) of a strain of Ustilago sphaerogena, from which Glitz and Dekker had isolated a guanyloribonuclease (109, 110). 106. T. Arima, T. Uchida, and F. Egami, BJ 108, 609 (1968). 107. J. M. Adams, P. G. N . Jeppessen, F. Sanger. and B. G. Barrell, Nature 223, 1009 (1969). 10s. N. W.Y. Ho, T. Uchida, F. Egami, and P. T. Gilham, Cold Spring Harbor Symp. Quunt. Biol. 34, 647 (1969). 109. D. G. Glitz and C. A. Dekker, Biochemistry 3, 1391 (1964). 110. D. G. Glitz and C. A. Dekker, Biochemistry 3, 1399 (1964).

9.

MICROBIAL RIBONUCLEASES

235

Ustilago sphaerogena was cultured in a medium containing glucose, glycine, and mineral salts a t 30" for 60 hr. After the cells were removed, the RNase U, fraction was roughly separated from the RNase U, fraction by batchwise treatment of the supernatant with DEAE-cellulose. Other contaminating proteins in the RNase U2 fraction were removed by batchwise treatment with CM-cellulose a t pH 4.0. The resulting RNase U, fraction, after dialysis, was subjected to DEAE-cellulose column chromatography, in which three peaks of RNase activity corresponding to RNases U,, U,, and U:], respectively, were eluted successively. The RNase U, was further purified by rechromatography with DEAE-cellulose and then gel filtration with Sephadex G-75. The RNase U, preparation obtained by this procedure was purified about 3700-fold with a yield of 2.6% from the culture broth of U . sphaerogena. Attempts to develop a simplified procedure are in progress in our laboratory. A large-scale purification procedure has recently been accomplished by Sankyo Co. for the commercial preparation of RNase U,. This procedure mainly consists of repeating the DEAE-Sephadex A-25 column chromatography several times and then using a Sephadex C-50 column chromatography. An RNase U, preparation with the specific activity of 138 could be obtained by this method with a high yield of 22% of the activity of the culture broth. 2. Properties Most of the basic properties of RNase U, are summarized in Tables I1 and IV. It is an acidic protein, most active a t pH 4.5 for RNA digestion (7 ) . It has far less hydrolase activity than phosphotransferase activity (SO). The enzyme preparation obtained possesses a specific activity of about 1.4 x lo3units per OD,,, unit, corresponding to about one-fifth that of RNase T, and twice of that of RNase T,. Ribonuclease U, is as stable as RNase T,. It is fairly resistant to heating a t 80" for 4 min in sodium phosphate buffer, p H 6.9 ( 7 ) . 3. Specificity Ribonuclease U, may be regarded as a purine specific endoribonuclease [ ribonucleate purinenucleotido-2'-transferase (cyclizing) 3 , but the specificity is not absolute. Its mode of action on high molecular weight yeast RNA has been investigated by the 3-terminal analysis of digestion products (SO). As shown in Table XI, RNase U, preferentially splits the phosphodiester bonds of purine nucleotides in RNA. Low concentrations of enzyme (less than 0.5 units/mg of RNA in 0.2 ml) can be used as a nearly purine specific ribonuclease (106'). With about 10-fold larger amounts some phosphodiester bonds of pyrimidine nucle-

236

T. UCHIDA AND F. EGAMI

TABLE XI OF PRODUCTS B Y RNase uz DIGESTION OF 3’-TERMINAL ANALYSIS HIGHMOLECULAR WEIGHT YEASTRNAo RNase UP 0.01 0.107 0.428 1.07 4.28 21.4

53.5 a

(3’-Terminal A/ Total A) X 100

(3’-Termiiial G/ Total G) X 100

(3’-Terminal C/ Total C) X 100

(%I

(%)

(%I

70

46 89 92 96 100 101 95

0 0 0

93 96 100 99 99 100

a 27 57 64

(3’-Terminal U/ Total U) x 100

(%I 0 0 0 0.2 6 20 30

From Uchida et al. (SO). Unit for 1 mg RNA.

otides in RNA are also split. The phosphodiester bond of cytidylic acid is split more readily than that of uridylic acid by RNase Up. According to F. Harada, F. Kimura, and s. Nishimura (unpublished), the phosphodiester bond of uridine-5-oxyacetic acid 3’-phosphate in tRNA can be split by RNase U,. The susceptibility of four nucleotide rcsiducs to RNase U, decreases in the order of A > G >> C > U. Deaminated RNA is more resistant to RNase U, than unmodified RNA. Thus, the presence of the amino group a t the 6 position of purine base (or a t the 4 position of pyrimidine base) seems to be favorable for susceptibility to RNase U,. This suggestion is compatible with the findings that the phosphodiester bond of N - [9- (P-D-ribofuranosyl) -purin-6-ylcarbamoyl] threonine, blocked at the 6 position of adenylyl residue with a large substituent, is resistant to RNase U? (90) and that thioguanosine 2’,3’-cyclic phosphate (240 pg/lOO 1.1) is scarcely hydrolyzed by a large amount of RNase U, (3 units) (S. Irie, T . Uchida, and F. Egami, unpublished). Recently, Hashimoto et al. (35) reported that 7-deazaApU (tubercidylyl- (3’-5’) -uridine) was quite resistant to RNase U2, suggesting that a nitrogen atom a t N-7 of purine base might participate in the preference of substrate to RNase U?. This suggestion was supported by the findings that RNase U2 prefers purine base to pyrimidine base and that methylated RNA including 7-methyl guanylyl residues is also more resistant to RNase Uz than yeast RNA. Both l-methylApU and glyoxalGpU are more resistant to RNase U, than ApU and GpU (SO).Furthermore, it was found that RNase U, does not attack the dinucleoside monophosphates in which N-1 of gua-

9.

MICROBIAL RIBONUCLEASES

237

nylyl residue is blocked with N-cyclohexyl-N'-p- (4-methylmorpholinium) ethylcarbodiimide p-toluene-sulfonate (CMCT) such as CMCGpA and CMC-GpC (108). Considering these results, blocking purine residue a t N-1 appears to decrease its susceptibility to RNase U,. The phosphodiester bonds of xanthylic acid in deaminated RNA were scarcely split by RNase U2 (SO). The susceptibility of purine nucleotide residues to RNase U, decreases in the order of A > 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

T. UCHIDA AND F. EGAMI

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

MICROBIAL RIBONUCLEASES

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

T. UCHIDA AND F. EGAMI

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

T. UCHIDA AND F. EGAMI

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<) than in acidic medium (pH 7 > ) , 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.

MICROBIAL RIBONUCLEASES

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.

MICROBIAL RIBONUCLEASES

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

T. UCHIDA AND F. EGAMI

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).

10.

BACTERIAL DEOXYRIBONUCLEASES

255

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).

256

I. R. LEHMAN

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

257

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).

258

I. R. LEHMAN

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

DEOXYRIBONUCLEASES

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.

BACTERIAL DEOXYRIBONUCLEASES

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

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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.

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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).

266

I. R. LEHMAN

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).

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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).

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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).

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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

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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.

SPLEEN ACID DEOXYRIBONUCLEASE

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).

11.

277

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).

280

GIORGIO BERNARD1

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

ACID DEOXYRIBONUCLEASE

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).

282

GIORGIO BERNARD1

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.

11.

SPLEEN ACID DEOXYRIBONUCLEASE

283

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 ) .

284

GIORGIO BERNARD1

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).

11. SPLEEN

ACID DEOXYRIBONUCLEASE

285

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).

286

GIORGIO BERNARD1

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).

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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

290

M. LASKOWSKI, SR.

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).

12. DEOXYRIBONUCLEASE

I

291

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).

12.

DEOXYRIBONUCLEASE I

293

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

x c 0 c

C 0 V

120

60

Effluent ml

Effluent ml

FIG.1A and B.

I80

12.

295

DEOXYRIBONUCLEASE I

0.2

E c 0 aD N L

0

w 0.I U

0

e

n 5:

a

Effluent ml

FIG.1. Chromatography of various samples of DNase on SE-Sephadex column, 0.9 by 55 cm; temperature, 25" ; initial eluent 0.2 M sodium acetate buffer, p H 4.70; linear gradient with 150 ml each of the initial and limit buffer, 1.OM sodium Enzymic activity. (A) Worthington D P grade DNase acetate buffer, pH 4.70. (0) (amorphous), load 30 mg, linear gradient begun a t 30 ml. (B) Worthington oncecrystallized DNase, load 45 mg, linear gradient begun a t 21 ml. (C) Worthington electrophoretically purified DNase, load 5 mg, linear gradient begun a t 20 ml. From Price et al. (36). Authentic sample of chymotrypsinogen B tested in the same system was eluted between 10 and 35 effluent ml. Chymotrypsin B gave several peaks in the range 15-65 ml.

1.2-

A

A280

0.6 -

mL

100

FIG. 2. Chromatography of DNase I (Worthington D P grade) on phosphocellulose (Whatman P-11, column 0.9 by 70 cm) linear gradient of sodium acetate buffer pH 4.7, 0.38-0.7 M (with respect to acetate), 200 ml of each. From Salnikow et al. (39).

296

M. LASKOWSKI, SR.

TABLE I AMINOACID COMPOSITION OF PANCREATIC DEOXYRIBONUCLEASE PEAKA a - b ~~

~

Nearest integral number of residues per molecule

Residue

Residues per 100 g of protein

Residues Per molecule

Price et al. (Ref. 56)

3.78 2.65 6.08 12.08 4.88 8.41 8.37 2.93 1.75 5.13 8.55 4.33 8.49 8.30 5.47 1.71 1.31 2.34 0.97 0.95

9.14 5.99 12.07 32.81 14.96 29.92 20.00 9.35 9.51 22.37 26.72 11.86 23.26 15.77 11.52 4.05 3.95 3.90 1.82 1.81

9 6 12 33 15 30 20 9 10 22 27 12 23 16 12 4 4 4 2 2

~

Lysine Histidine Arginine Aspartic acid Threoninec SerineC Glutamic acid Proline Glycine Alanine Valined Isoleucined Leucine Tyrosinec Phenylalanine Methioninea Half-cystinee Tryptophan/ Mannose Glucosamine Amide-NH3 Total

Lindherg (Ref. 34) ~~

9 6 12

34 15 30 20 9 9 23 27 12 24 16 12 4 4 4

22 98.589

272

From Price et al. (36). Duplicate analyses by ion exchange chromatography were performed on 24-, 48-, and 72-hr hydrolyzates. Unless indicated otherwise, the average of all of these values was used to compute the values in the second column. Replicate analyses agreed to *2.5y0. A molecular weight of 31,000 was used to calculate the residues per molecule. The average 24-, 48-, and 72-hr values for threonine, serine, and tyrosine have been extrapolated to zero time to correct for destruction during hydrolysis. The average 72-hr values were taken for isoleucine and valine. Methionine and half-cystine were determined in duplicate as methionine sulfone and cysteic acid on the performic acid-oxidized protein [S. Moore, JBC 238, 235 (1963)l. A molecular ratio of 3.80 was determined for tyrosine to tryptophan by the method of T. W. Goodwin and R. A. Morton [BJ 40, 628 (1946)l. g This recovery figure is based on the weight of the samples analyzed after correction for moisture and ash content.

12.

DWXYRIBONUCLEASE I

297

and six residues of mannose. Peak B contains one residue of sialic acid, three residues of N-acetylglucosamine and five residues of mannose. Peak C contains the same carbohydrate as peak A, but one residue less of histidine and one residue more of proline. All three forms have NH, terminal leucine, COOH-terminal threonine. Other changes from the values of Table I are in tryptophan, three rather than four residues, aspartic acid 34 rather than 33, and valine 25 rather than 27. The amino acid analysis (Table I) strongly supports the value of 31,000 for the molecular weight of DNase I. This value has been accepted in Stockholm (34) and in New York (35), making all previously reported values obsolete [see reviews (10, 26, 27, SO)1. The values obtained (Table 11) by ultracentrifugation methods sedimentation diffusion and approach to equilibrium closely agree with the value of 31,000 based on N-terminal determination. New information is also available (35) on stabilization of the DNase I molecule. The presence of 5 mM Caz+fully stabilizes the molecule against proteolytic digestion. For many years it has been known that pancreatic proteases, trypsin and both chymotrypsins, are protected from autolysis by 10mM Ca2+.The protection of DNase I is achieved by a lower Ca2+ concentration. Calcium also exerts a pronounced effect on the reduction of S-S bonds in DNase I. This protein is unusually susceptible to reduction: Both S-S bonds may be reduced within minutes by mercaptoethanol or similar reagents a t pH 7.2 without any denaturing agent (37). The reduced enzyme is inactive and remains inactive even after exposure for 24 hr to oxygen in the presence of ethylenediaminetetraacetate (EDTA) . The situation changes upon addition of 4 m M Ca2+.Activity is regained in minutes. If the Ca2+is present during reduction, only one S-S bond is reduced and activity remains unchanged. If Ca2+is added to a completely reduced protein even in the presence of a reducing agent one S-S bond is re-formed. In spite of exerting a strong stabilizing effect on the enzyme, Ca2+is not strongly bound to the protein. Simple gel filtration a t neutral pH removed .l6Ca2+completely from both the native and the partially reduced forms of the enzyme (37).

111. Active Center

Neither the three-dimensional structure nor the complete primary structure of DNase I has yet been announced. However, a histidine residue has been identified in the active center of DNase I fraction A by

298

M. LASKOWSK1, SR.

SUMMARY OF

THE

TABLE I1 PHYSICAL AND CHEMICAL CHARACTERISTICS OF DNase 1.

Characteristic analyzed

Ei:m a t 280 nm Experimental From amino acid composition Refractive index increment (ml/g)' szO,a ( X 10-13 sec) D20,w( X 10+ cm2/sec) Partial specific volume (from amino acid composition)

Results obtained 12.3b 13.9" 15.3d 0.196 f 0.007f 2.78 8.7 0.733

Molecular weight Approach to equilibrium Sedimentation diffusion N-Terminal determination Isoelectric point Experimental Total nitrogen (%) Experimental From amino acid composition Total sulfur (yo) Experimental From amino acid composition ~~

30 ,900 30,700 29 ,400 31,300 4.79 16.7 f 0.4f 16.2 0.75 f 0.02f 0.81

~

Reprinted from Lindberg (34) by permission of the copyright owner. Copyright 1967 by the American Chemical Society. b In 0.01 M KzHP04-KH2P04buffer, pH 7.6. I n 0.1 N NaOH, pH 13. d This value was calculated using the molar extinction coefficients for the aromatic amino acids tyrosine and tryptophan given by T. W. Goodwin and R. A. Morton [BJ 40, 628 (1946)). 8 Here dn/dc was determined at 20", using the ultracentrifuge as a differential refractometer. f The average deviation about the mean is given based on four determinations. 0 Determined by M. Kunitz [ J . Cen. Physiol. 33, 349 (1950)] and by A. Polson [BBA 22, 61 (1956)j. 0

Price et al. (36).The evidence is based on experiments in which DNase I was reacted with iodoacetate a t pH 7.2 in the presence of 0.1 M Mn2+. Under these conditions the enzyme is gradually inactivated and the loss of activity parallels the formation of one residue of 3-carboxymethyl histidine per molecule. The rate of the alkylation reaction is dependent on Mnz+concentration. Substitution of Mn2+by Cuz+in the presence of tris buffer greatly increases the rate of alkylation. A 29-residue peptide con-

12.

DEOXYRIBONUCLEASE I

299

taining the modified histidine residue has been isolated in 90% yield after tryptic hydrolysis of carboxymethylated DNase I. One structural requirement in the active center can be deduced from a study of specificity with small substrates (see Section VII). Oligonucleotides bearing a 3'-monophosphoryl group are easily cleaved to form d-pNwp (41), whereas neither N u nor pN@is formed. This suggests that a positively charged group (either from lysine or arginine) is placed in the vicinity of an active histidine and immobilizes the negative charge of 3'-monophosphoryl group. One can further speculate that the resistance of d-N"pN0 and d-pN"pN@to the action of DNase I is caused by the lack of such an anchor. However the resistance of a t least some compounds of the type d-pN"pN@cannot be explained without postulating the repulsive effect of the 5'-monophosphoryl group. The long-established preference for the Pu-pPy bond is presumably of lesser structural importance than the phosphoryl group because d-ApApTp can be cleaved to liberate d-pTp (@), whereas d-CpApC is resistant (43).

IV. Inhibitor

A naturally occurring inhibitor of DNase I was originally observed in hypertrophic epithelium of the crop gland of a pigeon (19) and later in several normal and neoplastic mammalian tissues (20, 2 1 ) . The early work has been reviewed (10). For several years, except for sporadic confirmation, the problem remained dormant until Lindberg (22, 23, 4.4, 45) resumed the work and purified two different proteins with inhibitory properties from calf spleen. The spleen inhibitor I1 was crystallized. It forms a uni-uni molecular complex with DNase I under conditions in which either the inhibitor or DNase I is in excess. Figure 3, reproduced from Lindberg's paper (23), illustrates this point. The complex can be irreversibly dissociated (with the loss of inhibitory activity) into its component parts a t pH 7.6 in the presence of 3 M urea. It can also be dissociated a t pH 11.3 and a t pH 3.5 again with loss of inhibitor activity, the activity of DNase I remaining intact. Recently, Lindberg and Skoog (45a) purified DNase I inhibitor from thymus. A 15-fold purification led to a homogeneous preparation. The thymus inhibitor has many properties identical with spleen inhibitor I1 but differs in molecular weight 41. S. Vanecko and M. Laskowski, Sr.,JBC 236, 1135 (1961). 42. J. L. Potter, U.-R. Laurila, and M. Laskowski, Sr.,JBC 233, 915 (1958). 43. H. G. Khorana, J . Cellular C o m p . Physiol. 54, Suppl. 1, 5 (1959). 44. U. Lindberg, BJ 92, 27p (1964). 45a. U. Lindberg and L. Skoog, European J . Biochetn. 13, 326 (1970).

300

M. LASKOWSKI, SR.

0.4

I("

1 ' " A,1

DNase I

p..'

0.2/

,

,

0

,q

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420 I

-0 - 60 -40

0 -20

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+

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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).

12.

DEQXYRIBONUCLEASE I

303

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).

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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.

This Page Intentionally Left Blank

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).

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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.

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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).

322

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).

111. Catalytic Properties

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).

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SPLEEN ACID EXONUCLEASE

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

SPLEEN ACID EXONUCLEASE

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

G. I. DRUMMOND AND M. YAMAMOTO

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

G. I. DRUMMOND AND M. YAMAMOTO

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

G. I. DRUMMOND AND M. YAMAMOTO

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

G. I. DRUMMOND AND M. YAMAMOTO

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.

NUCLEOTIDE PHOSPHOMONOESTERASES

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

G. I. DRUMMOND AND M. YAMAMOTO

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.

NUCLEOTIDE PHOSPHOMONOESTERASES

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

G. I. DRUMMOND AND M. YAMAMOTO

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

G. I. DRUMMOND AND M. YAMAMOTO

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

G. I. DRUMMOND AND M. YAMAMOTO

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.

NUCLEOSIDE CYCLIC PHOSPHATE DIESTERASES

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).

16.

NUCLEOSIDE CYCLIC PHOSPHATE DIESTERASES

365

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).

366

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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NUCLEOSIDE CYCLIC PHOSPHATE DIESTERASES

367

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).

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G. I. DRUMMOND AND M. YAMAMOTO

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

NUCLEOSIDE CYCLIC PHOSPHATE DIESTERASES

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

G. I. DRUMMOND AND M. YAMAMOTO

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.

NUCLEOSIDE CYCLIC PHOSPHATE DIESTERASES

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.

This Page Intentionally Left Blank

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

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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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374

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

ALKALINE PHOSPHATASE

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

T. W. REID AND I. B. WILSON

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.

REID AND I. B. WILSON

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

T. W. REID AND I. B. WILSON

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

C O l i ALKALINE PHOSPHATASE

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

T. W. REID AND I. B. WILSON

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

ALKALINE PHOSPHATASE

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

T. W. REID AND I. B. WILSON

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

T. W. REID AND I. B. WILSON

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 .

C O l i ALKALINE PHOSPHATASE

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.

REID AND I. B. WILSON

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 .

C O l i ALKALINE PHOSPHATASE

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

T. W. REID AND I. B. WILSON

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

111. Catalytic Properties

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

T. W. REID AND I. B. WILSON

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

ALKALINE PHOSPHATASE

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

T. W. REID AND I. B. WILSON

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

ALKALINE PHOSPHATASE

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

T. W. REID AND I. B. WILSON

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

ALKALINE PHOSPHATASE

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

T. W. REID AND I. B. WILSON

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

ALKALINE PHOSPHATASE

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

T. W. REID AND I. B. WILSON

(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

ALKALINE PHOSPHATASE

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 (
406

T. W. REID AND I. B. WILSON

functioning a t high concentrations of substrate. This is consistent with the following models: (1) An enzyme with two nonidentical sites lying along the twofold axis of symmetry and having different binding affinities. (2) A protein with two identical sites which changes into an enzyme with nonidentical sites by a conformational change induced by the binding of substrate to one of the sites. (3) An enzyme conformation with two identical sites in equilibrium with an enzyme conformation with two nonidentical sites and for which the substrate binds best to the conformation with nonidentical sites, pulling the equilibrium in that direction. Other models are also consistent with the X-ray data and the binding and kinetic data, but the three models are relatively simple. Models involving tetramer formation would not fit the above data, because it is found that the tetramer will bind up to 16 phosphates (49). Models 2 and 3 have been proposed by Lazdunski et al. (138) and Simpson and Vallee (134) ; however, their data do not distinguish between models 1 , 2 , and 3. The fact that the second site appears to be metal ion independent would support model 1. It should be noted that models 2 and 3 are thermodynamically equivalent.

F. TRANSPHOSPHORYLATION Some aspects of transphosphorylation have already been discussed (see Section II1,C). In studies with alkaline phosphatase it has been found that the enzymic activity measured by the release of p-nitrophenol from p-nitrophenyl phosphate increases with the concentration of tris buffer much faster than it increases with the ionic strength of other salts such as NaCl and MgrSO, (4, 5 0 ) . This behavior of tris was shown by Dayan and Wilson (122, 123) to result from a transphosphorylation reaction, where 0.5 M tris reacts with phosphoryl enzyme to form tris phosphate a t the same rate as does 55 M water to form orthophosphate. Thus tris is about 100 times better as an acceptor than water. It was later shown that as with other phosphatases ( 1 4 ) many compounds can participate as active acceptors in transferase reactions (see Table I X ) (123, 1.24). Other compounds, not shown in Table IX, which show transferase activity are L-glucose, glucosamine, and butanolamine (124). Compounds which do not show exceptionally marked transferase ability are sodium lactate, fluoroethanol, ethanol, ethylenediamine, catechol, 2140.

R.K. Morton, BJ 70,

139 and 150 (1958).

17. E .

407

C O l i ALKALINE PHOSPHATASE

TABLE IX PHOSPHATE SOURCES FOR VARIOUSACCEPTORS" Transphosphorylation rate: hydrolysis rate

Substance (1M ) Tris Ethanolamine Diethanolamine Triethanolamine Dimethylaminoethanol 3-Aminopropanol-1 L-Serine (0.7 M) L-Serine methyl ester Glycerol Ethylene glycol

0 . 1 M tris +l.OM NaCl

0.1 M tris

1.14 0.50

0.90 0.41 0.88

1.07 0.44 0.31

0.22 0.25 0.26 0.20

0.38

The values given are the ratios of p-nitrophenol to phosphate formed a t pH 8.0 with 0.1 M tris as buffer at 25", minus 1.09 and 1.14 in water and 1 M NaC1, respectively. (I

amino-1-phenylethanol, glycine, 4-aminobutanol-1, 5-aminopentanol-1, and 6-aminohexanol-1 (123). Neumann (124) showed that cysteamine S-phosphate, aminoethanol 0-phosphate, and serine 0-phosphate could serve as phosphate sources for the various acceptors listed in Table IX without changing the percent transphosphorylated product formed for a given acceptor. Barrett et al. (121) found similar results for nine different substrates with tris acting as an acceptor. As previously stated, these results prove that a phosphoryl enzyme intermediate occurs. Consider the scheme for the enzyme-catalyzed hydrolysis a t low substrate concentrations where the reaction is first order in substrate: Ester

kiO + enzyme + phosphoryl enzyme + alcohol (leaving group)

-G

1+'

KE-P

Enzyme

Pi

+ alcohol

Here lclo = kcat/Km (141), KE-P is the equilibrium constant for the hydrolysis of the phosphoryl enzyme (E-P) , and Kesteris the equilibrium constant for the hydrolysis of the phosphate ester. Thus the transphos-

408

T. W. REID AND I. B. WILSON

phorylation constant defining the acceptor ability of any leaving group is given by

Since kcat is fairly constant, with some exceptions, and K, does not vary more than an order of magnitude, there is a tendency for the acceptor ability of a leaving group to be proportional t o the thermodynamic stability of the conjugate ester, i.e., l/Kester.Thus glucose as an acceptor should form more glucose 6-phosphate than glucose 1-phosphate in agreement with studies of Anderson and Nordlie (142). I n the case of alkaline as large as Kester phosphatase KE+ is quite small and in general only for simple phosphate esters. This makes it quite difficult to find acceptors for which trawphosphprylation would be far in excess of hydrolysis. The scheme indicates that the leaving group of every substrate must be an acceptor, or, put another way, every compound must be an acceptor if its conjugate ester is a substrate. It will be a relatively good acceptor if the K , of the conjugate ester is low and if the conjugate ester is relatively stable (it is assumed that kcat will have the usual value). On this basis we should not expect phosphate to be a good acceptor even though the conjugate ester, pyrophosphate, is a good substrate, because pyrophosphate is relatively unstable. Dihydroxy alcohols and amino alcohols in which the functional groups are not separated by more than four carbon atoms are effective acceptors. There is a general difference in the behavior of these compounds, however, in that hydroxyl alcohols decrease the rate of formation of p-nitrophenol from p-nitrophenyl phosphate and amino alcohols increase the rate. Amino alcohols are relatively good nucleophiles in reaction with phosphate esters, perhaps because the amino group serves as a general base catalyst (143, 144). The ability of tris and ethanolamine to serve as acceptors does not vary greatly with pH (30, 123) even though phosphate ester hydrolysis depends very much upon pH. Also, Trentham and Gutfreund (98) found that hydroxylamine does not serve as an acceptor a t pH 8.0, but it does a t pH 5.8. Tris and glycerol are not acceptors with Co(I1) alkaline phosphatase over the p H range 4.5-11, yet the Co(I1) enzyme, like the Zn(I1) enzyme, catalyzes the hydrolysis of different phosphate esters a t the Same rate (91, 94, 96). Also, the K, values of various substrates show little 142. W. B. Anderson and R. C. Nordlie, JBC 242, 114 (1967). 143. C. Dekker and J. Lecocq, Experientia 15, 27 (1959). 144. J. Lecocq, Experientia 22, 361 (1966).

17. E . coli

409

ALKALINE PHOSPHATASE

variation (91, 94, 96). Thus, from examination of Eq. (10) it can be seen that either the equilibrium constant for the hydrolysis of the phosphoryl enzyme is much smaller for the cobalt enzyme than for the zinc enzyme, or tris and glycerol esters have smaller values for kCat/K, relative to k3' for the Co(I1) enzyme than for the Zn(I1) enzyme.

G. KINETICSTUDIES Early kinetic studies on alkaline phosphatase-catalyzed hydrolyses were consistent with the following scheme (106, 123, 128, 1&, 14.6):

E

+

k;

0

II RO-P-OH

I

OH

G k-1

0 II

e E.RO-P-OH I

-

0 II E-P-OH

k;

OH

RI-0-P-OH I OH

+

E

+

I

ROH

OH

E

+

Pi

(11)

This reaction is the same as the hydrolysis of a substrate involving a covalent intermediate ( I l l ? ) , with the addition of a transphosphorylation pathway, where E is the free enzyme, E . S is the enzyme-substrate complex, E-P is the phosphoryl enzyme, and ROH is the acceptor. The reaction is formulated for negligible concentrations of ROH, Pi, and 0 RO-LH

I

OH

so that the reverse reactions need not be considered. The steady state solution of the initial rate of solvolysis of substrate, v = d(ROH)/dt = - d ( S ) / d t , has the form of the Michaelis-Menten equation:

where (E") is the total concentration of enzyme in all its forms. The catalytic rate constant kcat and the Michaelis constant K , are: 145. W. T. Jenkins and L. D' Ari, JBC 241, 295 (1966). 146. W. N. Aldridge, T. E. Barman, and H. Gutfreund, BJ 92, 23c (1964).

T. w REID AND I. B. WILSON

410

For simplification purposes the concentrations of acceptor, R’OH, and water are included in their respective rate constants k,‘ and k3‘. When (R’OH) is set equal to zero, the equations for kCRt and K , reduce to the form for hydrolysis without a second acceptor. A difficulty with Scheme I1 is that there is conflicting data as t o what is the rate determining step. The fact that all phosphate esters are hydrolyzed a t the same rate (see Table VI) would tend to indicate that the rate determining step was the hydrolysis of a common intermediate 0

II

(E-P-OH)

I

OH

and therefore step k,’ would be rate determining. I n agreement with this was the finding that the addition of tris accelerates the rate of release of ROH but not the rate release of Pi (123).Similar results were also found with ethanolamine (30, 123). It is assumed that ethanolamine acts only as a kinetic acceptor; thus, the latter observation would imply that k,’ is much larger than k3‘. If k,’ were not much larger than k,‘ there would be considerable limitation on the availability of phosphoryl enzyme, and dephosphorylation brought about by an acceptor would decrease the level of phosphoryl enzyme and therefore diminish the rate of formation of phosphate. I n disagreement with the above indications was the finding of Aldridge et al. (146) that for enzyme which was phosphorylated a t pH 5.5 with inorganic phosphate and rapidly mixed with buffer a t pH 8.4, the rate of dephosphorylation was twice as fast as the turnover of the enzyme a t pH 8.0. Also, transient state kinetic studies by Fernley and Walker (99, 110) showed a rapid release (burst) of phenol followed by a steady state release of phenol, only a t pH < 7. Thus, these data would seem to indicate that a t pH >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

C O l i ALKALINE PHOSPHATASE

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

T. W. REID AND I. B. WILSON

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

C O l i ALKALINE PHOSPHATASE

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,' << ki. This inequality, since the ratio of ROH to Pi,

makes kpi and K , more or less independent of the concentration of acceptor ( k 6 ' ) .Neuman found that tris increases the rate of hydrolyses of cysteamine S-phosphate and p-nitrophenyl phosphate but does not change K,. Lazdunski et al. (138) have found that a t high concentrations of ATP M ) if the ensymesubstrate solution a t p H 8.0 is rapidly or AMP (> quenched during steady state hydrolysis, one mole of phosphate is bound per mole of enzyme. This result would be inconsistent with both Schemes I11 and IV since it would require dephosphorylation to be rate determining in order to maintain such a high level of phosphoryl enzyme

0

In contrast, Reid and Wilson (139) found that at low concentrations of ATP, PP,, and p-nitrophenyl phosphate M ) if the enzymesubstrate solution a t pH 8.0 is rapidly quenched during steady state hydrolysis, 0.1 mole of phosphate is bound per mole of enzyme. This result would

414

T. W. REID AND I. €3. WILSON

appear to indicate that dephosphorylation is ten times faster than phosphorylation. Thus this would be consistent with Schemes I11 and IV. The scheme for oxygen exchange in Pi based upon the standard scheme for ester hydrolysis (Scheme V) is

The rate of exchange has the Michaelis-Menten form

where

In the reaction the implicit assumption is made that the step ESP"+ E,P removes "0, whereas it is conceivable that during the lifetime of EaP," there might be a reorientation that would enable the removal of other oxygen atoms. Thus the k , used here might be smaller, perhaps by a factor of four, than the value of k, used in the other part of the reaction and in other reactions. If k, is small with respect to k-l, which is probable, this question does not affect the results. It is quite probable that both k , and k-, are small with respect to k-l; therefore, the above equations can be simplified. With this approximation K,,, becomes identical with Ki (see the section on the phosphoryl enzyme). Then the maximum velocity of oxygen exchange (at pH <6) can be compared with the maximum velocity of substrate hydroIysis, where k , would be the rate constant for substrate hydrolysis

+

kcst (oxygen exchange) - k z / ( l kz/k--d - 5 - (E-P) kcat (hydrolysis of S ) k,' Kz (E") The result is that the ratio of maximum velocities is the same as the fraction of phosphoryl enzyme intermediate that is in equilibrium with enzyme and phosphate (see Table VIII). At higher pH for substrate

17. E .

415

C O l i ALKALINE PHOSPHATASE

hydrolysis, where Ic3' is larger than the rate determining step, the equation would be

+

kcat (oxygen exchange) - h / ( l kz/k-2> = 5 5 1 (E-P) ~koat (hydrolysis of S) 6k,' 6 K2 6 (E") where 6 is less than 1 for pH > 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.

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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

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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.

MAMMALIAN ALKALINE PHOSPHATASES

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).

II. Molecular Properties

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.

MAMMALIAN ALKALINE PHOSPHATASES

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

MAMMALIAN ALKALINE PHOSPHATASES

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.

MAMMALIAN ALKALINE PHOSPHATASES

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).

111. Catalytic Properties

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.

MAMMALIAN ALKALINE PHOSPHATASES

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

MAMMALIAN ALKALINE PHOSPHATASES

-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.

MAMMALIAN ALKALINE PHOSPHATASES

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.

MAMMALIAN ALKALINE PHOSPHATASES

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.

MAMMALIAN ALKALINE PHOSPHATASES

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

MAMMALIAN ALKALINE PHOSPHATASES

(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.

MAMMALIAN ALKALINE PHOSPHATASES

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.

MAMMALIAN ALKALINE PHOSPHATASES

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.

MAMMALIAN ALKALINE PHOSPHATASES

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.

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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

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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 . . . . .

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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

VINCENT P. HOLLANDER

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

VINCENT P. HOLLANDER

(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

VINCENT P. HOLLANDER

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

VINCENT P. HOLLANDER

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

VINCENT P. HOLLANDER

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

VINCENT P. HOLLANDER

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

VINCENT P. HOLLANDER

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

VINCENT P. HOLLANDER

- 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

VINCENT P. HOLLANDER

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

VINCENT P. HOLLANDER

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

VINCENT P. HOLLANDER

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

VINCENT P. HOLLANDER

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

VINCENT P. HOLLANDER

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

VINCENT P. HOLLANDER

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

VINCENT P. HOLLANDER

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

VINCENT P. HOLLANDER

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

VINCENT P. HOLLANDER

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

VINCENT P. HOLLANDER

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

VINCENT P. HOLLANDER

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

VINCENT P. HOLLANDER

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

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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.

II. Molecular Properties

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

J. JOSSE AND S. C. K. WONG

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.

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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.

INORGANIC PYROPHOSPHATASE OF

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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).

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INORGANIC PYROPHOSPHATASE OF

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

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INORGANIC PYROPHOSPHATASE OF

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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

J. JOSSE AND S. C. K. WONG

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 <1

loo@ 1lo@ 1600-1800 <1

Recovery of activity

(%I 100 0 50 55 80-90 0

~

From Wong el al. (90). b In the denaturation treatment enzyme was dialyzed a t 24" against the indicated buffer for 7 days. The denatured proteins were then dialyzed a t 3" for 1-2 days against the indicated renaturatiori treatment buffer. Buffers:NaCl-NaPOd: 0.1 M NaCl, 0.01 M sodium phosphate, pH 7;MENaC1-NaP04: NaCl-Nap04 buffer 0.01 M 2-mercaptoethanol; GuHCl: NaCl-Nap04 buffer 5 M guanidine hydrochloride; GuHCl-DTT: GuHCl buffer 0.01 M dithiothreitol; GuHCl-NEM: GuHCl buffer-protein was initially alkylated with N-ethylmaleimide in the presence of GuHCl buffer. c In both of these situations there was opportunity for partial reduction of disulfide bonds before polypeptide refolding and reaggregation began as the guanidine salt diffused out of the dialysis sac. 4

+

+

+

20.

INORGANIC PYROPHOSPHATASE OF

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511

formation (absence of reducing agent, No. 2 of Table 11) or cysteine sulfhydryl groups in the protein were alkylated (No. 6 of Table 11), activity was not recovered (30). The reconstituted enzyme (No. 5 of Table 11) was indistinguishable from original native enzyme by the following criteria: molecular weight optical rotatory dispersion, (sedimentation equilibrium), s&,, D,",,, ultraviolet absorption spectrum, and catalytic behavior (see Section II1,D) (30). Polyacrylamide gel electrophoresis of reconstituted enzyme invariably yielded a single, sharp band which migrated a t the same rate as that of native enzyme (Fig. 4 ) . There was never evidence of the slower moving band noted in some preparations of native protein, even when the enzyme was reconstituted from preparations containing this band (as in Fig. 4). Possibly this second band represented traces of the 130-140-W aggregate detected by electron microscopy (Section I1,E).

(a) FIG.4. Polyacrylamide gel electroplioresis of native ( a ) and reconstituted (b) E. cola pyrophosphatase.

512

J. JOSSE AND S. C. K. WONC

This would be dissociated in 5 M guanidine hydrochloride and obviously does not re-form upon renaturation.

H. CHEMICALCOMPOSITION 1. Amino Acid Content

Results of amino acid analyses are tabulated in Table I11 (34-38), both as numbers of residues per native enzyme particle (MW, 120,000) and as residues per polypeptide subunit (MW, 20,000) ( 3 4 ) .The values listed for half-cystine were obtained from the cysteic acid and S-carboxymethylcysteine contents, respectively, of performic acid-oxidized and iodoacetic acid-treated preparations. The identity of these two results implies that the 12 cysteine residues in the native enzyme are all in the reduced form. This conclusion was confirmed by studies with the Ellman reagent [5,5’-dithiobis (2-nitrobenzoic acid) ] ( 3 9 ) ,which indicated 9-12 moles of reactive sulfhydryl per 120,000 g of protein in the presence of protein denaturing agents (5 M guanidine hydrochloride or 0.5% sodium lauryl sulfate) ( 3 4 ) . However, in the absence of denaturants there was no reactivity whatever, indicating that all cysteine residues in the native enzyme are “buried” or otherwise inaccessible to this reagent. Tryptophan content of the protein was measured with 7 different methods, both spectrophotometric and chemical ( 3 4 ) ; results ranged from 5 to 20 residues per molecule of enzyme, and we have arbitrarily selected a value of 12 tryptophans per protein molecular weight of 120,000. A careful search for trace metals in the enzyme was conducted in the laboratory of Dr. Bert L. Vallee, and none was found in significant amounts. Microanalyses for bound phosphate also gave a negative result. Dialyses against either cation or anion exchange resins or against metal chelating reagents did not alter the sedimentation coefficient or enzymic activity of the protein. Charcoal treatment was similarly without effect (34). 2. NH2-Terminal and COOH-Terminal Amino Acids

The NH2-terminal residue of the enzyme was determined by the cyanate-hydantoin procedure of Stark and Smyth (40) and by Edman 34. P. M. Burton, D. C. Hall, and J. Josse, JBC 245, 4346 (1970). 35. S. Moore and W. H. Stein, “Methods in Enzymology,” Vol. 6,p. 819, 1963, 36. D.H.Spackman, “Methods in Enzymology,” Vol. 11, p. 3, 1967. 37. S. Moore, JBC 238, 235 (1963). 38. R. D. Cole, W. H. Stein, and S. Moore, JBC 233, 1359 (1958). 39. G.L.Ellman, ABB 82, 70 (1959). 40. G.R.Stark and D. G. Smyth, JBC 238, 214 (1963).

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E . coli

513

TABLE I11 AMINO ACID COMPOSITION OF E . C O l i PYROPHOSPHATASE"~b

Amino acid Lysine Histidine Ammonia Arginine Aspartic acid Threonine Serine Glutamic acid Proline Glycine Alanine Half-cystine As cysteic aci& As CM-cysteined Valine Methionine Isoleucine Leucine Tyrosine Phenylalanine Tryptophan' Total

Residues in native enzyme (mole/120,000 g protein)

Residues in subunit (mole/20,000 g protein)

98.0 30.1 (65.3) 25.2 128.1 24.9 47.4 104.7 80.3 55.7 100.7

16.3 5.0 (10.9) 4.2 21.3 4.2 7.9 17.4 13.4 9.3 16.8

12.1 12.0 100.0 17.5 60.9 93.8 47.5 36.5 12.0

2.0 2.0 16.7 2.9 10.2 15.6 7.9 6.1 2.0

1075.3

179.2

From Burton et al. (34). Analyses were performed by Dr. Darrell H. Spackman with a Beckman Model 120B instrument equipped with an automatic integrator (36,36). Results have been corrected for decomposition and recovery losses. The cysteic acid value was obtained from analyses of performic acid-oxidized protein (37). The S-carboxymethylcysteine (CM-cysteine) value was obtained from analyses of iodoacetic acid-treated protein which had been hydrolyzed for 20 and 70 hr (36, 38). The tryptophan value was obtained from independent spectrophotometric and chemical analyses; see text. a

degradation ( 4 1 ) . Serine was the only residue detected, but there were serious difficulties in recovery of serine derivatives with both techniques and quantitation was not possible. A second cycle of Edman degradation yielded 5.6 residues of penultimate leucine [isoleucine (42)] per molecule 41. R. F. Doolittle, BJ 94, 742 (1965). 42. I n the paper chromatographic systems which were employed it was not possible to discriminate clearly between the phenylthiohydantoin derivatives of leucine and isoleucine.

514

J. JOSSE AND S. C. IC. WONG

of enzyme, and the third degradative cycle also yielded leucine [isoleucine (@)I ; the fourth cycle yielded results too obscure to interpret with confidence (34). The COOH-terminal residue, determined both with carboxypeptidases A and B (43), was lysine (5.4 moles/mole of enzyme) , and the penultimate residue was alanine. Additional components were not detected in either digestion (34). 3. Peptide Maps of Tryptic Digests

Protein was denatured either with performic acid oxidation (44) or by heat treatment ( 3 4 ) ,and tryptic digestion was carried out a t pH 7.5. Peptide maps were obtained with two different separation techniques, and in all cases the number of ninhydrin-positive spots was in the range of 17-20 (34). From Table I11 the expected number of tryptic peptides is 21 if the protein consists of 6 identical subunits (16 lysines 4 arginines 1). Selective staining procedures for arginine-, histidine-, tyrosine-, and tryptophan-containing spots yielded slightly fewer than the predicted number of peptides ( 3 4 ) . Altogether these results were consistent with the presence of 6-8 identical subunits in the native enzyme.

+

+

4. Products of Cyanogen Bromide Cleavage

Cyanogen bromide cleavage of the polypeptide subunit should yield 4 peptides if the subunit contains 3 methionines (Table 111).Cleavage products were isolated by gel filtration through Sephadex G-75 columns

and analyzed for purity with polyacrylamide gel electrophoresis in 8 M urea containing 0.1 M thioglycolate. Four peptide products were obtained (34).

I. CHEMICALMODIFICATION OF RESIDUES ESSENTIAL FOR ENZYMIC ACTIVITY All of the studies thus far carried out have clearly implicated lysine residues as essential in maintaining the functional integrity of the enzyme. As with all inhibition studies of the sort described below, a decision cannot be made concerning whether the implicated residues are actually in the active site of the enzyme or if they are indirectly involved in maintaining the protein in a catalytically functional three-dimensional state. It must be emphasized that we have not yet succeeded in identi43. R. P. Ambler, “Methods in Enzymology,” Vol. 11 p. 155, 1967. 44. C . H. W. Him, JBC 219, 811 (1956).

20.

INORGANIC PYROPHOSPHATASE OF

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515

fying groups in the active site of this protein. Experiments with p hydroxymercuribenzoate, N-ethylmaleimide, diisopropylphosphorofluoridate, N-acetylimidazole, and diazonium-1H-tetrazole have been useful in the negative sense in that they have indicated no reactivity whatever of cysteines, serines, tyrosines, or histidines in the native enzyme (4). 1 . Reaction with 2,4,6-Trinitrobenzene Sulfonic Acid

2,4,6-Trinitrobenzene sulfonic acid (TNBS) is known to react with protein lysine residues, forming colored trinitrophenylated (TNP) derivatives with absorption maxima a t 345 mp ( 4 6 ) .When E . coli pyrophosphatase was incubated with a 400-fold molar excess of the reagent, there was progressive decay of enzymic activity accompanied by increasing absorbance of the solution at 345 mp (Fig. 5 ) . Half the enzymic activity had disappeared by the time 16% of the protein lysines had been modified; eventually (6 hr) all 98 enzyme lysines reacted (Table I11 and Fig. 5 ) . Isolation and hydrolysis of the product showed total conversion of all lysine to E-TNP-lysine without formation of any other

Time (rnin)

FIG.5. Reaction of E . coli pyrophosphatase with TNBS a t pH 8.5 in 0.5M NaHCOs. The reaction waa followed in a spectrophotometer, and the concentrations of s-TNP-lysine were determined from the optical density at 345 mp. The solid line shows these data as percentage of total protein lysines; by 6 hr the optical density had stabilized at the level indicated by the arrow on the right-hand ordinate. Aliquots were removed a t the times shown (circles) and assayed for remaining enzymic activity. 45. P.M.Burton and J. Josse, JBC 245, 4358 (1970). 46. T. Okuyama and K. Satake, J . Biochem. (Tokyo)47, 464 (1960).

516

J. JOSSE AND S. C. K. WONG

TNP-amino acids (45). Various levels of TNBS were then reacted with the enzyme, and the inactivation kinetics were analyzed in terms of a multi-order reaction, according to the treatment of Levy et al. (47)

-d[E1 = kl[E][TNBSln (7) dt where E is remaining active enzyme a t time t , k , is the rate constant, and n is the number of TNBS molecules reacting with an enzyme site to inactivate it (E n TNBS 3 E(TNBS),). With TNBS present in excess the process obeys pseudo-first-order kinetics down to as little as 25% remaining activity (Fig. 6 ) . The slopes of the lines in Fig. 6 were in turn analyzed to determine both k , and n ( 4 7 ) . The results ( k l = 86 M-l min-l and n = 1) indicated that interaction of one molecule of TNBS with an enzyme lysine leads to inactivation (4.6). Although it has not been structurally identified, this is almost certainly a particular and specific lysine, but is it in the active site of the enzyme and concerned with binding of anionic substrates and competitive inhibitors (e.g., P P i ; see Section III,E below)? I n the presence of PP,, known to bind strongly to the enzyme active site (Section IIIIE), there was a weak protective effect. The experimental points fell in the shaded area of Fig. 6, and the data were analyzed with equations developed by Scrutton and Utter (48). The results of this treatment led to the conclusion that TNBS can react with both free enzyme and enzyme-PPi complex to cause catalytic inactivation; the differences are only quantitative ( 4 5 ) . Either TNBS can displace PP, from an active site lysine or TNBS modifies a different lysine, apart from the active site, and the presence of PPi on the enzyme partially protects against TNBS inactivation by some indirect mechanism. Unfortunately, as discussed above, this issue cannot be settled with these kinetic analyses. Furthermore, because all of the enzyme lysines are to some extent reactive with TNBS (Fig. 5 ) , the single “superreactive lysine” whose modification leads to inactivation cannot be isolated and identified, as, for example, in a particular peptide fragment. A variety of interpretations are possible, as discussed elsewhere (&) .

+

2. Reactions with Cyanate and with Diazonium-1H-tetrazole

Cyanate is another reagent known to react with protein amino groups (49), and kinetic analyses of inactivation of E. coli pyrophosphatase 47. H. M. Levy, P. D. Leber, and E. M. Ryan, JBC 238,3654 (1963). 48. M. C. Scrutton and M. F. Utter, JBC 240, 3714 (1965). 49. G. R. Stark, Biochemistry 4, 1030 (1965).

20.

INORGANIC PYROPHOSPHATASE OF E .

517

coli

2.0

4.0

40

20 Time Irnin)

FIQ.6. Pseudo-first-order inactivations of E. coli pyrophosphatase by different concentrations of TNBS at pH 8.5 (0.5 M NaHCOd. The number given alongside each least squares line indicates the TNBS concentration in mM. The shaded area represents the zone where the experimental points fell when PPI (0.1-70 mM) was included in mixtures containing 0.5 mM TNBS (46).

have demonstrated a situation similar to that described for TNBS in the preceding section. However, the rate constant [ Ic, in an expression analogous to Eq. ( 7 ) ] was much lower. There was also partial protection against inactivation by PP, (46). Diazotization of proteins with diazonium-1H-tetrazole in previous work has been studied in terms of reaction with histidine and tyrosine residues (50, 5 1 ) . When E . coli pyrophosphatase was incubated with this reagent, there was pseudo-first-order inactivation, but analysis of the 50. H. Horinishi, Y. Hachimori, K. Kurihara, and K. Shibata, BBA 88, 477 (1964). 51. M. Sokolovsky and B. L. Vallee, Biochemistry 5, 3574 (1986).

518

J. JOSSE AND S. C. K . WONG

product indicated that only lysine residues had reacted ; neither histidine nor tyrosine derivatives were detected (46).

111. Catalytic Properties

Escherichia coli pyrophosphatase can be assayed in vitro by means of a simple two-step procedure. The enzyme is first incubated with PPi in the presence of a divalent cation, and the amount of liberated PI is then determined colorimetrically by the method of Fiske and SubbaRow ( l a , 69). PPi

+ H20

2 Pi

(8) As noted in Section I, this reaction is highly exergonic (At"= -5 kcal) and except under special conditions (Section II1,C) essentially unidirectional. When greater sensitivity is required, szPPican be employed, and liberated "PI, separated by paper electrophoresis or chromatography, can be directly counted (18). --t

A. pH EFFECTS

In assay mixtures containing Mg2+the enzyme had a pH optimum of 9.1 with a relatively broad range of activity (20% of maximum activity at pH 7 and a t pH 11)( l a ) . Although the ionization state of the PPI substrate is no doubt important, this alone does not explain the high pH optimum. Titration of a solution containing 0.67 mM Na4PPI and 1.33 mM MgCl, (same as standard assay mixture but without buffer) indicated that ionization of PPI was more than 95% complete a t pH 8.5. As noted in the next section, the pH optimum in assay mixtures containing Zn2+or Co2+in place of Mg2+was lower (pH 7.5).

B. EFFECTS OF IONSAND INHIBITORS There was no detectable activity in the absence of divalent cation. Of the several salts tested, only those with MgZ+,MnZ+,Znz+,or CoZt permitted enzymic activity (Table IV) . The anion was of no consequence as long as the salt remained soluble, but, as described in the preceding section and documented in Table IV, there were definite pH effects. When more than one of these cations were present together, there was augmentation of activity if both were a t less than 0.5 mM; however, 62. C . H. Fiske and

Y.SubbaRow, JBC 66,376

(1926).

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INORGANIC PYROPHOSPHATASE OF

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519

TABLE IV EFFECTS OF CATIONS ON PYROPHOSPHATASE ACTIVITY~ Conc. giving maximal activity

Activity (mpmole PPi hydrolyzed in 15 min by 5 mfig of enzyme)

(a)

Cationb None

w+

1.33 0.67 0.67 0.67

MnP+ Znp+ cop+ a

pH 9.1

pH 7.5

<0.001 98 28 6

<0.001 51 16 77 22

6

From Josee (12).

* Salts with chloride, sulfate, or acetate anions gave identical results. Activity was not observed with Na+, K+, Cu+, Baz+,Caz+,Cuz+,Fez+,Niz+, or Fe3+ (12).

the level of activity was seldom as great as that obtained with either cation alone at twice its original concentration. At concentrations above 0.5 mM there was usually antagonism with lower activity than with either ion alone ( l a ) . Enzymic activity in the routine assay was inhibited by N a F (9% activity a t 1 mM, < 0.01% activity a t 10 mM) and by guanidine hydrochloride (36% activity at 1 M, 3% activity at 2 M ) , but not by KCN (100% activity at 0.1 mM, 87% activity a t 10 d ) or p-hydroxymercuribenzoate (100% activity a t 0.1 mM, 67% activity a t 10 mM) (12). C. REVERSAL OF REACTION

Although exchange of isotopically labeled Pi into PP, has been convincingly demonstrated with the inorganic pyrophosphatase of yeast (53),we were unable to detect exchange of 32P1(1 mCi/pmole) into PPi in the presence of the E . coli enzyme. However, by coupling the system to the very highly exergonic conversion of glucose-6-P to 6-P-gluconate and trapping the synthesized PP,, net reversal was readily demonstrated. 282pi TDP-glucose

+ 3rFszPi

8Zp3'2pi

$ glucose-1-P

glucose-1-P e glucose-6-P

TPN+

+ TPazPs*P

+ glucose-6-P + 6-P-gluconate + TPNH + H+

53. M. Cohn, Biochemistry 2, 623 (1964)

(9) (10) (11) (12)

520

J . JOSSE AND S. C. K. WONG

After E. coli pyrophosphatase and 32Piwere mixed with the appropriate substrates and purified enzymes a t pH 8.5, reduction of TPN+ proceeded as thymidine triphosphate with eventual trapping of 10% of the 32PI (TP32P32P). The net stoichiometry of the coupled reactions was 32Pi, -0.51 pmole; TP32P32P, +0.27 pmole; TDP-glucose, -0.26 pmole; and TPNH, +0.21 pmole (12).

D. SUBSTRATE SPECIFICITY AND STOICHIOMETRY A wide variety of phosphate esters were tested for activity with the enzyme, as measured by release of Pi. Only those listed in Table V served as substrates. The enzymic hydrolyses of purified inorganic tripolyphosphate (P3,i)and tetrapolyphosphate (P4,1) increased linearly with time or with increasing amounts of enzyme, provided that adequate concentrations of substrate and divalent cation were present. Total hydrolysis of each to PI was easily obtained. As with PPi (Table IV), the requirement for divalent cation was absolute, and activity with Mgz+or Mn2+was maximal a t pH 9.1 while that with Znz+or Co2+was highest a t pH 7.5 (18). The stoichiometry of PPI hydrolysis was in accordance with Eq. (8) ; there was no detectable condensation to form P,,,, P,,,, or other higher polyphosphates (12).I n following the course of P3,i or P4,ihydrolysis (using 32P-labeled materials), there was also stoichiometric conversion only to P i ; no traces of intermediate PPi (or P3,iin the case of P4,, TABLE V SUBSTRATE SPECIFICITY OF E . coli INORGANIC PYROPHOBPHATASE~ ~~

Conc. (mM)

Activitp (mpmoles substrate hydrolyzed per 15 min)

Mpl+ Znz+

1.3 0.7

98.0 77.0

9.1 7.5

Mg*+ Znz+

20.0 2.7

1.6 0.5

9.1 7.5

Mgz+ Zn2+

33.3 6.7

0.7 0.2

Conc. (mM)

pH

Cation

PPi

0.67 0.67

9.1 7.5

PJ,i

1.33 1.33

P4.i

2.67 2.67

Substrate

From Josse (12). All activity values are extrapolated to the levels expected for 5 mfig of enzyme. The concentrations of substrate and cation listed are those affording maximal activity without formation of precipitate. a

b

20.

INORGANIC

PYROPHOSPHATASE OF

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521

hydrolysis) could be found ( 5 4 ) . Intermediates occurring during the digestions of these higher polyphosphates are apparently tightly bound to the enzyme, and catalytic hydrolysis presumably occurs stepwise from an end. It is clear from Table V that enzymic velocity diminishes progressively as substrate chain length is increased. Preparation of even higher purified linear polyphosphates was not attempted because of their extreme lability to nonenzymic hydrolysis during isolation procedures; with very large amounts of E. coli pyrophosphatase their degradation might also be enzymically catalyzed. The presence of tri- and tetrapolyphosphatase activities in purified E. coli pyrophosphatase raised the question of whether these were properties of the same protein or of contaminating enzymes. For example, yeast inorganic pyrophosphatase does not act upon P3,i (55) ; there is a separate tripolyphosphatase in that organism (66). Three lines of evidence indicate that hydrolyses of PP,, Ps,i, and P4,, are catalyzed by the same E. coli protein: (1) The three activities purify together with the same relative ratios to one another throughout all stages of the isolation procedure. (2) When the purified enzyme is subjected to ion-exchange chromatography (DEAE-cellulose) or gel filtration (Sephadex G-150) , the three activities elute together with constant ratio in all fractions. (3) When the enzyme is inactivated by heating in the presence or absence of Mg2+, the inactivation kinetics of the three activities are indistinguishable (Fig. 7). The following classes of phosphorus-containing compounds were not affected by large amounts of the enzyme with MgZ+,Mn2+,Zn2+,or Co2+ at either pH 9.1 or 7.5: (1) ribo- or deoxyribonucleoside mono-, di-, or triphosphates ; (2) ribo- or deoxyribopolynucleotides ; (3) nucleotide coenzymes (e.g., DPN+, UDP-glucose) ; (4) phosphomonoesters (e.g., glucose-6-P, p-nitrophenyl phosphate) ; (5) cyclic tri- or tetrametaphos(inorganic phosphorofluoridate, phates; (6) phosphorofluoridates adenosine 5'-phosphorofloridate) ; and (7) phosphonates (e.g., methylenebis-phosphonate) (1.2,57). 54. J. Joese, JBC 241, 1948 (1966).

55. L. A. Heppel and R. J. Hilmoe, JBC 192, 87 (1961). 56. 9. R. Kornberg, JBC 218, 23 (1958). 57. The various compounds were tested in the routine assay under all of the

conditions noted in Table IV; liberation of Pi after incubation with large excesses of purified enzyme was not detected. Compounds of groups (2) and (3), containing phosphodiester or internal phosphoanhydride linkages, were additionally tested with human semen phosphomonoesterase after incubation with E . coli pyrophosphatase. No PI was liberated after this dual incubation, indicating absence of phosphodiesterase or coenzyme-degrading activity in the pyrophosphatase (18).

522

J. JOSSE AND S.

C.

K. WONG

1.0

2. t

.-'5

0.1

4-

0 0

u .-

E, N

C 0)

0

.-m .-

z

,& 0

0.04

c 0 .+ u

?

LL

0.004 Time at 90° (min)

FIU. 7. Inactivation kinetics of (0) pyrophosphatase, ( 0 )tripolyphosphatase, and ( 0 )tetrapolyphosphatase activities at 90". At 80" there was 9Ei-l00% retention of all activities in 0.01 M Mg2+ (10 min), and a t 70" there was 95100% retention of activities even without

Me.

E. NATUREAND BINDINGOF ACTIVESUBSTRATE AND THE ROLEOF MAGNESIUM

It was shown in Section III,B that a divalent cation was essential for enzymic hydrolysis of PP,. The optimal concentrations of cation were of approximately the same level as that of PPr (Table IV) , suggesting that metal ion was necessary for stoichiometric combination with PPi anion. Considering the case of Mg2+and PP:- at pH 9.1, the situation can be analyzed in terms of specific Mg-PP, complexes.

20.

INORGANIC PYROPHOSPHATASE O F

E.

523

COli

Titrimetric analyses have established that K , = 105.4120.06 M-l and K .2 -- 102.34*0.03 M-l (54, 5 8 ) . With use of these values and of conservation equations for both Mg and PP,, the respective concentrations of Mg2+,PPt-, MgPPF, and MgzPPP in any given assay mixture can be readily calculated. Enzymic hydrolysis (PPi + 2 Pi) was measured in each of a large number of solutions in which the concentrations of the various PPI species varied widely. The enzyme velocities so obtained were correlated with these concentrations and analyzed for mathematical fit to a number of possible kinetic models. [At lower pH additional ionic species are present, for example, H P E - and MgHPPt-. However, at pH 9.1 where enzymic activity is maximal (Section III,A), these protonated species are virtually nonexistent and can be neglected.] Some of the data are shown in Fig. 8, where it can be seen that a t a given input level of PPi, activity increases with addition of Mg up to a point; after that activity diminishes with additional Mg. A computer program to analyze these data was designed by Dr. R. A. Dammkoehler of Washington University, St. Louis, Missouri (69). Because so many of the tested kinetic equations were nonlinear and contained interdependent parameters, the computer procedure required extensive trial-and-error strategy to determine optimal global fit for each possible 2500

0

2500

5000 40,000 45QOO

Concentration of Mg

(pM)

FIG.8. Effect of total magnesium concentration (MgtotaI) on the activity of E . coli pyrophosphatase a t different levels of input PPI (PPl,t,t.l). 58. S. M. Lambert and J. I. Watters, JACS 79, 5606 (1957). 59. R. A. Dammkoehler, JBC 241, 1955 (1966).

524

J . JOSSE AND S. C. K. WONG

model. Of all of the equations tested, there was convergence with good fit to give physically meaningful solutions for the kinetic constants in only one basic scheme. EPPt*PPr

TIKi

*MgPP:-

,

E ,pEMgPP'-

-+ +Ha0

E

Mg2+

+ 2 P?-

( 14)

fMgsPP: /]Xi' Kn

EMpPP;

In this model all three of the PP, species of Eq. (13) compete for the same site on the enzyme, but only MgPPT- is the active substrate; PP4- and MgzPPP are competitive inhibitors. The equation for this model is

where v is the measured zero-order enzyme velocity, V,, is the maximum velocity, K , is the Michaelis constant for MgPPq- as substrate, and Ki and Ki' are the competitive inhibition constants for PP4- and Mg,PPP, respectively. The overall mathematical fit of this model to the experimental data is 91.5% (expressed as percentage of variance in experimental reaction velocities which is accounted for by the model). A value of 100% would represent perfect fit. The kinetic constants obtained by the computer for this model are as follows.

Km = 4.9 f 0.7 X 1W'M Ki = 8.5 f 1.6 X 10-8M Ki' = 2.8 f 1.4 X M ,,,Ti = 2800 f 100 units/mg of enzyme (19) (The unit of enzyme activity is defined in Table I.) A very similar model in which no role is assigned to MgzPPY also gives very good fit (90.0%) and virtually identical values for K,, Ki, and V,,,,,. This is not surprising in view of the relatively high value obtained for Ki' in Eqs. (15) and (18). Either Mg,PP! binds to the enzyme relatively weakly or it does not bind at all; the accuracy of the experimental data is not sufficient to differentiate between these possibilities (54). It is clear from this analysis that the actual substrate of E. coli pyrophosphatase is not simply PP, but rather a particular metal complex species which, in the case of catalytic hydrolysis at pH 9.1 in the presence of Mg, is MgPP:-. Much the same conclusion has been reached in earlier

20.

INORGANIC PYROPHOSPHATASE OF

E.

525

COli

studies of inorganic pyrophosphatases from other sources (60, 61). Free PP, (not combined with metal = PPf-), although it is bound to the active site of the enzyme roughly 50 times more strongly, is not a substrate, nor is the MgzPPb species, which may or may not bind weakly to the protein. The complete absence of activity when milligram amounts of enzyme are mixed with PPI alone (Table IV and Fig. 8) is now easily explained on two counts. First, divalent cation is required in stoichiometric amounts to form the active substrate; second, there is powerful competitive binding of enzyme by excess PPj-. Inhibition of activity with excess magnesium (Fig. 8) is accounted for by sequestering of PP, as inactive Mg2PPq according to Eq. (13).

F. INTERACTIONS WITH INHIBITORS Crude estimates of the affinities of the enzyme for other compounds have been made by study of their capacity to inhibit hydrolysis of PPI. If the observed inhibition is assumed to be competitive, a simplified kinetic treatment yields the inhibition constants (Ki" values) listed in Table VI (54).The data indicate that several of the compounds whose hydrolysis is not catalyzed (Section II1,D) are nevertheless bound weakly to the enzyme (e.g., ADP, methylene-bis-phosphonate). There is also very weak binding of PI, the product of the enzymic reaction [Eq. TABLE VI COMPETITIVE INHIBITION OF PPi HYDROLYSIS' Inhibitor Pi Pa,i p4.1

Trimetaphosphate Tetrametaphosphate AMP ADP ATP Methylene-bis-phosphonate 0

300-600 15-36 110-230 220-460 280-580 01

110-210 OD

250-500

From Josse (64).

60. E. Bauer, 2. Physbl. Chem. 284, 213 (1937). 61. E. A. Robbins, M. P. Stulberg, and P. D. Boyer, ABB 54, 215 (1965).

526

J . JOSSE AND S. C . K. WONG

(S)], but its estimated affinity constant is 100 times that of substrate MgPPi- ( K , = 4.9 x M).

IV. Conclusions

Only a limited amount of information about the active site of this enzyme can be inferred, and even less can be deduced about the catalytic mechanism of action. The active site probably carries a positive charge at pH 9.1 because the strength of binding of small molecule substrates and inhibitors a t this p H can be directly correlated with the amount of negative charge on the ligand. For example, PPj- is bound to the enzyme very strongly, although it is not a substrate ( K i = 8.5 x M ); MgPP:-, the active substrate, is bound less well ( K , = 4.9 X M ); and Mg2PPo is bound very weakly if a t all. Positive charge on a protein at pH 9.1 must be carried either on arginine or lysine residues. The chemical modification studies described in Section II,I did implicate an essential lysine residue in the protein, but whether or not this lysine was in the active site of the enzyme could not be ascertained. The site is apparently restricted in “size.” Among the polyphosphate substrates (PP,, P3,i, and P,,,), binding as well as velocity diminishes markedly as chain length increases (Tables V and VI). However, once the longer polyphosphates are attacked, their hydrolysis products, which become increasingly more strongly bound as chain shortening proceeds, apparently remain on the enzyme until degradation to Pi is complete (Section II1,D). Enzymic hydrolysis, as contrasted to the binding phenomena just described, has more stringent restrictions, Substituents on the phosphate groups are not tolerated (except for additional phosphate moieties as with P , , , ) ; for example, ADP is weakly bound (Table V I), but it is totally inactive as substrate (Section II1,D). More importantly, divalent cation plays a central role which apparently is not primarily concerned with binding. Only the MgPPT- complex serves as substrate even though it is bound less well than PP;-. After hydrolytic cleavage of the pyrophosphate linkage has taken place on the enzyme surface, the products of the catalytic reaction, P:- and Mg2+, readily dissociate from one another and from the enzyme. The exact role of the divalent cation in the enzymic mechanism is not known. The effect may be primarily one of electron induction with withdrawal of electrons from the phosphorus nucleus, rendering it more susceptible to nucleophilic attack by oxygen of water (61). Possibly steric factors (restrictions on the rotational con-

20.

INORGANIC PYROPHOSPHATASE OF

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527

formations of the two central P-0-P pyrophosphate bonds in the MgPP?- complex) are also important (64). Compared to most other enzymes, this protein catalyzes a relatively simple chemical reaction and with very rigid specificity and high efficiency (turnover number, 1.5 X lo6). Data from the kinetic studies cited in Section II1,E can be interpreted in terms of straightforward Michaelis-Menten analyses without resort to considerations of allosteric effects or subunit interactions. Yet the enzyme protein has a somewhat complex structure and appears to consist of six identical polypeptide chains arranged to form a very stable globular particle. Results of unpublished equilibrium dialysis experiments have been inconsistent, and it cannot yet be decided how many PP, binding sites are present on the native enzyme particle of molecular weight 120,000. The problem seems to have reached the point where further definitive structural characterization of the enzyme must await two very expensive and laborious undertakings : sequence determination of the polypeptide subunit and high resolution X-ray diffraction analysis of the three-dimensional structure of the protein molecule. Both opcrations would appear technically feasible in view of thc size of the subunit (molecular weight, 20,000) and of the tendency of the protein to form large, cubic-type crystals. ACKNOWLEDQMENTS The experimental work described in this chapter represents the joint efforts of several colleagues: Drs. Dennis C. Hall, Pamela M. Burton, and Robin C. Valentine (deceased), Mr. Fletcher Smith, and Mrs. Ann Hoyt.

This Page Intentionally Left Blank

Yeast and Other Inorgartic Pyrophosfihatases LARRY G. BUTLER

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

1. Introduction. 11. Yeast Inorganic Pyrophosphatase A. Molecular Properties . . B. Catalytic Properties . . 111. Other Inorganic Pyrophosphatases

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

. .

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

529 530 530 534 539

I. Introduction

A great variety of biosynthetic pathways are dependent on reactions which release inorganic pyrophosphate ( PPI) ; a convenient summary of some of these pathways has been provided by Kornberg ( 1 ) . The widely distributed enzymes which are capable of hydrolyzing this PPL to orthophosphate (Pi),thus driving the biosynthetic reactions toward completion by removal of a product, were previously reviewed in the second edition of this series (a). Until relatively recently, the only inorganic pyrophosphatase which had been characterized and studied in a systematic, detailed manner was Kunitz’ crystalline preparation from baker’s yeast (3)which is the main subject of this chapter. Extensive investigations of this yeast enzyme are currently in progress in a t least 1. A. Kornberg, in “Horizons in Biochemistry” (M. Kasha and B. Pullman, eds.), p. 251. Academic Press, New York, 1962. 2. M. Kunitz and P. W. Robbins, “The Enzymes,” 2nd ed., Vol. 5, p. 169, 1961. 3. M. Kunitz, J . Gen. Physiol. 35, 423 (1952). 529

530

LARRY G . BUTLER

two laboratories ; because these studies are incomplete a definitive survey cannot be provided at this time. Researches on microsomal and E . coli inorganic pyrophosphatases are treated separately in Chapter 22 by Nordlie and Chapter 20 by Josse and Wong, this volume; other pyrophosphatases are reviewed briefly in the last section of this chapter.

II. Yeast Inorganic Pyrophosphatase

A. MOLECULAR PROPERTIES 1. Purification Kunitz ( 4 ) simplified his original method (3) for the isolation of pyrophosphatase from baker’s yeast and its crystallization from 22% alcohol by incorporating the alcohol fractionation procedure of Heppel and Hilmoe ( 5 ) . The rather inconvenient alcohol fractionation has more recently been supplanted by chromatographic techniques (6-8) ; in particular. Cooperman has elegantly exploited the difference in chromatographic properties of the enzyme in the presence and absence of Mg2+ to obtain efficient purification ( 6 ) .A simple procedure for crystallization of the enzyme from ammonium sulfate solution in a stabIe, homogeneous form has been developed by Ridlington and Butler (8). Although direct comparisons have not been made, no marked differences have been observed in the properties of enzymes obtained by the different methods (9). 2. Physicochemical Parameters

Some of the physicochemical parameters of the enzyme, as determined on the Kunitz preparation (S),are summarized in Table I. Sedimentation equilibrium analysis of molecular weight carried out on the enzyme crystallized from ammonium sulfate gave a value of 71,000 ( 8 ) , in fair agreement with the value of 63,000 obtained by sedimentation velocity measurements on the Kunitz preparation ( 1 0 ) . The enzyme has been reported to dissociate into subunits in the presence of sodium dodecyl 4. M. Kunitz, ABB 92, 270 (1981). 5. L. A. Heppel and R. J. Hilmoe, JBC 192, 87 (1951); L. A. Heppel, “Methods in Enzymology,” Vol. 2, p. 570, 1955. 6. B. Cooperman, manuscript in preparation. 7. S. M. Avaeva, E. A. Braga, and A. M. Egorov, Biofizika 13, 1126 (1988). 8. J. Ridlington and L. Butler, manuscript in preparation. 9. 0. A. Moe and L. Butler, manuscript in preparation. 10. H. K. Schachman, J . Gen. PhysioZ. 45, 451 (1952).

21. INORGANIC

531

PYROPHOSPHATASES

TABLE I PHYSICOCHEMICAL PARAMETERS" Diffusion coefficient (water, Z O O ) Sedimentation constant (water, 20") Molecular weight (sedimentation velocity) Electrophoreticmobility (0.01 M cacodylate, pH 6 . 7 , 0 . 1 8 M NaCl) Weight intrinsic viscosity Optical density (280 nm, 1 mg/ml) Isoelectric point

6 . 8 X 10-7 cm*/sec 4.4s 63,000 - 1 . 8 x 10-5 cm/sec/V/cm 3.3 1.45 4.75

0

Data, from Kunitz (3)and Schachman (10).

sulfate (SDS) (7');the apparent molecular weight as determined by gel electrophoresis in the presence of SDS is 35,000 (8). Thus, the native enzyme very likely consists of two subunits of similar molecular weight. Gel electrophoresis in 8 M urea rather than SDS gave a single protein band, suggesting that the subunits, if not identical, are electrophoretically similar (8). Analysis in two different laboratories of the amino acid composition of the enzyme crystallized from ethanol gave similar results with the exception of the values for cysteine. Hausmann (11) found only a trace, but Eifler et al. (12) reported 3 cysteine residues per mole of 60,000 molecular weight. The latter investigators found that three different sulfhydryl reagents reacted with the enzyme in amounts approaching one mole per mole of enzyme. A total of three -SH per mole were found after reduction with sodium borohydride. The investigators suggested that the molecule contains one -S-S- and one -SH, which, if true, forbids identity of subunits. The essentiality of a free -SH for activity has not been established (8, 1 2 ) . Maleimides and mercurials are somewhat inhibitory, but neither is absolutely specific for -SH groups. Iodoacetate and oxidizing agents such as ferricyanide are not inhibitory. Under certain conditions the enzyme is inactivated by reducing agents such as dithiothreitol, suggesting the presence of a 43-S- which must be intact for activity. Reactivation in oxidizing conditions of enzyme inactivated by reducing agents has not been observed (8).

3. Reversible Binding of Divalent Cations The enzyme is considerably more stable in the presence of certain divalent cations than in their absence, suggesting binding of the cations to the protein (8). Cooperman found that the relaxation rate of the 11. W. Hausmann, JACS 74, 3181 (1952). 12. R. Eifler, R. Glaesmer, and S. Rapoport, Acta Biol. Med. Ger. 17, 716 (1966).

532

LARRY G . BUTLER

nuclear magnetic resonance of water protons in the presence of the paramagnetic ion Mn2+ is enhanced 12-fold on binding of the Mn2+to the protein (6). By using this technique, Cooperman found 2 Mn2+binding sites per mole of dimer, with dissociation constant KD = 9.0 x M. For these sites, Mg2+ competes with MnZ+;the corresponding KD’s for Mg2+,Znz+, and Ca2+are 7.2 x M , 5.7 x M , and 16 x M, respectively. The apparent binding of Mg2+ and Ca2+ has also been estimated indirectly by measuring their ability to protect the enzyme against inactivation by reducing agents as a function of Mg2+or Ca2+ concentration (8). The KD’s for Mg2+ and Ca2+estimated in this way M , respectively. The cation specificity of M and 7 x are 5 X this protection differs considerably from the specificity as activator of the catalytic reaction. While Mg2+and Ca2+can give complete protection, Mn2+and Go2+ are only marginally effective, and Zn2+destabilizes the enzyme (8). 4. Metalloenzyme

I n addition to reversibly binding to the enzyme and to formation of a complex with PPi which is the actual substrate (see below), a divalent tation may be involved in yet another way in this simple hydrolase enzyme ( 1 3 ) . Observation of time-dependent inactivation by EDTA and other chelators suggested the possibility that the enzyme contains tightly bound metal ions necessary for activity. The relative effectiveness of several chelators in causing this inactivation parallels the strength of their association with Mg2+ more closely than that of other ions. Direct analysis for Mgz+by atomic absorption spectroscopy on several enzyme preparations after exhaustive dialysis against 1 mM EDTA, pH 7.2, a t 3” (under these conditions there is negligible loss of activity) indicated the presence of approximately one mole of Mg2+per mole of active enzyme dimer. At higher temperatures and pH the strongly bound Mg2+is removed by EDTA treatment, with loss of activity. All chelator-dependent enzyme inactivation is presumed to result from removal of Mg2+,but because of the large amount of enzyme required for measurement of protein-bound Mg2+ this correlation has so far been established under only one set of conditions. No reversal of the inactivation by addition of excess Mgz+ or other cations has yet been observed. Although these observations are very recent and the system has not yet been studied intensively, it appears at this time to conform to Vallee’s rigorous definition of a metalloenzyme (14). 13. J. Ridlington, Y. Yang, and L. Butler, manuscript in preparation, 14. B. L. Vallee, “The Enzymes,” 2nd ed., Vol. 3, Part B, p. 22.5, 1960,

21.

633

INORGANIC PYROPHOSPHATASES

The rate of inactivation by chelators is strongly dependent on temperature, pH, and protein concentration (IS). Between 16" and 30" the activation energy for the chelator-dependent loss of activity is 41 kcal at pH 8.2. At 30" the rate of inactivation is over 200-fold faster a t pH 8.7 than a t pH 7.2. The inactivation is much faster in dilute than in concentrated enzyme solutions ; as the protein concentration is increased, correspondingly more rigorous conditions must be employed to observe inactivation. The rate of inactivation appears to exhibit saturation kinetics with respect to chelator concentration. At high EDTA levels the inactivation rate approaches a maximum which is independent of chelator concentration (IS). Titration with chelators of a metalloenzyme preparation from which extraneous metals and chelators have been removed produces a characteristic enhancement of the intrinsic protein fluorescence (excitation at 280 nm, emission a t 350 nm) (IS). This fluorescence enhancement by nonfluorescent chelators is instantaneous, reversible by excess added divalent metal ions, and can occur without loss of activity. Different chelators give different characteristic amounts of fluorescence enhancement a t saturation, demonstrating the specific effect of the chelator on the fluorescence of the apparent metalloenzyme-chelator complex. I n contrast, if the effect of chelators were simply to complex with Mg2+ after its dissociation from the metalloenzyme, the resulting apoenzyme should have identical fluorescence properties regardless of which chelator was utilized. The above observations are consistent with the following model for the metalloenzyme-chelator system: E(Mgz+)

+C

KD

'k

E(Mg2+C)--t P

+ Mgz+C

Metalloenzyme is represented by E(MgZ+) , C is chelator, E (Mg*+C) represents chelator reversibly bound to the metalloenzyme with dissociation constant K , and with characteristic enhancement of the protein fluorescence, and k' is the first-order rate constant for irreversible formation of inactive protein P and the cation-chelator complex, MgZ+C.Values of both K , and Ic' may vary according to the nature of the chelator, the nature of the metal ion (if other cations can replace Mg2+),and the intrinsic stability of the cation-chelator complex, as well as pH, temperature, and protein concentration. Values of K , have been estimated by fluorescence titration and by measurement of the rate of inactivation as a function of chelator concentration (IS). In general, those chelators that form the most stable complexes with Mg2+ also bind most strongly to the metalloenzyme, though far less strongly than to free Mg2+.Chelators with bulky non-

534

LARRY G. BUTLER

functional groups are relatively less effective than compact chelators such as EDTA; this may reflect steric limitations imposed by the protein molecule a t the metal binding site. Several important questions about the structure are yet to be resolved. The subunits appear to be electrophoretically identical, and the dimer contains two Mnz+ binding sites, consistent with subunit identity. If the dimer contains three cysteine residues, however, the subunits cannot be identical. Recalculation of the results of -SH analysis using the more recent molecular weight of 71,000 (8) rather than 60,000 (12) gives close to four -SH per mole, allowing for identity of subunits. The finding of one free -SH and one Mg2+per mole of dimer argues against identity of subunits, but there may be a common explanation: The bound Mg2+ may protect a -SH group. If the enzyme contains one -&& per mole of dimer this must either link the subunits or the subunits are not identical. It is not yet known how or whether dissociation of subunits is related to Mg2+ removal. It does seem likely that the role of the tightly bound Mgz+ is structural rather than catalytic. The Mg2+ might aid in maintenance of dimer structure by binding between the monomers, but binding of chelators to such a shared Mgz+would not be expected.

B. CATALYTIC PROPERTIES 1 . Assay The enzyme is routinely assayed by a modification of the Fiske and SubbaRow technique for measuring Pi production from PPi (8). Cooperman has developed a continuous automated pH-stat pyrophosphatase assay which is very convenient for repetitive analyses (6). Kinetic studies over a wide range of conditions require a more sensitive assay such as the production of 32Pifrom szPPi,as used by Moe and Butler (9). 2. Specificity a. Substrate. With Mg2+as the activating cation the enzyme is quite specific for hydrolysis of PP, ; inorganic polyphosphates and metaphosphates are not cleaved (3,15).In the presence of Zn2+the specificity is broadened to include a wide variety of organic pyrophosphates (16-18) 15. L.A. Heppel and R. J. Hilmoe, JBC 192, 87 (1951). 16. M.J. Schlesinger and M. J. Coon, BBA 41, 30 (1980). 17. D.L. Miller and F. H. Westheimer, JACS SS, 1511 (1966). 18. S. M. Avaeva, S. N. Kara-Murza, and M. M. Botvinik, Bwkhimiya 32, 205 ( 1967).

21. INORGANIC

PYROPHOSPHATASES

535

but not acyl phosphates (19). Inorganic ortho-, meta-, and polyphosphates and organic phosphates appear to inhibit PPi hydrolysis only to the extent that they compete for Mg2+ (9, 16). The substrate analog methylene diphosphonate, which structurally resembles PP, rather closely (go), is not inhibitory in the presence of Mg2+;in the presence of Mn2+, it is a weak competitive inhibitor (6). Hydroxymethylene diphosphonate is strongly inhibitory with MnZ+and weakly inhibitory with Mgz+ (6). b. Metal Zons. The relative effectiveness of various divalent cations as activators of the enzyme depends upon the pH, concentration of cation, and ratio of cation to PP, ( 3 ) . Under conditions near optimal for Mg2+but which may be far from optimal for other cations, the relative activities are Mg2+> Zn2+> Co2+> Mn2+>> Ca2+ (9). In the presence of Mgz+ most other cations are inhibitory. The decreasing order of effectiveness as inhibitors of the Mgz+-activated reaction is Ca2+> Cd"' > Mn2+> Co2+> Ni2+> Znz+. It appears that for Ca2+,a t least, the inhibitory form is the Caz+-PPl complex (9). The relative effectiveness of the divalent cations as activators or inhibitors and in stabilization of the activity does not correlate well with ionic radius or other known properties of the cations. There are interesting contrasts in the specificity of cations as activators or inhibitors of the reaction and their specificity for stabilization of the activity of the enzyme. The most effective activator, Mg", and the most effective inhibitor, Ca2+,have a similar capacity to stabilize the enzyme; the second most effective activator, Znz+, is completely ineffective in stabilizing the activity (IS). 3. Kinetics

The different ionic forms of PP, which are present in significant amounts a t pH 6-9 in the presence of Mg2+, as activating cation, and KCl, to adjust the ionic strength, are shown in Fig. 1. Moe and Butler have attempted to assess the catalytic significance of these forms and of free Mg2+which is also present (9). The values of the equilibrium constants for all these interactions have been evaluated under conditions similar to those of the pyrophosphatase assay (bl), and a computer program has been devised to calculate the concentration of each of these ionic species as a function of pH and of total added PPi and MgC1, (9). Rates of PP, hydrolysis were determined over a wide range of Mg2+and PP, concentrations a t pH 7.4, 8.1, and 9.0. The observed rates of PPi hydrolysis and computed concentrations of the ionic species 19. J. Sperow and L. Butler, ABB, submitted for publication. 20. M. Larsen, R. Willett, and R. G . Yount, Science 166, 1510 (1969). 21. S. M. Lambert and J. I. Watters, JACS 79, 5606 (1957).

536 KPPi-3

LARRY G . BUTLER

-

- PPi-4

--

MgPPi-2

-

-

MCzPPi"

L

HZPPi-'

RG.1. Ionic forms of PP, which are present in significant amounts at pH 6-9 in the presence of MgCL and KCl. containing Mg2+ and PP, were analyzed to determine which of several plausible kinetic models for the reaction gives the best fit to the data. The data were analyzed by two different techniques: estimation of apparent values for kinetic constants by graphical methods with subsequent refinement by trial and error fitting to the kinetic equations ( 9 ) , and by a computer program for nonlinear regression ( 2 2 ) . The two techniques gave values for kinetic constants which are in good agreement (9). The only model which provides an excellent fit (RZ= 0.99) (22) for the kinetic data and which is, in addition, consistent with all other available data is shown in Fig. 2 along with the velocity expression and the computer generated values for the kinetic constants. The proposed model involves activation by free Mg2+before the enzyme can bind substrates or inhibitors. Uncomplexed PPi is a strong competitive inhibitor, and two different Mg2+-PPi complexes are hydrolyzed a t kinetically distinguishable rates. This latter feature, which may be unique to this enzyme, is illustrated in Fig. 3, which presents a portion of the kinetic data. At extremely high Mg2+ concentrations where effectively all the PPi is present as the di-magnesium complex the velocity levels off to a constant value instead of decreasing to zero a s the concentration of the monomagnesium species diminishes to extremely low values. Kinetic studies of inorganic pyrophosphatases from mouse (25) and rat (24) liver cytoplasm have previously implicated free Mg2+ as an 22. J. Jew, JBC 241, 1948 (1966). 23. A. Horn, H. Bornig, and G . Thiele, European J . Biochem. 2, 243 (1967). 24. M. Irie, A. Yabuta, K. Kimura, Y. Shindo, and K. Tomita, J . Biochem. (Tokyo) 67, 47 (1970).

21.

537

INORGANIC PYROPHOSPHATASES

1 PPi

I

EM(M,PP,)

-

EM

+

Products

V , ( A / K , ) + Vb(B/Kb)

v =

I

+

A/Ka

+ BIK, + IIK, + K , IM

x x 4.9 x 3.8 x

K , = 4.9

IO-bM

K b = 5.4

10-6ll.I

K, = K, =

10-6 M 10-3ICI

V r / V a = 0.28 F I G . 2. Proposed kinetic model for yeast inorganic pyrophosphatase. Here M represents MgZ+but may also apply to any divalent cation with which the enzyme is active. I n the rate equation A represents all mono-magnesium PPi complexes, B represents the di-magnesium complex, and I represents free PPi. Hydrogen ion equilibria are not considered. Kinetic runs were done at pH 7.4, 30” (9). Best values for kinetic constants were obtained from a computer program for nonlinear regression (2.%?).

activator of the reaction. It is not yet clear whether the kinetic effect of free MgZt on yeast pyrophosphatase corresponds to the previously described binding of Mg2+and other metal ions, for there are as yet unexplained discrepancies in the values of the dissociation constants obtained by the two types of measurements (see Section II,A,3, and Fig. 2 ) . In any case it seems likely that “activation” of the yeast enzyme by free Mgz+actually involves formation of a binding site for free PPI (inhibitor) and the magnesium-PP, complexes (substrates). Cooperman has recently obtained results with the NMR technique which indicate that Mn2+probably serves as a bridge between PPi and the enzyme when PPi is bound in the presence of Mn2+(6). At present it is not known whether binding of the cation-PPi complexes is on the previously bound divalent

538

LARRY G. BUTLER

-log [MgCl,1,

M

FIG.3. Rate of PPI hydrolysis as a function of total added MgCl,. Experimental conditions: 0.45M tris (Cl-), pH 7.4, ionic strength adjusted to 1.0 with KCl, 30", 2.58 X lod M PPI. Experimental data are shown by the points. The solid line is that predicted by the kinetic model described in Fig. 2; the broken line is that predicted by a model which best fits data obtained with E. coli inorganic pyrophosphatase (22), which postulates no effect of Mg,PPi or free Mg* on the reaction rate.

cation or at some other site which is created as a result of the previous binding of cation to the metalloenzyme. 4. Mechanism

Cohn observed that yeast pyrophosphatase catalyzes exchange of oxygen between Pi and water a t a rate which is five hundred times as fast as the rate of reversal of the overall reaction (26). On the basis of the oxygen exchange activity, Cohn classified the enzyme as a phosphoryl-transferring enzyme. However, compounds other than water are not phosphoryl acceptors (6, 9). Even in the presence of Zn2+no detectable transfer of szP from 32PPito ADP is observed under conditions where ADP must be bound to the enzyme because it is a substrate and competes with s2PP, (9). The rigid specificity for water as phosphoryl acceptor suggests the presence of a specific acceptor binding site which cannot effectively accommodate compounds other than water. The rapid oxygen exchange is consistent with a mechanism involving a phosphorylated intermediate. Numerous attempts have been made to directly detect and isolate such an intermediate using high specific 25. M. Cohn, JBC 230, 369 (1958).

21.

INORGANIC PYROPHOSPHATASES

539

activity s2PP,in a wide variety of conditions (9). No covalently bound has been detected using techniques which have successfully demonstrated phosphoryl enzymes in other systems which include both acidand alkali-labile forms. From the lack of phosphoryl transfer and lack of detectable phosphoryl enzyme it appears that the mechanism may involve concerted nucleophilic attack of water, perhaps assisted by the metal ion. An attractive model for the role of the metal ion, conversion to unsymmetrically charged species which are very rapidly hydrolyzed, appears to have been ruled out by the nonenzymic model experiments of Cooperman ( 2 6 ) . The role of the metal ion may be the key to the problem of the mechanism of this enzyme. Although the reaction is relatively simple compared to many other enzymic reactions, metal ions appear to participate in three different ways; thus, elucidation of the exact catalytic role is unlikely to be straightforward. S2P

111. Other Inorganic Pyrophosphatases

Many nonspecific phosphatases hydrolyze PP, at a rapid rate, but only enzymes relatively specific for hydrolysis of PP, will be considered here. Pyrophosphatases with properties similar to those of the yeast enzyme (neutral or alkaline pH optima, Mg2+or other cations required) are distributed universally. Pyrophosphatases obtained from a variety of bacterial species have been classified according to heat stability and inducibility by PP, (27). Tono and Kornberg have purified the pyrophosphatases from both vegetative cells and spores of Bacillus subtilis (28) and B. megatarium (29).The physicochemical and catalytic properties of the enzymes isolated from the spores are essentially identical to those isolated from vegetative cells. The activity and stability of these enzymes are profoundly enhanced by Mn2+. An inorganic pyrophosphatase which is tightly bound to chromatophores of Rhodospirillum rubrum has been implicated as the reverse of the final reactions in the photosynthetic production of PP, by this organism which can also utilize PP, as energy donor in a phosphorylating electron transport system (SO). 26. B. S. Cooperman, Biochemistry 8, 5005 (1969). 27. B. I. Blumenthal, M. K. Johnson, and E. J. Johnson, Can. J. Microbial. 13, 1695 (1967). 28. H. Tono and A. Kornberg, JBC 242, 2375 (1967). 29. H. Tono and A. Kornberg, J . Bacterial. 93, 1819 (1967). 30. M. Baltscheffsky, H. Baltscheffsky, and L.-V. von Stedingk, Brookhaven Symp. Bbl. 19, 246 (1966).

540

LARRY G . BUTLER

Alkaline pyrophosphatase dependent on Mgz+ was found in every sample examined from a broad spectrum of the plant kingdom (31). Plants which fix CO, by the dicarboxylic acid pathway have characteristic high levels of alkaline pyrophosphatase in their chloroplasts ; presumably this performs the rather specific function of driving the synthesis of phosphoenolpyruvate, the immediate precursor of CO, fixation ( 3 2 ) . Biosynthesis of the maize chloroplast enzyme is controlled by light acting through the phytochrome system (33). Pyrophosphatase from spinach chloroplasts has been partially purified (34, 35). An alkaline pyrophosphatase from rat liver cytoplasm has been partially purified and characterized (24) ; the corresponding enzyme from mice is inhibited by Mg2+-ADP and free PPi, and free Mg2+has been implicated as an allosteric activator ( 2 3 ) . Partial heat inactivation results in loss of the apparent allosteric effects. R a t liver mitochondrial pyrophosphatase, which is inhibited by adenine nucleotides ( 3 6 ) ,appears to be bound to the inside of the inner mitochondrial membrane (37'). This enzyme, after solubilization, has been separated into two fractions which have somewhat different specificity (24, 38). A pyrophosphatase strongly simulated by sulfhydryl reagents (39) has been partially purified from brain tissue (40). The mono-magnesium PP, complex appears to be the true substrate for this enzyme (41). Pynes and Younathan have purified a pyrophosphatase 1800-fold from human erythrocytes (4.2). The properties of this enzyme are strikingly similar to those of the yeast enzyme; the major difference appears to be the more rigid substrate specificity of the erythrocyte enzyme in the presence of Znz+. In general, pyrophosphatases with acidic pH optima are less specific than corresponding enzymes with optimal activity in the neutral or alkaline range and have no requirement for divalent cations (43). Oginsky and Rumbaugh have described an unusual cobalt-dependent 31. B. Nagana, B. Venugopd, and C. E. Sripathi, BJ 60, 224 (1955). 32. S. Simmons and L. Butler, BBA 172, 150 (1969). 33. L. G. Butler and V. Bennett, Plant Physiol. 44, 1285 (1969). 34. A. M. El-Badry and J. A. Bassham, BBA 197, 308 (1970). 35. A. E. Karu and E. N. Moudrianakis, ABB 129, 655 (1969). 36. R. C. Nordlie and H. A. Lardy, BBA 53, 309 (1961). 37. L. Schick and L. G. Butler, J. Cell Biol. 42, 235 (1969). 38. M. Irie, A . Yabuta, T. Negi, and K. Tomita, J . Biochem. ( T o k y o ) 67, 59 (1970). 39. J. J. Gordon, BJ 66, 50 (1957). 40. U. S. Seal and F. Binkley, JBC 228, 193 (1957). 41. E. A. Robbins, M. P. Stulberg, and P. D. Boyer, ABB 54, 215 (1955). 42. G. D. Pynes and E. S. Younathan, JBC 242, 2119 (1967). 43. R. Brightwell and A . L. Tappel, ABB 124, 333 (1968).

21.

INORGANIC PYROPHOSPHATASES

541

bacteria1 pyrophosphatase which has optimal activity near pH 5 and which is rather specific for PPi as substrate (44).The enzyme is strongly activated by low concentrations of histidine, and other amino acids can partially replace histidine.

44.

E. L. Oginsky and H. L. Rumbaugh. J. Bncteriol. 70, 92 (1955).

This Page Intentionally Left Blank

22 Glucose.6.Phosfihatase. Hydrolytic and Synthetic Activities ROBERT C. NORDLIE I. Introduction

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

. . . . . . . . . . . A.Historica1 B. Distribution of Glucose-6-Phosphatase Phosphotransferase C . Relation to Other Enzymes . . . . . . . I1. Molecular Properties . . . . . . . . . . A . Solubilization and Attempted Purification . . . . B. Phospholipids and Glucose-6-Phosphatase . . . . C. Effects of Detergents an$ Detergent-like Effects . . D . Possible Significance of the Membranous Nature of . . . . . . . . Glucose-6-Phosphatase 111. Catalytic Properties A . Methods of Assay . . . . . . . . . B. Reactions Catalyzed C. Thermodynamic Considerations . . . . . . D . Kinetic Studies and Reaction Mechanism . . . . E . Control of Glucose-6-Phosphatase Phosphotransferase . . . . . . . . . . . Activities I V . Metabolic Roles and Regulation, in Vivo . . . . . . A . Control of Enzymic Activities. in Vivo . . . . B . A Final Speculation . . . . . . . . . Appendix . . . . . . . . . . . . . .

. . . .

543 545 547

.

552 553 553 554 556

.

562

.

.

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

565 566

.

567 571 572

. . . .

596 597 599

.

.

592

600

.

1 Introduction

Glucose-6-phosphatase (~-glucose-6-phosphatephosphohydrolase. EC 3.1.3.9) is a unique enzyme in several respects . It is the only principal 543

544

ROBERT C. NORDLIE

enzyme of carbohydrate metabolism associated with the endoplasmic reticulum. And, although traditionally considered an important hydrolytic catalyst, it also possesses more potent synthetic activity than any other hydrolase known. Indeed, as discussed in detail in Sections I11 and IV, the synthetic capacity of the enzyme under certain conditions exceeds that for hydrolysis. The enzyme has been reviewed briefly by Byrne in the preceding edition of “The Enzymes” ( I ) and elsewhere ( 2 ); it has also been reviewed by Manners (S), and by Swanson ( 4 ) and Nordlie and Arion ( 5 ) in two volumes of “Methods in Enzymology.” The roles played by hepatic glucose-6-phosphatase in regulating carbohydrate metabolism have been described in excellent reviews by Cahill et al. (6) and by Ashmore and Weber ( 7 ) . The latter work also contains a comprehensive review of catalytic properties of the phosphohydrolase activity of the enzyme covering studies carried out prior to 1958. Glucose-6-phosphatase, along with a number of other enzymes involved in carbohydrate metabolism, has also been reviewed in Japanese (8). Phosphotransferase activities of the enzyme (see below) have been considered in recent reviews by the present author (9, l o ) ,by Cohn et al. (II), and, along with other allosteric enzymes, by Stadtman ( I d ) . It is the author’s intention in this chapter to emphasize newer findings relating to the hydrolytic activities of the enzyme and to place considerable emphasis on the variety of phosphotransferase activities of this multifunctional catalyst which have been elucidated in the past 6 years.

+

Glucose-6-P He0 -+ Glucose-6-P sugar or polyol R-P glucose -+ R-P H20 -+

+

+

--f

+

+

glucose Pi glucose sugar-P or polyol-P R glucose-6-P R Pi

+ +

+

(1) (2)

(3) (4)

1. W. L. Byme, “The Enzymes,” 2hd ed., Vol. 5, p. 73, 1961. 2. W. L. Byrne, Fructose-l,6-Diphosphatase,Its Role Gluconeogenesis, Symp., univ. Virginia, 1961 p. 89. Am. Inst. Biol. Sci., Washington, D. C., 1964. 3. D. J. Manners, Ciba Found. Symp., Control Glycogen Metab. p. 321 (1964). 4. M. A. Swanson, “Methods in Enzymology,” Vol. 2, p. 541, 1955. 5. R. C. Nordlie and W. J. Arion, “Methods in Enzymology,” Vol. 9, p. 619, 1966. 6. G. F. Cahill, Jr., J. Ashmore, A. E . Renold, and A. B. Hastings, Atm. J. Med. 26, 264 (1959). 7. J. Ashmore and G. Weber, Vitamins Hormones 17, 91 (1959). 8. T. Oda and H. Koide, Saishin l g a k u 16, 2 (1961) ; C A 55, 7515a (1961). 9. R. C. Nordlie, Ann. N . Y . Acad. Sci. 166, 699 (1969). 10. R. C. Nordlie, in “Control of Glycogen Metabolism” (W. J. Whelan, ed.), p. 153. Academic Press, New York, 1968. 11. R. M. Cohn, R. H. Herman, and D. Zakim, A m . J. Clin. Nutr. 22, 1204 (1969). 12. E. R. Stadtman, Advan. Enzymol. 28, 41 (1966).

22. GLUCOSE-6-PHOSPHATASE

545

where R - P = PPi, nucleosidetriphosphate, nucleosidediphosphate, mannose-6-P, fructose-6-P, carbamyl-P, phosphoenolpyruvate, phosphoramidate, or certain other phosphoryl compounds.

A. HISTORICAL 1. Phosphohydrolase Activity Work prior to 1940 by Cori and co-workers (13, 14) and by Ostern, Herbert, and Holmes (15)indicated that blood sugar was produced from hepatic glycogen by the combined action of phosphorylase plus a phosphatase acting on a hexosemonophosphate ester. Because of the presence in their liver extracts of phosphoglucomutase, Cori et al. (14) were unable to establish whether glucose-1-P or glucose-6-P was the actual substrate for this phosphatase. On the basis of studies of the hydrolysis of a variety of phosphate esters by rabbit extracts, Fantl and Rome (16) concluded in 1945 that hepatic tissues contain an enzyme-which of catalyzing the breakthey termed glucose-6-phosphatase-capable down of glucose-6-P to glucose and P i . The enzyme was shown to be optimally active between pH 6 and 7, to be inhibited by phlorizin and Zn2+,and to be relatively unstable to dialysis. I n 1948, Broh-Kahn and Mirsky ( 1 7 ) extended the earlier work of the Cori’s (1.9, 14) and Ostern et al. (15) and established that glycogenolysis involved the production of glucose-1-P which was obligatorily converted to the hexose-6-P ester prior to hydrolysis by phosphatase action, thus confirming the existence of a specific glucose-6-phosphatase system in liver. They further demonstrated the inhibition of hydrolysis of glucose6-P (and not of the interconversion of glucose-1-P to glucose-6-P) by fluoride ion, phlorizin, Zn2+,alloxan, and high concentrations (0.4-2.1 %, w/v) of glucose. I n 1949, both Swanson (18, 19) and de Duve et al. (20, 21) described 13. G. T. Cori and C. F. Cori, Proc. SOC.Ezptl. Biol. Med. 39, 337 (1938). 14. G.T.Cori, C. F. Cori, and G. Schmidt, JBC 129, 629 (1939). 15. P. Ostern, D. Herbert, and E. Holmes, BJ 33, 1858 (1939). 16. P.Fantl and N. M. Rame, Australian J . Exptl. Biol. Med. Sci. 23, 20 (1945). 17. R. H. Broh-Kahn and I. A. Mirsky, A B B 16, 87 (1948). 18. M. A. Swanson, Federation Proc. 8, 258 (1949). 19. M.A. Swanson, JBC 184, 647 (1950). 20. C. de Duve, J. Berthet, H. G. Hers, and L. Dupret, Proc. 1 s t Intern. Congr. Biochem., Cambridge, 1949 Abstr. Commun., p. 403. Cambridge Univ. Press, London and New York, 1949. 21. C. de Duve, J. Berthet, H. G. Hers, and L. Dupret, Bull. SOC.Chim. Biol. 31, 1242 (1949).

546

ROBERT C. NORDLIE

the partial separation from nonspecific acid and alkaline phosphatases of a distinct, rather labile liver enzyme which was capable of hydrolyzing fairly specifically glucose-6-P. Catalytic properties of hydrolase activity of this enzyme, as described by these workers in their initial (19, 21) and succeeding (22-27) reports, as well as by others, are considered in Sections I1 and 111. 2. Phosphotransferase Activities

Hass and Byrne (28-30) and Segal (31) described in 1959 and 1960 the ability of liver microsomal glucose-6-phosphatase to catalyze an exchange reaction involving transphosphorylation between glucose-6-P and 14C-gliicose [reaction (2a) 1. The former workers (SO) demonstrated that Glucose-6-P

+ 14C-glucoseF? 14C-glucose-6-P+ glucose

(24

D-fructose could also function, to a lesser extent, as phosphoryl group acceptor with this system. I n 1960, Rafter (32) first described an activity in mouse liver mitochondrial preparations which was capable of catalyzing the hydrolysis of PP, and the transfer of a phosphoryl group from PPi to glucose to produce glucose-6-P [reaction ( 3 a ) l . This same activity was also observed PPi

+ glucose

glucose-6-P

+ Pi

(34 later by Nordlie and Lardy (33) in rat liver mitochondria1 preparations and by Nordlie and Gehring (34) in rat liver microsomal preparations. Subsequent studies, first reported in consecutive papers in 1964 by Nordlie and Arion (35) and by Stetten ( 3 6 ) , indicated that the PPi-glucose --*

22. C. de Duve and H. Beaufay, Bull. SOC.Chim. Biol. 33, 421 (1951). 23. H. G. Hers and C. de Duve, Bull. SOC.Chim. BWZ. 32, 20 (1950). 24. H. G. Hers, J. Berthet, L. Berthet, and C. de Duve, Bull. SOC.Chim. Biol. 33, 21 (1951). 25. H. Beaufay and C. de Duve, Bull. SOC.Chim. Biol. 36, 1525 (1954). 26. H. Beaufay, H. G. Hers, J. Berthet, and C. de Duve, Bull. SOC.Chim. Biol. 36, 1539 (1954). 27. H. Beaufay and C. de Duve, Bull. SOC.Chim. Biol. 36, 1551 (1954).

28. L. F. Hass and W. L. Byrne, Proc. 4th Intern. Congr. Biochem., Vienna, 1958 Abstr., Vol. 15, p. 39. Pergamon Press, Oxford, 1960. 29. L. F. Hass and W. L. Byrne, Science 131, 991 (1960). 30. L. F. Hass and W. L. Byrne, JACS 82, 947 (1960). 31. H. L. Segal, JACS 81, 4047 (1959). 32. G. W. Rafter, JBC 235, 2475 (1960). 33. R. C. Nordlie and H . A. Lardy, BBA 53, 309 (1961). 34. R. C. Nordlie and A. W. Gehring, Proc. N . Dakota h a d . Sci. 17, 73 (1963). 35. R. C. Nordlie and W. J. Arion, Federation Proc. 23, 534 (1964). 36. M. R. Stetten, Federation Proc. 23, 534 (1964).

22. GLUCOSE-6-PHOSPHATASE

547

phosphotransferase activity and the hydrolysis of PP, were catalyzed by rat hepatic microsomal glucose-6-phosphatase (37, 38). Activity noted previously with mitochondria1 preparations appears to have resulted from contamination of such preparations with small amounts of microsomes which could be removed by repeated washings (39). Further studies from a number of laboratories, described in detail in Sections III,B,l and 2 have confirmed the identity of these activities with classic microsomal glucose-6-phosphatase. Mannose-6-P ( 4 0 ) ,a variety of nucleoside diphosphates and nuileoside triphosphates (41), carbamyl-P and phosphoenolpyruvate (42, 4 3 ) , and phosphoramidate (44) have also been shown to function as phosphoryl group donors, with various degrees of efficiency, with this system, which also has been demonstrated in kidney (&), intestine (46, 47), and pancreas (48) as well as liver.

B. DISTRIBUTION OF GLUCOSE-6-PHOSPHATASE PHOSPHOTRANSFERASE. 1. Phylogenetic and Tissue Distribution

“Glucose-6-phosphatase” has been described as present in a wide variety of tissues and organs from a large number of species of mammals, fishes, amphibians, birds, plants, and microorganisms. Reports of the occurrence of this activity in liver, kidney, small intestine, and pancreas are presented in Tables XII, XIII, XIV, and XV, respectively, in the Appendix. Reports on the occurrence of this activity in other mammalian organs and tissues are considered in Table XVI, while papers describing activity from all other sources-bird, fish, amphibian, plant, and microorganism-are listed in Table XVII. In compiling these tables, the author has attempted to include all references in the literature to “glucose-6-phosphatase.” Studies refer37. R. C. Nordlie and W. J. Arion, JBC 239, 1680 (1964). 38. M. R. Stetten, JBC 239, 3576 (1964). 39. W. J. Anon, Master’s Thesis, University of North Dakota, 1964. 40. W. J. Arion and R. C. Nordlie, JBC 239, 2752 (1964). 41. R. C. Nordlie and W. J. Arion, JBC 240, 2155 (1965). 42. R. C. Nordlie, J. D. Lueck, and T. L. Hanson, Federation Proc. 29, 913 (1970) (abstr.). 43. J. D. Lueck and R. C. Nordlie, BBRC 39, 190 (1970). 44. R. Parvin and R. A. Smith, Biochemistry 8, 1748 (1969). 45. R. C. Nordlie and J . F. Soodsma, JBC 241, 1719 (1966). 46. D. G. Lygre and R. C. Nordlie, Biochemistry 7, 3219 (1968). 47. D. G. Lygre, Doctoral Dissertation, University of North Dakota, 1968. 48. D. B. M. Scott and G. Jones, in “Enzymes and Isoenzymes” (D. Shugar, ed.), Abstr. 364. Academic Press, New York, 1970.

548

ROBERT C. NORDLIE

enced include detection of activity by histochemical techniques as well as by direct biochemical analysis. A word of caution appears to be in order regarding the information in Tables XII-XVII, since in many instances workers have equated the hydrolysis a t acid p H of glucose-6-P with L‘glucose-6-phosphatase activity.” I n certain instances [for example, blood (see references 49-56) 1 , subsequent studies (57-59) have revealed that nonspecific phosphatase activity (alkaline phosphatase in the case of blood) actually was responsible for the noted hydrolyses. I n other instances [see, for example, Thompson’s study (60) of bean cotyledon enzyme and that of Uno (61) with preparations from Aspergillus oryzae] attempts have been made to establish the involvement of an enzyme discrete from nonspecific phosphatase. It appears to the author that the existence of a distinct glucose-6phosphatase is well established for endoplasmic reticulum of liver, kidney, small intestine, and pancreas; but additional studies appear to be required before a similar conclusion may be reached regarding the enzyme from other sources. It should be pointed out, however, that while glucose6-P hydrolysis by enzymes from many sources may not result from specific “glucose-6-phosphatase,” this hydrolysis in these tissues may nonetheless be of metabolic significance. 2. Intracellular Distribution

a. Phosphohydrolase Activity. I n 1951, Hers et al. (24) fractionated tissue homogenates by differential centrifugation technique and studied quite thoroughly the subcellular distribution of glucose-6-P phosphohydrolase activity in liver and kidney. Results of their studies, summarized in Table I, indicated that the enzyme from rat, guinea pig, and rabbit was principally microsomal. A variety of subsequent studies, which have been summarized in tabular form by Ashmore and Weber (7), 49. G. C. Secchi, A. Rezzonico, and N. Gervasini, Enzymologia 33, 134 (1967). 50. S. diBella, G. Richetta, and U. Pichierri, Clin. Chim. Acta 8, 788 (1963). 51. S. Szmigielski, J. Litwin, and B. Zupanska, Arch. Immunol. Therap. Ezptl. 14, 362 (1966) ; B A 48, 13649 (1967). 52. J. Kellen and K. Belaj, Clin. Chim. Acta 6, 595 (1961). 53. S. Fossa, F. Dall’Orso, and C. Marino, Minerva Anestesiol. 32, 325 (1966). 54. G. A. Dosta and Y. M. Ostrovskii, Vopr. Med. Khim. 8, 477 (1962). 55. G. A. Dosta, Byul. Ekserim. Biol. i Med. 55, 59 (1963). 56. H. Koide and T. Oda, Clin. Chim. Acta 4, 554 (1959). 57. M. Foe, Clin. Chim. Acta 17, 13 (1967). 58. K. Zuppinger, Clin. Chim. Acta 6, 759 (1961). 59. R. Wagner and A. Yourke, A B B 54, 174 (1955). 60. J. E. Thompson, Can. J . Biochem. 47, 685 (1969). 61. K. Ono, Master’s Thesis, University of Maryland, 1968.

549

22. GLUCOSE-6-PHOSPHATASE

TABLE I SUBCELLULAR DISTRIBUTION OF G ~ u c o s ~ - 6 -PHOSPHOHYDROLASE~ P ~

Live+

Kidney”

Tissue fraction

Rat

Guinea Pig

Rabbit

Guinea Pig

Rabbit

(%)

(%)

(%I

(%I

(%)

Nuclei plus mitochondria Microsomes Soluble fraction Recovery ( %)

5

5

-

20

20

87 6 98

87 6 98

85 3 106

58 16 94

75 9 104

Summarized from Hers el al. (34). are expressed as percent of homogenate activity per gram liver recovered in each fraction. “Recovery” is total percent of homogenate activity recovered in the various subcellular fractions. Assays were carried out a t pH 6.5 and 38”. Animals had been fasted for 12 hr before use.

* Data

confirmed the fact that this enzyme is present principally in the endoplasmic reticulum of the cell. Although glucose-6-phosphatase activity in the nuclear fraction which has generally been observed may in part result from contamination of nuclear preparations with unruptured cells and debris, activity has also been observed with purified nuclei (62). Further, Kashnig and Kasper (83) have recently identified glucose-6-phosphatase as a component of rat liver nuclear membrane. Small amounts of activity which often have been reported for mitochondrial preparations appear to result from contamination of such preparations with small amounts of microsomes, as indicated above (S7, 39). b. Phosphotransferase Activities. As discussed in detail in Section III,B, studies during the past several years have revealed that liver, kidney, intestinal, and pancreatic glucose-6-phosphatases also catalyze a variety of phosphotransferase reactions [see Eqs. (2) and ( 3 ) ] . Unlike relatively minor phosphotransferase activities exhibited by nonspecific acid and alkaline phosphatases ( 6 4 ), some of these phosphotransferase activities of glucose-6-phosphatase can, under certain conditions, equal or actually exceed the rate of hydrolysis of glucose-6-P (10, 40, 42, 4S, 4.5). Among other types of studies supporting the common identity of 62. C. C. Widnell and J. R. Tata, BJ 92, 313 (1964). 63. D. M. Kashnig and C. B. Kasper, JBC 244, 3786 (1969). 64. R. K. Morton, Dkcussions Faraday Sac. 20, 149 (1955).

550

ROBERT C. NORDLIE

these activities, and of acid inorganic pyrophosphatase activity, with classic microsomal glucose-6-phosphatase has been the demonstration of identical patterns of subcellular distribution of all. Results of studies of the subcellular distribution of rat hepatic activities (38,65), rabbit intestinal activities (46), and human liver activities (66) are summarized briefly in Tables 11-IV. I n all instances identical patterns of distribution of all activities were observed, with the bulk of the activity residing in the microsomal preparations and most of the remaining activity present in the nuclei plus cellular debris fraction. I n interesting recent studies, Pollak and co-workers (67, 68) have observed glucose-6-P phosphohydrolase, acid inorganic pyrophosphatase, and PPi-glucose phosphotransferase activities to be considerably higher in lipid-poor reticulosomes than in microsomal preparations. They hypothesized that reticulosomes, which they prepared by treatment of microsomal preparations from rat and chick livers with ribonuclease and deoxycholate (68), may be the precursors of endoplasmic reticulum, and TABLE I1 SUBCELLULAR DISTRIBUTION OF INORGANIC PYROPHOSPHATASE, PPi-G~ucos~ PHOSPHOTRANSFEKASE, AND GLUCOSE-~-P PHOSPHOHYDROLASE ACTIVITIES O F RAT LIVERMICROSOMAL GLUCOSE-6-PHOSPHATASeO ~~~

~~

Activityb Tissue fraction Nuclei plus cellular debris Heavy mitochondria Light mitochondria Microsomes Cytosol

Inorganic pyrophosphatase

PPi-glucose phosphotransferase

Gluco~e-6-P phosphohydrolase

(%I

(7%)

17.9

18.5

17.6

3.2 71.8

1.7 4.3 75.5

3.9 6.8 71.7

C

C

C

(%I

7.1

Calculated from Stetten (38).Assays were carried out a t pH 5.4with 0.08 M phosphate substrate, 0.4 M glucose (phosphotransferase), and 0.0497,Triton X-100 present. Data are expressed as lOOX activity in indicated fraction/sum of activities in all fractions. c Cytosol contained less than 1% of total activity in all instances. 65. M. R. Stetten and D. Rounbehler, JBC 243, 1823 (1968). 66. R. C. Nordlie, unpublished observations (1970). 67. J. K. Pollak, R. Malor, M. Morton, and K. A. Ward, in “Autonomy and Biogenesis of Mitochondria and Chloroplasts.” North-Holland Publ., Amsterdam (in press). 68. K.A. Ward and J. K. Pollak, BJ 114, 41 (1969).

551

22. QLUCOSE-6-PHOSPHATASE

TABLE I11 SUBCELLULAR DISTRIBUTION OF RABBIT INTESTINAL GLUCOSE-~-PHOSPHATASE AND ASSOCIATED INORGANIC PYROPHOBPHATABE AND PHOGPHOTRANSFERABE ACTIVITIEBO Specific activityb X 100 Tissue fraction Nuclei plus cellular debris Mitochondria1 Microsomal Soluble

Glucose-6-P phosphohydrolase

PPi-glucose CDP-glucose Inorganic phosphotransphosphopyrophosphatase ferase transferase

2.2

3.3

0.89

0.25

1.2 6.6 0

2.3 5.9 0.4

0.36 2.3

0.14 0.74 0

0

Data are from references 46 and 47. Assay mixtures, pH 6.0, contained 3.3 mM phosphate substrate and 180 mM D-glucose (phosphotransferaae). Specific activity = pmoles substrate hydrolyzed (hydrolase) or pmolea glucose-6-P synthesized (phosphotransferase) per min per mg protein a t 30". 0

have demonstrated membrane formation by recombination of reticulosomes with microsomal lipid extracts. A decrease in enzymic activity concomitant with the latter process was also observed. The possible role of phospholipids in glucose-6-phosphatase phosphotransferase action is further considered in Section I1,B. TABLE I V SUBCELLULAR DISTRIBUTION OF G ~ u c 0 s ~ d -PHOSPHOHYDROLASE, P INORQANIC PYROPHOSPHATASE, AND PPi-GLUCOSE PHOSPHOTRANSFERASE ACTIVITIESOF HUMANLIVER^ Activityb Tissue fraction Nuclei plus cellular debris Mitochondrial Microsomal Cytosol

Inorganic pyrophosphataae

PPi-glucose phosphotransferase

Glucose-6-P phosphohydrolase

(%)

(%I

(%)

20.1

37.7

36.1

16.4 54.1 9.4

0 62.3 0

6.5 75.4 0

From Nordlie (66). Relative activity is expressed as in Table 11. Assays were a t pH 6.0 with 10 mM phosphate substrates and 180 mh4 glucose (phosphotransferase). Deoxycholate was added to tissue preparations, to 0.27'0 (w/v), before assay. b

552

ROBERT C. NORDLIE

C. RELATION rn OTHER ENZYMES Glucose-6-P occupies a central, crossroad position in carbohydrate metabolism, as indicated in Fig. 1. This hexosephosphate ester may be formed by glucose phosphorylation, by gluconeogenesis, and, through the intermediation of phosphoglucomutase, by glycogenolysis. The relative normal rates of disposition of glucose-6-P through glycolysis, glycogenolysis, direct oxidation by glucose-6-P dehydrogenase, and hydrolysis by glucose-6-P phosphohydrolase, as demonstrated by Ashmore e t al. (69),are indicated in the figure. The last process appears predominant among these alternate pathways for glucose-6-P utilization in liver. Physiological roles for both phosphohydrolase and phosphotransferase activities of the enzyme are considered briefly in Section IV.

(or PPi, etc.)

GLC

Gluconeogenesis

Glycolysis

(251 I

PYRUVATE

FIG.1. Some interrelationships between glucose-6-phosphatase and other enzymes of carbohydrate metabolism [from Nordlie (10) ; copyright (1968), Academic Press, Inc. Reproduced by permission1 . Numbers in parentheses indicate the relative disposition per 100 molecules of glucose phosphorylated of glucose-6-P via four alternate metabolic pathways, according to Ashmore et al. (69). Further details are given in the text. 69. J. Ashmore, G. F. Cahill, Jr., A. B. Hastings, and S. Zottu, JBC 224, 225 (1957).

22. GLUCOSE-6-PHOSPHATASE

553

II. Molecular Properties

A. SOLCBILIZATION AND ATTEMPTED PURIFICATION Glucose-6-phosphatase is either a part of, or strongly attached to, the lipoprotein membrane of the endoplasmic reticulum ( 7 0 ) .Because of the enzyme’s particulate nature, attempts at purification have, a t best, thus far met with only indifferent success (5). Techniques which have been successful in the solubilization and purification of other particulate enzymes [for example, butanol extraction and adsorption on and elution from calcium phosphate gel with nonspecific acid phosphatase (see reference 64)] have simply destroyed glucose-6-phosphatase activity (26, 27, 37, 71-73). Solubilization of the enzyme from liver microsomal preparations of a number of species of mammals has been reported involving treatments with deoxycholate (27, 37), digitonin (71,72, 7 4 ) , digitonin plus deoxycholate (75, 76), digitonin, deoxycholate, and glycerol (77), proteolysis with trypsin or papain in combination with digitonin (78, 79), Lubrol W, a nonionic detergent ( 7 3 ) , and mechanical disruption in the French press ( 8 0 ) . It has been the author’s experience (37) that, while detergent-treated microsomes are dispersed such that the enzyme may be considered “solubilized,” such preparations slowly reaggregate with time in the absence of detergent and regain their particulate nature, presumably because of their high content of lipid. Moderate enrichments of activity have been reported following fractional precipitation and differential centrifugation of microsomal preparations treated with detergents (37, 70, 72, 7 8 ) or proteolytic enzymes (78), or fractional precipitation of detergent-dispersed microsomal preparations with acetone in the presence of Mg2+(46, 8 1 ) . The isolation and partial purification of soluble glucose-6-phosphatase L. Ernster, P. Siekevitr, and G. E. Palade, J . Cell Biol. 15, 541 (1962). H. L. Segal, M. E. Washko, and C. W. Lee, Science 128, 251 (1958). H. L. Segal and M. E. Washko, JBC 234, 1937 (1959). C. Carruthers and A. Baumler, ABB 99, 458 (1962). R. G. Langdon and D. R. Weakly, Federation Proc. 16, 208 (1957). M. Gorlich and E. Heise, Nature 194, 376 (1962). M. Gijrlich and E. Heise, 2.Naturforsch. 17b, 465 (1962). C. Stahlmann, L. Richter, C. Dorow, E. Schutte, W. Schreiber, and T. Gunther, 2. Physiol. Chem. 348, 633 (1967). 78. M. Gorlich and E. Heise, Nature 197, 698 (1963). 79. E. Heise and M. Gorlich, Nature 197, 1311 (1963). 80. R. C. Nordlie, J. R. Gilsdorf, R. N. Horne, and R. J. Paur, BBA 158, 157 70. 71. 72. 73. 74. 75. 76. 77.

(1968). 81. M. C. Ganoza, Doctoral Dissertation, Duke University, 1964.

554

ROBERT C . NORDLIE

from pig liver microsomes has been claimed by Stahlmann et al. (77), who also have described the chemical composition and physical properties of their partially purified preparations. The enzyme, solubilized by treatment with glycerol, digitonin, and deoxycholate, and partially purified by chromatography on Sephadex G-100, sedimented as a single peak a t pH 6.8 in the analytical centrifuge. These authors describe their solubilized enzyme as a monomer with a molecular weight of 325,000 and one active site per molecule. They indicate that freeze-dried, dialyzed preparations contained, by weight, 53% residual digitonin, 26% lipids (approximately half of which were phosphatides) , 19% protein, 4% water, and 0.1% cholesterol. Since bands for protein, lipid, and enzymic activity, obtained after agar gel electrophoresis, were superimposable, Stahlmann et al. (77) concluded that the enzyme is a lipoprotein. These authors indicated a recovery of 92% of original activity in their most highly purified preparations. I n view of the fact that the specific activity of this purified fraction is only 3.8 times that of microsomal fraction, one is left with the conclusion either that glucose-6-phosphatase constitutes an extremely large portion of total microsomal protein (approximately 25% ) or that Stahlmann et al. (77) were working with very crude enzyme preparations. The latter conclusion is supported by the observations of Ernster et al. (70), who obtained a six- to twentyfold enrichment in glucose-6-phosphatase activity following treatment of microsomal preparations with deoxycholate and centrifugation a t 105,000 x g for 2 hr. The resulting phospholipid-rich, “fluffy” upper layer (“M” fraction) had a specific activity more than six times that of original microsomes and more than 20 times that of the pellet resulting from such treatment. It is the present author’s opinion that a significant purification of glucose-6phosphatase is yet to be accomplished.

B.

PHOSPHOLIPIDS AND

GLUCOSE-6-PHOSPHATASE

In 1954, Beaufay and de Duve (27) first suggested a relationship between microsomal phospholipid and glucose-6-phosphatase. They observed a loss of enzymic activity from phospholipid-rich microsomal preparations concomitant with extraction with such organic solvents as butanol or treatment with lecithinase. Various studies were carried out to demonstrate that the latter effect was not produced through inhibition of enzymic activity by accumulated products of the hydrolysis of phospholipids. On the basis of their observations that deoxycholate treatment labilized microsomes to phospholipase action, they concluded that . . . the detergent did not exert its primary effect on the dissociation of phospholipids from microsomal protein, but that it probably disrupted

22. GLUCOSE-6-PHOSPHATASE

555

the association between the lipids and lipoproteins of microsomes. . . . l l Bcaufay and de Duve (27) suggested that one function of membrane lipids was to stabilize the particular conformation of the protein which is required for catalytic activity, a view currently held by the present author in light of the results of work recently carried out in his laboratory (82, 83). An excellent review of this earlier work on the involvement of phospholipids with glucose-6-phosphatase is given in a doctoral dissertation by Ganoza (81). These original studies on the possible roles of phospholipids in glucose-6-P phosphohydrolase activity have been extended by Byrne and co-workers (81, 84-87), and comparative studies of the effects of phospholipase treatment and phospholipid supplementation on both phosphohydrolase and phosphotransferase activities of the rat-liver enzyme have been carried out by Snoke and Nordlie (82, 83). While Beaufay and de Duve (27) were unable to restore glucose-6phosphatase activity by phospholipid supplementation of lecithinasetreated microsomes, Duttera et al. (87), using microsomal preparations from fasted rats, have demonstrated (a) a progressive loss of glucose-6-P phosphohydrolase activity correlative with phospholipase C-catalyzed release of acid-soluble phosphate (phosphoryl choline) from microsomes, (b) lack of inhibition of glucose-6-phosphatase activity by hydrolysis products of phospholipase C action, and (c) restoration of glucose-6-P phosphohydrolase activity to original (or, indeed, supraoriginal) levels by supplementation of phopholipase C-treated preparations with various phospholipids. Snoke and Nordlie (82, 83) have extended these studies of Duttera et al. (87) to include PPi-glucose phosphotransferase as well as glucose6-P phosphohydrolase activity of the enzyme and have also noted essentially parallel progressive losses of both activities concomitant with release of acid-soluble phosphate resulting from phospholipase C action. A partial (fed animals) or total (fasted rats) restoration of both activities was effected by phospholipid supplementation of phospholipase-treated preparations. However, as indicated by detailed studies of catalytic properties of the various preparations, the enzyme was not restored to its 82. R.E.Snoke and R. C. Nordlie, Abstr. 166th Natl. ACS Meeting, Atlantic City Abstr. Biol. 102 (1968). 83. R.E.Snoke, Doctoral Dissertation, University of North Dakota, 1970. 84. M.C.Ganosa and W. L. Byrne, Federation Proc. 22, 535 (1963). 85. M.C.Ganoza and W. L. Byrne, Abstr. 146th Natl. ACS Meeting, N e w York Abstr. 85-C (1963). 86. W.L. Byrne and S. M. Duttera, J . Am. Oil Chemists’ Sac. 44, 363A (1967). 87. S. M. Duttera, W. L. Byrne, and M. C. Ganoza, JBC 243, 2216 (1988).

556

ROBERT C. NORDLIE

initial form following phospholipase treatment and lipid supplementation. The former treatment produced a significant decrease in K, values for all substrates as well as in V,,, values. The addition of phospholipids to phospholipase C-treated preparations decreased the affinity of the enzyme for PP, and glucose-6-P and increased V,,, values a t pH 6.0. Most dramatically, the last treatment shifted the pH optimum of phosphotransferase activity toward neutrality in much the same manner as supplemental detergents (80, 88, 89),cortisone administered in vivo ( 8 0 ) , or mechanical disruption of microsomes (80). Clearly, the “regenerated” enzyme differs catalytically from that of untreated microsomes. The experimental observation cited would appear to be consistent with two catalytically different forms of the enzyme, characterized most strikingly by differences in pH optima for phosphotransferase activity and interconvertible as a result of conformational alterations induced through changes in lipid-protein interactions which in turn may be modified by phospholipase-phospholipid treatments (82, 83), glucocorticoid therapy (80), detergent supplementation (80, 88, 89),or mechanical stress (80). Consistent, also, with the idea of intimate interrelationships between glucose-6-phosphatase activity and microsomal phospholipids is the observation of Ernster e t al. (70) that both phospholipids and enzymic activity are found together in markedly elevated amounts in the loose sediment obtained following centrifugation of deoxycholate-treated microsomes. I n marked contrast with this observation, however, is the recent finding of Pollak et al. (67, 68) that lipid-depleted reticulosomes (see Section I,B,2,b) are considerably richer (specific activities up to 20 times greater) in glucose-6-phosphatase and associated phosphohydrolase and phosphotransferase activities than are microsomes from which the reticulosomes are prepared, and that supplementation of such preparations with microsomal lipid extracts leads to a diminution of enzymic activities. Clearly, the precise details of the interrelationships of phospholipids and glucose-6-phosphatase phosphotransferase are yet to be worked out. The potential significance for differential control of the various hydrolytic and synthetic activities of the enzyme through lipidinduced or lipid-mediated changes is, however, already quite obvious. OF DETERGENTS AND DETERGENT-LIKE EFFECTS C. EFFECTS

As might reasonably be expected on the basis of the intimate relationships between glucose-6-phosphatase and phospholipids discussed above, 88. R. C. Nordlie, T. L. Hanson, and P. T. Johns, JBC 242, 4144 (1967). 89. J. F. Soodsma and R. C. Nordlie, BBA 191, 636 (1969).

22. GLUCOSE-6-PHOSPHATASE

557

activities of the enzyme are influenced, in some instances quite markedly, by a variety of detergents and detergent-like treatments. The latter include the action of ammonium hydroxide a t pH 9.8 (90-92), mechanical disruption (75), and glucocorticoids acting in vivo (80, 93-95). 1. Direct Eflects of Detergents

Beaufay and de Duve (27) first demonstrated that deoxycholate exerts a biphasic effect on glucose-6-phosphatase activity of microsomal preparations, activating a t relatively low concentrations (approximately 1 g/liter) and inhibiting a t higher concentrations. They also showed that deoxycholate treatment labilized the enzyme to thermal inactivation. Generally similar effects of deoxycholate have been observed more recently by Ashmore and Nesbett (96), Carruthers and Baumler (73), Segal and Washko (72) , Nordlie et al. (93) , and Snoke and Nordlie (97). In comparative studies (73, 97) , stimulations of enzymic activities have been noted with certain anionic (deoxycholate and cholate) , nonionic (Triton X-100 and Lubrol W) , and cationic (cetyltrimethylammonium bromide) detergents, while other such compounds in each group (the anionic detergents sodium lauryl sulfate, sodium cetyl sulfonate, sodium dodecylbenzene sulfonate ; the nonionic detergents Tween 20 and Tween 80 ; and the cationic detergents cetavlon, laurylamine, and cetyldimethylbenzylammonium chloride) inhibited and did not activate appreciably a t any concentration tested. Phosphotransferase activity of the enzyme has been found to respond much more extensively than does phosphohydrolase activity to detergent treatment (see, for example, references 86, 89, 92). This observation led us to investigate in detail alterations in catalytic properties of both phosphotransferase and phosphohydrolase activities produced by detergent treatment. Deoxycholate supplementation has been found to shift the pH optimum of PP,-glucose phosphotransferase from approximately 4.5 noted with fresh microsomal suspensions to approximately pH 5.7 with optimal (0.2076, w/v) deoxycholate concentrations added to microsomes (see Fig. 2 and references 80 and 98). Preparations obtained by 90. M. R. Stetten and F. F. Burnett, BBA 139, 138 (1967). 91. M. R. Stetten and F. F. Burnett, BBA 128, 344 (1966). 92. M. R. Stetten, S. Malamed, and M. Federman, BBA 193, 260 (1969). 93. R. C. Nordlie, W. J. Arion, and E. A. Glende, Jr., JBC 240, 3479 (1965). 94. W. J. Arion and R. C. Nordlie, JBC 242, 2207 (1967). 95. R. C. Nordlie and R. E. Snoke, BBA 148, 222 (1967). 96. J. Ashmore and F. B. Nesbett, Proc. SOC.E xpt l . Biol. Med. 89, 78 (1955). 97. R. E. Snoke and R. C. Nordlie, BBA 139, 190 (1967). 98. R. C. Nordlie, W. J. Arion, T. L. Hanson, J. R. Gilsdorf, and R. N. Horne, JBC 243, 1140 (1968).

ROBERT C. NORDLIE

PH

PH

Frc. 2. Activity-pH profiles of liver PPI-glucose phosphotransferase activity assayed with ( A ) untreated microsomes and (B) microsomal preparations supplemented to 0.270, w/v, with deoxycholate. Preparations are from normal, diabetic, or cortisone-treated rats, as indicated. [Modified from Nordlie et al. (80). Copyright (19681, Elsevier Publishing Co. Reproduced by permission.1

fractional ammonium sulfate precipitation of deoxycholate-dispersed microsomal preparations also exhibited this latter pH optimum (see references 37, 41, and 4 5 ) . The pH optimum for inorganic pyrophosphatase activity likewise was shifted toward neutrality by detergent treatment of microsomes, as was, to a lesser extent, that of glucose-6-P phosphohydrolase activity (98). Significant reductions in K,,, values for all phosphate substrates, and to a lesser extent K , for glucose, were also noted following deoxycholate supplementation (93, 98, 99), Detailed kinetic studies (89) on the effects of the cationic detergent cetyltrimethylammonium bromide (cetrimide) on rat kidney microsomal glucose-6-phosphatase phosphotransferase, in which effects of varied concentrations of the detergent on pH optima, v values, V,,,, values, and K , values were assessed, have revealed that this surface-active agent, too, effects shifts in pH optima and alters various kinetic parameters as well. Based on observed polyphasic effects of various concentrations (0.05-0.30%, w/v) of the detergent on v values, together with decreases in K , values for phosphate substrates resulting from all concentrations of detergent tested, and decreases of V,,, values effected by the higher concentrations of cetrimide (0.20 and 0.300/0,w/v) , it has been concluded that the detergent functions both as a classic “coupling” activator (100) 99. T. L. Hanson and R. C . Nordlie, BBA 198, 66 (1970).

22.

GLUCOSE-6-PHOSPHATASE

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a t lower concentrations and as a noncompetitive inhibitor a t higher concentrations (89). Interestingly, the lowest K , values for PP, (0.6 mM, see reference 89) and for carbamyl-P (1.6 mM; references 42, @), functioning as phosphoryl donors with the enzyme, have been noted with cetrimide-supplemented microsomes. Palmityl-CoA (88) and other long-chain fatty acyl-CoA esters (9, 10) exert rather dramatic concentration and pH-dependent activity discriminating effects on phosphotransferase and phosphohydrolase activities of liver and intestinal (46’) enzyme. Both inhibitions and activations were observed in the range 4-40 pM fatty acyl-CoA. For example, 40 pM palmityl-CoA stimulated PP,-glucose phosphotransferase activity to 350% of control values although increasing glucose-6-P phosphohydrolase less than lo%, while 4.1 pM palmityl-CoA inhibited the two activities 50 a,nd 17% (88),respectively. Supplemental long chain fatty acyl-CoA compounds also shifted the pH optimum of phosphotransferase (but not glucose-6-P phosphohydrolase) even more markedly toward neutrality than did deoxycholate. A pH optimum as high as p H 6.0 or 6.5 has been observed in the presence of 40 pit4 palmityl-CoA, for example, and quite significant amounts of activity are demonstrable even a t pH 7.0 in the presence of this (88) and other (9, 10) acyl-CoA esters. These observations suggest the possibility that long-chain acyl-CoA esters, which accumulate in liver in diabetes, may serve under these conditions as a mechanism for “turning on” phosphotransferase activity which in turn may serve as a replacement for insulin-dependent glucokinase in the diabetic animal (9, 10, 88). Concomitant depression of glucose-6-P phosphohydrolase activity by these same acyl-CoA esters likewise may serve as part of a mechanism for placing an upper limit on the elevated rate of net glucose release from gluconeogenic tissues in diabetes (9, 10, 88) (see also Sections II,C,3 and IV). 2 . Detergent-like Effects

a . N H 4 0 H at High p H . Stetten and co-workers (90-92) have demonstrated that in a fashion similar to activation by detergents the exposure of rat liver microsomes to N H 4 0 H a t pH 9.5-9.8 for 30 min a t 30”,or for many hours a t 0”, also quite significantly activates both hydrolytic and synthetic activities of such preparations, although the enzyme is not solubilized by such treatment. The effect is especially pronounced with the latter activity. A decrease in K , values for phosphate substrates is 100. J. S. Friedenwald and G. D. Maengwyn-Davies, in “The Mechanism of Enzyme Action” (W. D. McElroy and B. Glass, eds.), p. 180. Johns Hopkins Press, Baltimore, Maryland, 1954.

560

ROBERT C. NORDLIE

effected by such treatments, as is a shift toward neutrality in the pH optimum of the phosphotransferase activity. Stetten and Burnett (90) suggested that high pH treatment may be superior, for practical purposes, to activation by detergents, since enzyme activated in the former manner is not labilized to thermal denaturation. Activation by ammonium hydroxide a t high pH is suggested to be accomplished through conformational changes in the microsomal membrane (92).

b. Mechanical Disruption. Nordlie and co-workers (80) have found that passage of liver microsomes through a French press under 5000 psi pressure effects a solubilization of approximately 35% of total microsoma1 glucose-6-phosphatase phosphotransferase and that the pH optimum of the phosphotransferase activity of the solubilized preparation (pH 5.5) lies considerably nearer neutrality than does that of unsolubilized microsomal enzyme (pH 4.5) which sediments during centrifugation a t 3 x lo8 g-min.

c. Effects of Glucocorticoids in Vivo. As discussed briefly in Section IV,A below, the administration of glucocorticoids to rats appears to increase glucose-6-phosphatase and associated phosphotransferase activities principally by activating existing enzyme rather than by inducing synthesis of new enzyme protein (80, 93, 94). Pyrophosphate-glucose phosphotransferase activity from liver of both normal control and cortisonetreated rats displayed maximal activity a t pH 4.5 in the absence of detergent (Fig. 2 ) . However, a definite shoulder appeared between pH 5.5 and 6.5 in preparations from glucocorticoid-treated animals. On supplementation of preparations with deoxycholate to 0.2% (w/v) , differences in activity levels for control and cortisone-treated animals disappeared, and the p H optimum was shifted to pH 6.0, with a shoulder remaining a t pH 4.5-5 (80). I n addition, the extent of stimulation by deoxycholate of enzymic activity was much less in glucocorticoid-treated rats than in normal animals (95). These observations suggest that glucocorticoids, functioning in vivo, produced effects resembling in some ways those of detergent supplementation or mechanical disruption in altering the enzyme with respect to pH optima (80). I n contrast to detergent supplementation, however, no changes were noted in K , values resulting from glucocorticoid therapy (72, 93). 3. Modifying Effects of Detergents on the Action of Inhibitors, in Vitro The observations cited above strongly suggest intimate interrelationships between structural features and catalytic behavior of glucose-6phosphatase phosphotransferase. Such interrelationships are even more strongly supported by interesting recent observations on the modifying

22. GLUCOSE-6-PHOSPHATASE

561

effects of certain natural and synthetic detergents on the action of some inhibitors, and of certain in vivo treatments, on phosphohydrolase and phosphotransferase activities of liver and kidney microsomal glucose-6phosphatase. Cetyltrimethylammonium bromide, a cationic detergent which by itself activates both phosphotransferase and phosphohydrolase activities of glucose-6-phosphatase, rather markedly potentiates inhibition by phlorizin of the former activity while a t the same concentrations (0.05-0.30%, w/v) significantly ameliorating inhibition by phlorizin of the latter activity (101, 102). Values of K i for phlorizin, inhibiting noncompetitively vs. glucose-6-P, were increased in the presence of cetrimide, while those determined for the compound inhibiting similarly with respect to PPI or glucose in the phosphotransferase reaction were decreased ( 1 0 2 ) . While neither cetrimide nor phlorizin is a physiological compound in mammals, these observations do point up the fact that hydrolase and phosphotransferase activities of the enzyme are sensitive to differential control by common factors. The effects observed with long-chain fatty acyl-CoA esters (9, 10, 88),as described above, also fit into a similar general category of agents capable of differential regulation of activities of this multifunctional enzyme and in addition may be regarded as “physiological.” Cetrimide, as well as other detergents including lysolecithin and palmityl-CoA, also have been found to potentiate quite significantly the inhibition, by Pi (103,104) and a variety of nucleosidetriphosphates and nucleosidediphosphates (10s), of glucose-6-P phosphohydrolase activity. For example, a t p H 7.5, the Ki value for ATP, a competitive inhibitor of glucose-6-P phosphohydrolase activity, was reduced from a value of approximately 20 mM observed with untreated microsomes to 2 mM with microsomes supplemented with certrimide to 0.1%, w/v, prior to assay; net inhibition by 6 mM ATP of the hydrolysis of 0.13 mM glucose-6-P was concomitantly increased from 22% to 75%. The potentiation by these detergents of inhibition by Pi of PPi-glucose phosphotransferase activity was much less extensive than that with phosphohydrolase activity (104). Interesting studies remain to be carried out relating to mechanisms by which these effects are produced. However, it appears clear even a t this preliminary stage that conformational features, quite probably 101. J. F.Soodsma, B. Legler, and R. C. Nordlie, JBC 242, 1955 (1967). 102. D.G.Lygre and R. C. Nordlie, BBA 185, 360 (1969). 103. R. C. Nordlie, T. L. Hanson, P. T. Johns, and D. G. Lygre, Proc. Natl. Acad. Sci. U. S. 60, 590 (1968). 104. A. L. Vianna and R. C. Nordlie, JBC 244, 4027 (1969).

562

ROBERT C. NORDLIE

relating a t least in part to the lipid-protein nature of the enzyme, may be intimately related to the selective control of synthetic and hydrolytic capabilities of the multifunctional catalyst. SIGNIFICANCE OF THE MEMBRANOUS NATURE D. POSSIBLE O F GLUCOSE-6-PHOSPHATASE While occupying a strategic, crossroad metabolic position (Fig. l ) , glucose-6-phosphatase is unique among enzymes involved in carbohydrate metabolism with respect to its intracellular location. The presence of this enzyme as a part of the membrane of the endoplasmic reticulum, as well as its hydrolytic function, and its presence in tissues which all release glucose into the bloodstream-liver, kidney, and intestine-have prompted Ernster et al. (70) to suggest that the phosphohydrolase activity of the enzyme may be involved in transport of glucose between the intracellular compartment (cytoplasmic matrix) and canalicular system of endoplasmic reticulum which is continuous with the extracellular medium. Similarly, de Duve (105) has suggested that glucose6-phosphatase may be oriented in the membrane of the endoplasmic reticulum in such a manner as to permit the direct release of hydrolysis products to the extracellular compartment. On the basis of the multifunctional nature of the enzyme and the observation that PPi-glucose phosphotransferase activity of the enzyme is much more extensively activated by detergents and high pH treatment than is phosphohydrolase activity, Stetten and Burnett (90) have suggested that “. . . the sites for binding of glucose are chiefly located on the membrane surface within the endoplasmic reticulum lumen with the enzyme oriented in such a way as to permit synthesis from PPi and glucose to channel the glucose-6-P formed into the true intracellular compartment.” Requisite to this conclusion is the hypothesis of these authors that the enzyme “. . . is so oriented within the endoplasmic membrane that the active binding sites for glucose-6-P are predominantly on the outer surface of the microsomes that is readily accessible to the cytoplasm of the cell, while the sites for glucose are largely on that inner surface which corresponds to the space between the ergastoplasmic membranes and are, in effect, outside of the cell.” (90) Such an hypothesized functional role for the enzyme is extremely attractive. However, the basic mechanism of action of glucose-6-phos105. C. de Duve, Ciba Found. Sump., Enzymes Drug Action, p. 505 (1962).

22. GLUCOSE-6-PHOSPHATASE

563

phatase phosphotransferase, involving a single binding site for free glucose and for the glucose moiety of glucose-6-P, based on kinetic studies of Hass and Byrne (SO), Segal (31), and Arion and Nordlie (4O),appears on first consideration to be inconsistent with this hypothesis. Segal (31) has pointed out that his kinetic observations (since confirmed and extended in the author’s laboratory, see reference @) on the nature of glucose inhibition of glucose-6-P hydrolysis are not compatible with a reaction mechanism involving more than a single binding site for glucose. Thcse same arguments hold for kinetic data obtained by Arion and Nordlie (40) with glucose-6-P phosphohydrolase, PPi-glucose phosphotransferase, and mannose-6-P: glucose phosphotransferase activities. It is the author’s feeling that the “two-site” hypothesis relating to a physiological role for the enzyme must be rationalized with the kinetics of the system before the former can become generally acceptable. I n this regard, however, it may be pointed out that the kinetic studies cited above were, of necessity, carried out with activated or partially purified preparations. Conceivably, the enzyme may behave catalytically somewhat differently in its natural state and environment within the cell. For example, enzyme molecules exposed to the internal milieu of the cell may function principally as a phosphohydrolase while those near the canalicular lumen of the endoplasmic reticulum, and hence more directly under the influence of the extracellular environment, may act principally as phosphotransferase. The differential effects of long-chain fatty acyl-CoA esters (9, 10, 88), and of phlorizin in combination with cetrimide (89)’mentioned above, indicate that a variety of factors can discriminantly turn on or turn off hydrolytic and synthetic activities of the enzyme. A suggested scheme incorporating these concepts of the selective participation of hydrolytic and synthetic activities of the enzyme in cellular glucose release and uptake is presented in Fig. 3. Also, it should be pointed out in this connection that the reaction mechanism (see Fig. 4 and Section II11D15,a)postulated on the basis of kinetic observations may be an oversimplification of the actual situation. For example, the possibility exists that separate phosphoryl-enzyme intermediates may be formed with various phosphate substrates (10). If such intermediates were interconvertible with a single phosphorylenzyme complex which was common to reactions involving all such substrates, and if the rates of these interconversions were very rapid relative to the other steps in the catalytic process, the system would be kinetically indistinguishable from that depicted in Fig. 4. Further work relating to the mechanism of reaction, and the experimental assessment of the possible involvement of this multifunctional catalyst in intra- and extracellular glucose transport, would appear to be in order.

564

ROBERT C. NORDLIE

Canalicula of endoplasrnic reticulum (continuous with extracellular medium)

L

IntroceUular Compartment

Z

GLC-6-P

FIG. 3. Proposed mechanism involving hydrolytic and synthetic activities of glucose-6-phosphatase in the transport of glucose between intracellular and extracellular compartments. The shaded area represents the cross-sectional view of endoplasmic reticulum. E’ and E” are modified forms of glucose6phosphatase displaying principally phosphohydrolase and principally phosphotransferase activities, respectively. Differential influences of the intra- and extracellular milieu are postulated to maintain molecules of the enzyme selectively as E or E”. Additional details are given in Section II,D.

E-P-R

E- P- GLUCOSE

REACTION 3

E-P

FIG.4. Scheme depicting the order of interactions of enzyme with substrates. E denotes enzyme ; R-P represents PP,, carbamyl-P, nucleosidetriphosphate, nucleosidediphosphate, or mannoseb-P. [Modified from Arion and Nordlie (40).I

22.

565

GLUCOSE-6-PHOSPHATASE

111. Catalytic Properties

Glucose-6-phosphatase from rat liver microsomes has been investigated most extensively, although more recently catalytic properties of both hydrolytic and synthetic activities of the kidney and intestinal enzymes, and to a lesser extent that of pancreas, have also become the subjects of study. Although, in our hands, subtle differences have occasionally been noted among activities from liver, kidney, and intestine, catalytic properties of the enzyme from the various sources in general are strikingly similar, and the discussion of catalytic properties to follow holds generally for enzyme from the various sources. It should, however, be pointed out in this regard that (a) James (106) has suggested that differences do exist between enzyme from small intestine and that from liver or kidney, and (b) Lygre and Nordlie (46, 47) noted that under identical conditions the ratio of phosphotransferase to phosphohydrolase activity of the kidney enzyme was approximately double that from liver or small intestine (see Table V ) . It is the author's feeling a t present that these apparent differences most likely reflect variations in the state of the enzyme as it is obtained from the different tissues rather than TABLE V COMPARISON OF LEVELSOF PHOSPHOTRANSFERASE AND PHOSPHOHYDROLASE ACTIVITIESO F GLUCOS~-6-PHOSPHATASEI N LIVER,KIDNEY,AND SMALLINTESTINE OF RABBIT" Enzymic activity

Enzyme source Liver Kidney Small inteatine

Glucose-6-P phosphohydrolase

Inorganic pyrophosphatase

PPi-glucose phosphotransferase

CDP-glucose phosphotransferase

7.8 8.0 6.6

8 . 7 (1.1) 7.2 (0.90) 5 . 7 (0.86)

2 . 6 (0.33) 5 . 6 (0.70) 2 . 1 (0.32)

0.6 (0.08) 1 . 7 (0.21) 0 . 7 (0.11)

5Data are from references 46 and 4'7. Assay mixtures, p H 6.0, contained 40 mM sodium cacodylate buffer, 3.3 m M phosphate substrate, and 180 mM D-glucose (phosphotransferase) . * Enzymic activity values without parentheses are expressed as pmoles of substrate hydrolyzed (hydrolase) or pmoles of glucose-6-P formed (transferase) per min per mg protein a t 30". Values in parentheses represent the ratios of indicated activity/glucose6-P phosphohydrolase activity.

106. J. James, Doctoral Dissertation, University of Texas, 1965.

566

ROBERT C. NORDLIE

being indicative of basic differences in the intrinsic properties of the enzyme from the assorted tissues. As discussed both in Section II,C and Sections II1,E and IV,A, activities of the enzyme are very sensitive to various effects-often activity discriminating-exerted by a large number of metabolites and other compounds, which may well serve to control activities of the enzyme in vivo. Any definitive conclusions regarding possible differences in catalytic properties of the enzyme from various sources ultimately must await the availability of the purified enzyme, although some insight might be gained by a study of mixtures of the enzyme from the various sources. A straightforward discussion of catalytic properties of this enzyme is complicated by the facts that (a) the enzyme has not been successfully purified to an appreciable degree (see Section I1,A) , and (b) as discussed in Section II,C, various detergents, mechanical treatment, and hormonal therapies exert profound effects on such characteristics as pH optima, K,,, and Ki values, and V,,,,, and v values. Phosphotransferase activities of the enzyme are especially sensitive in the latter respect. Accordingly, account is taken in the following discussions of the nature of the enzyme preparations studied. Conveniently, preparations may be categorized generally as follows: (1) crude tissue homogenates or isolated microsoma1 preparations, (2) detergent-supplemented microsomal preparations, and (3) “partially purified” enzyme preparations. Properties of preparations in the latter two categories are, in general, quite similar since preliminary treatment by detergents generally has been employed in attempts to purify the enzyme. A. METHODSOF ASSAY Methods for assay of both hydrolase and phosphotransferase activities of the enzyme are outlined in detail by Swanson ( 4 ) , Harper (lor), Nordlie and Arion ( 5 ) , and Barman (108). I n the former assay, extent of hydrolysis is monitored by colorimetric assay of Pi liberated or determination of freed glucose with the aid of glucose oxidase. Glucose6-P produced in phosphotransferase reactions is conveniently measured with the use of glucose-6-P dehydrogenase, while assessment of activity with other sugars or polyols, including glycerol ( 6 5 ) , as phosphoryl acceptors involves the use of isotopically labeled substrates ( 5 ) . The assay of glucose-6-P phosphohydrolase activity has also been 107. A. E. Harper, in “Methods of Enzymatic Analysis” (H. U. Bergmeyer, ed.), p. 788. Academic Press, New York, 1965. 108. T. E. Barman, “Enzyme Handbook,” Vol. 2, p. 530. Springer, Berlin, 1969.

22. GLUCOSE-6-PHOSPHATASE

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recently considered by Baginski et al. (109). Belfield and Goldberg (110) have suggested a novel method for measuring this activity in the presence of acid and alkaline phosphatases. Pitot et a2. (111) have described methods for the continuous automated assay of PPi-glucose phosphotransferase activity of the enzyme. B. REACTIONS CATALYZED As discussed briefly in Section I,A, glucose-6-phosphatase is now known to be a multifunctional enzyme capable of catalyzing potent phosphohnsferase as well as phosphohydrolase reactions [see Eqs. (1)-(4)]. Compounds demonstrated to function as effective phosphoryl donors include fructose-6-P (SO), mannose-6-P ( 4 O ) , PP, (35-SS), a variety of nucleosidetriphosphates and nucleosidediphosphates-most effectively CTP, CDP, deoxy-CTP, ATP, ADP, GTP, GDP, and ITP (41, 46)carbamyl-P (43), phosphoramidate (44), phosphopyruvate (42, 43) and glucose-6-P itself (SO, 31). The various phosphoryl donors are also hydrolyzed by action of the enzyme (see preceding references). Eqqations (1)-(4), which describe these various activities, are given in Section 1,A.

1. Multifunctional Nature

Evidence for the involvement of a single, multifunctional enzymemicrosomal glucose-6-phosphatase-in the above variety of hydrolytic and synthetic reactions has been summarized recently by the author (10) as follows: (1) Inseparability of activities. The various activities depicted [in Eqs. (1144) 1 have been found associated in microsomal fractions prepared from liver (37, 38, 41, 1121, kidney (46,113), and intestine (46). Activities were also present together in preparations from all species tested, i.e., rat (9, 38, 41, fl2), rabbit (46), guinea pig (46, l l d ) , and man (116,116). Activities

109. E. S. Baginski, P. P. Foa, and B. Zak, Anal. Biochem. 21, 201 (1967). 110. A. Belfield and D. M. Goldberg, Life Sci. 8, 129 (1969). 111. H. C. Pitot, M. Poirier, A. Cutler, and C. Shannon, Automat. Anal. Chem., Technicon Sump., New York 1966 Vol. 1, p. 494. Mediad, Inc., White Plains, New York, 1967. 112. M. R. Stetten and H. L. Taft, JBC 239, 4041 (1964). 113. C. J. Fisher and M. R. Stetten, BBA 121, 102 (1966). 114. R. C. Nordlie and D. G. Lygre, in “Biochemistry of Mitochondria” (E. C. Slater, Z. Kaninga, and W. Wojfczak, eds.), Abstr., p. 2. Academic Press, New York, 1967. 115. B. Illingworth and C. Cori, BBRC 19, 10 (1965). 116. P. J. Collip, ABB 118, 106 (1967).

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ROBERT C. NORDLIE

appear simultaneously during embryological development (117). Both hepatic acid pyrophosphatase and glucose-&phosphatase are markedly depressed in von Gierke's disease (116). All activities remain together in constant ratios during partial purification (57, 4 1 ) . (2) Effects of inhibitors. All activities tested were inhibited by molybdate (32, 37), citrate (1181, oxalate (1181, and other metal-binding agents (119, 120), L-cysteine (46, 114, 119, 120), dithiothreitol (46, l l 4 ) , and phlorisin (101). Where determined, K I values calculated for the inhibition of the various activities by a given compound agreed closely (101, 118-180). (3) Thermal inactivation. All activities were equally susccptible to inactivation by relatively mild heating (37,41, &). (4) Kinetic studies. Excellent agreement was obtained for various K , and Ki values determined for compounds common to a number of the reactions catalyzed (37, 40, 4 1 , 45, 112). Activities observed when several substrates were combined in assay mixtures agreed well with theoretical values calculated on the basis of the involvement of a single enzyme (37,4 5 ) . ( 5 ) Responses to hormonal manipulations and fasting. Activities responded in a parallel fashion to glucocorticoid administration (93-95, 1 1 9 , alloxan diabetes and insulin therapy (41, 113, 117), growth hormone treatment (1181, and acute and prolonged fasting (90, 98, 116, 121).

Generally similar approaches have been employed more recently to establish the additional involvement of phosphoramidate (44) and carbamyl-P (42, 43) as phosphoryl donors and of glycerol (65) as a phosphoryl acceptor. 2. Substrate Specificity

a . Phosphohydrolase Activities. Comparative studies o f the relative rates of hydrolysis of a variety of phosphate esters by liver microsomes under conditions optimal for glucose-6-P hydrolysis have been carried out by Beaufay and de Duve (85), Crane (I,%?), and Maley and Lardy (183). Although a simple comparison of relative rates of hydrolysis is complicated by the fact that identical concentrations of all substrates were not employed, a general idea of relative reactivity may be obtained from their data, which have been compiled by Ashmore and Weber (7'). Relative rates a t pH 6.5-6.7, with substrate concentrations generally approaching saturation, were as follows: glucose-6-P 100; allose-6-P 90 ; 1,5-sorbitan-6-P 74; mannose-6-P 50 ; galactose-6-P 28; ribose-5-P 30 ; phenyl-P 24; a-glycerol-P 20 ; 2-deoxyglucose-6-P 20 ; 117. G.Bot and G. Vereb, Acla Biochim. Biophys. Acad. Sci. Hung. 1, 169 (1966). 118. R. C. Nordlie and D. G. Lygre, JBC 241, 3136 (1966). 119. R. C. Nordlie and P.T. Johns, BJ 104, 37P (1967). 120. R.C.Nordlie and P.T. Johns, Biochemistry 7, 1473 (1968). 121. W.J. Arion and R. C. Nordlie, BBRC 20, 606 (1965). 122. R. Crane, BBA 17, 443 (1955). 123. G.F.Maley and H. A. Lardy, JACS 78, 1393 (1956).

22. GLUCOSE-6-PHOSPHATASE

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fructose-6-P 19 ; tagatose-6-P 11; glucosamine-6-P 8; fructose-1-P 7; P-glycerol-P 5 ; 6-phosphogluconate 4 ; N-acetyl glucosamine-6-P 2; fructose-1,6-diphosphate 1; and glucose-1-P 0.5. Values for K, were reported by Crane (122) as follows: glucose-6-P 2 m M ; 2-deoxyglucose6-P 13 mM; galactose-6-P 50 mM; 1,5-sorbitan-6-P 10 m M ; allose-6-P 7 mM; glucosamine-6-P 10 mM; ribose-5-P 5 mM; glucoheptulose-7-P 18 mM; mannose-6-P 20 m M ; and L-sorbose-1-P 8 mM. Under approximately these same conditions, PPI is hydrolyzed approximately two-thirds as rapidly as glucose-6-P (98).. The rates of hydrolysis of nucleosidetriphosphates and diphosphates have not been adequately assessed because of contamination of microsomal preparations with additional nucleosidases (41, 124).

b. Phosphotransferase Activities. i. Phosphoryl donor specificity. Values of K , for PP, and a variety of nucleosidetriphosphates and nucleosidediphosphates, functioning as phosphoryl donors with glucose as acceptor a t pH 5.2, are tabulated in Table VI, as are relative V,,, values extrapolated for infinite concentrations of phosphoryl donor and 180 mM D-glucose. A “partially purified” enzyme preparation from rat liver microsomes was employed in these studies (41). A K , value of 1.6 mM has been obtained for carbamyl-P a t pH 7.0 with detergent-activated rat liver microsomes. At pH 5.5, with 10 mM phosphoryl donor and 180 m M glucose present, carbamyl-P: glucose TABLE VI

PHOSPHORYL DONORSPECIFICITY‘ Phosphoryl donor substrate

PPi CTP CDP GTP GDP dCTP ATP ADP ITP

Relative apparent VILIhXJL

1.8 3.1 3.8 3.6 2.7 2.6 4.7 11.1 6.1

1.0 0.59 0.56 0.25 0.23 0.27 0.27 0.32 0.29

Data from Nordlie and Arion (41). Details are given in text. Apparent maximal reaction velocities are expressed relative to PPi-glucose phosphotransferase activity, for which a value of 1.0 is arbitrarily assumed. 0

124. L. Ernster and L. C. Jones, J. Cell B i d . 15, 563 (1962).

570

ROBERT C. NORDLIE

phosphotransferase activity was approximately 1.6 times that of PPIglucose phosphotransferase activity, while a t pH 7.0 the ratio was approximately 5.8 (&, 83). A K , value of 2.8 m M for phosphoramidate serving as a phosphoryl donor with untreated rat liver microsomes has been observed a t pH 6.5 by Parvin and Smith ( 4 4 ) , who also reported values of approximately 0.45 for the ratio of phosphoramidate-glucose phosphotransferase/PP,glucose phosphotransferase with both rat liver and kidney microsomes. The latter assays were carried out a t pH 6.5 with 0.4 M glucose and 20 mM phosphate substrates. ii. Phosphoryl acceptor specificity. Stetten (125) has assessed the relative ability of more than 40 compounds to serve as phosphoryl acceptors at pH 5.4 with PP, as phosphoryl donor and Triton X-100activated rat liver microsomes as enzyme source. Pyrophosphate concentration was 0.08 M and phosphoryl acceptor 0.4 M in all instances, and incubations were carried out a t 30" for 30 min. Relative activities, based on arbitrary value of 100 with D-glucose, were noted as follows (125a) :

(A) Hexoses and derivatives: D-Glucose 100, 2-deoxy-~-glucose121, 3-deoxy-~-glucose 59, 3-O-methyl-~-glucose 93, a-methylglucoside 81, D-glucosamine 46, acetyl-D-glucosamine 19, D-mannose 117, L-rhamnose 0 , D-altrose 105, L-altrose 18, D-galactose 43, L-galactose 43, 2-deoxyD-galactose 23, D-galactose amine 12, fucose 0 , D-allose 62, D-talose 22, D-fructose 20, ~-fructose-6-P0, L-sorbose 19. (B) Heptoses and heptuloses: D-Sedo (altro)heptulose 30, D-mannoheptulose 33, D-glycero-D-guloheptose 8, D-glucoheptulose 20, L-glucoheptulose 8, L-galactoheptulose 40. (c)Pentoses: D-Ribose 24, 2-deoxyribose 14, D-arabinose 12, L-arabinose 12, D-lyxose 0, L-lyxose 0, D-xylose 0. (D) Triose: D,L-Glyceraldehyde 11. (E) Alcohols: D-Mannitol 25, sorbitol 20, galactitol 10, glycerol 36, i-inositol 0 , ethanol 0. (F) Other compounds: Maltose, gluconic acid S-lactone, and serine 0.

Of particular interest is the finding of Stetten and Rounbehler (65) that glycerol-l-P, rather than glycerol-3-P which is formed by glycerol kinase, is produced by PPi-glycerol phosphotransferase action of glucose-6phosphatase. Nordlie and Arion (41) have, in a few instances, assessed K , values 125. M. R. Stetten, JBC 240, 2248 (1965). 125a. Copyright (1965), American Society of Biological Chemists, Inc. Reproduced by permission.

57 1

22. GLUCOSE-6-PHOSPHATASE

for phosphoryl acceptors other than D-glucose. At pH 5.2, with 10 rnM PPi and partially purified rat microsomal enzyme, K,,, values of 100 mM for D-mannose, 133 mM for 2-deoxy-~-glucose,and 571 mM for wgalactose were evaluated. Lygre and Nordlie (1%) have studied the relative ability of a variety of sugars, sugar derivatives, and polyols to inhibit PP,-glucose phosphotransferase activity of rabbit intestinal glucose-6-phosphatase. These inhibitions, which were competitive with respect to glucose in all cases, presumably involve the tested compounds' functioning as alternate phosphoryl acceptors with glucose. Values of Ki obtained a t pH 6.0 were as indicated in Table VII. It was concluded on the basis of these observations and a consideration of the findings of Stetten (125) that, ii. . . while an absolute structural specificity is not exhibited by this enzyme, those sugar molecules with the structural and configurational features given in [Fig. 51 bind much more effectively to the enzyme than do other compounds tested" (126).

C. THERMODYNAMIC CONSIDERATIONS With the exception of transphosphorylations involving sugar phosphate esters and sugars (for which AF" 0 ) , all reactions catalyzed by

=

TABLE VII INHIBITION OF INTESTINAL PP~-GLUCOSE PHOSPHOTRANSFERASE ACTIVITY BY SUGARS, SUGAR DERIVATIVES, AND POLYOLP Ki

Compound Hexoses and derivatives 2-Amino-~-glucose 2-Deoxy-~-glucose a-Methylglucoside 6-Methylglucoside 3-0-Methyl+-glucose D-Mannose D-Galactose D-Fructose Heptose D-Glucoheptose

128. D. G. Lygre and R.

(MI 0.12 0.38 0.17 1.4 0.17 0.16 0.95 1.7

1.5

Compound Pentoses D-Xylose GXylose D-Ribose D-habinose D-Lyxose Tetrose D-Erythrose Poly01s i-Erythri to1 D-Mannitol

C.Nordlie, BBA 178, 389 (1969)

Ki (MI 0.75 0.90 1.1 1.8 1.9 5.0 1.7 2.2

572

ROBERT C. NORDLIE

Fro. 5. Haworth formula indicating structural and configurational features essential for efficient binding of aldohexoses with glucose-6-phosphatase. From Lygre and Nordlie (126). Copyright (1965), Elsevier Publishing Co. Reproduced by permission.

glucose-6-phosphatase are exergonic. At physiological pH, A F O for the hydrolysis of glucose-6-P [reaction (1) ] is -2600 cal/mole (127), while a considerably more negative value (-7000 cal/mole) obtains for the hydrolysis of PP, (and the various nucleoside triphosphates and nucleoside diphosphates) [reaction (4) ] (128). While the coupling of the hydrolysis of PP, produced in synthetasecatalyzed reactions may tend to drive these otherwise energetically endergonic synthetic processes toward completion, it has been suggested that some of the energy which otherwise would be dissipated accompanying the hydrolysis of PPI might be conserved for further use by the organism if the phosphoanhydride were to serve as a phosphoryl donor in physiologically significant reactions (1.29,130). Pyrophosphate-glucose phosphotransferase activity of microsomal glucose-6-phosphatase would appear to serve such a role, for in the dynamic, living cell (in contrast with a closed thermodynamic system approaching equilibrium) this activity serves to remove PP, while a t the same time generating the important metabolite glucose-6-P. The AF” for this phosphotransferase reaction [reaction (3a)l may be calculated as the difference in AF” values for the hydrolysis of PP, minus that for hydrolysis of glucose-6-P. A value of -400 cal/mole is thus obtained.

D. KINETICSTUDIES AND REACTION MECHANISM Because pH optima, K,, v, V,,,, and Ki values for the various activities of this enzyme are highly dependent upon the physical state of the enzyme preparation used-i.e., whether untreated microsomes, detergent-supplemented microsomes, partially purified enzyme preparation, 127. E. S. West, W. R. Todd, H. S. Mason, and J. T. VanBruggen, “Textbook of Biochemistry,” 4th ed., p. 884. Macmillan, New York, 1966. 128. E. S. West, W. R. Todd, H. S. Mason, and J. T. VanBruggen, “Textbook of Biochemistry,” 4th ed., p. 891. Macmillan, New York, 1966. 129. A. Kornberg, Advan. Enzymol. 18, 191 (1957). 130. D. Stetten, Am. J . Med. 28, 867 (1960).

TABLE VIII SOMECATALYTIC PROPERTIES OF HYDROLYTIC AND SYNTHETIC ACTIVITIESOF HEPATICGLUCOSE~-PHOSPHATASEO Without detergent Activity

Ref.

Glucose-6-P phosphohydrolase PPi Phosphohydrolase PP,-Glucose phosphotransferase

PPi D-Glucose ATP-Glucose phospho transferase ATP D-Glucose CDP-Glucose phosphotransferase (partially purified preparation) CDP D-Glucose Carbamyl-P-glucose phospho transferase Carbamyl-P D-Glucose PPi-Glycerol phospho transferase (high pH-treated microsomes) PPi Glycerol Phosphoramidate-glucose phosphotransferase Phosphoramidate D-Glucose

K,(mM)

Detergenetreated preparations

PH PH Optimum Kmb((mM) Optimum

1.8 3.1

6.0 5.0

1.2 2.0

6.5 6.0

2.9 127

4.5

1.3 80-100

6.0

-

4.5

4.7 90

u.2-5.5

3.8 90

5.2-5.5

1.6 80

5.5-6.0

(9)

-

-

-

-

5. 300.

5.0

(44,131)

13 60-80

~

8.3

2.8

~

Unless otherwise indicated, data presented are from reference 98 and were obtained with liver homogenates or microsomal suspensions from normal, fed rats. K, values somewhat lower than those recorded have been observed with partially purified preparations (37, 40, 46). K, values were observed a t the following pH values: glucose-6-P phosphohydrolase, pH 6.5;PP, phosphohydrolase, pH 5.5; PP,-glucose phosphotransferase, pH 5.5; nucleotide-glucose phosphotransferases, pH 5.2; carbamyl-P-glucose phosphotransferase, pH 7.0; PP,-glycerol phosphotransferase, pH 5.4; and phosphoramidate-glucose phosphotransferase, pH 6.5 (no detergent) or pH 8.0 (detergent treated). * K, values for PP, as low as 0.96 mM (46) have been observed with partially purified preparations, and values of 3.3 X lO-'M have been noted in the presence of 0.3% cetrimide (89). K, values of 80 mM for glucose have been obtained with partially purified preparations (40). 573

574

ROBERT C. NORDLIE

phospholipase-treated microsomes, or mechanically disrupted preparations (see Section I1,C)-and because kinetic parameters are highly pH dependent (118), an all-inclusive consideration of the kinetics of the large variety of reactions which have been characterized is impossible here. Some representative catalytic properties of the various hydrolytic and synthetic activities of the enzyme are presented in Table VIII. For a more comprehensive consideration of the various activities, the interested reader is directed to individual articles by Nordlie and coworkers (9, 10, 37, 40, 41, 43, 45, 80, 98, 118), Stetten and colleagues (38, 65, 112, 125), Segal and Washko (72), and Parvin and Smith

(a).

1. Eflects of p H

Pyrophosphate-glucose and nucleotide-glucose phosphotransferase activities exhibit maximal activity a t pH 4.5 in untreated microsomal preparations or homogenates, and a t p H 5.5-5.7 after detergent treatment and/or partial purification of the enzyme. Typical activity-pH profiles obtained with microsomal preparations from normal control, diabetic, and cortisone-treated animals, without and with supplemental detergent added, are presented in Fig. 2. Nordlie and Lygre (118, 132) have carried out detailed pH kinetic studies of PPi-glucose phosphotransferase, CDP-glucose phosphotransferase, and glucose-6-P phosphohydrolase activities with partially purified rat liver enzyme preparations, and they have concluded, on the basis of sharp, unit-changes in slopes of Dixon (133) plots of pK, for phosphate substrates vs. pH near 6.5, that enzyme-bound histidine may participate directly in the catalytic process. Values of K , for glucose were unaffected by reaction mixture p H ; K , values and relative V,,, values calculated for infinite concentrations of all substrates are tabulated in Table IX. It is apparent from these and other data that, while v and K , values change quite markedly with assay pH, V,,, values for reactions involving PPi, CDP, and glucose-6-P are considerably less sensitive to change in pH, especially between pH 5 and 6.5. Segal and Washko (72) have made similar observations for glucose-6-P phosphohydrolase previously. It thus appears that the rather marked inflections in activity-pH profiles generally observed with activities of the enzyme within this range of hydrogen ion concentrations result principally (but not ex131. M. E. Holaer, K.D. Johnson, and R. A. Smith, BBA 122, 232 (1966). 132. D. G. Lygre and R. C . Nordlie, unpublished observations (1966). 133. M. Dixon, BJ 55, 161 (1963).

575

22. GLUCOSE-6-PHOSPHATASE TABLE IX VARIATIONS IN K, VALUESAND V,,

Clucose-6-P phosphohydrolase

PPi-Glucose phosphotransferaseb

4.65 5.10 5.60 6.05 6.50 7.00 7.43 8.09 8.58 8.92

(a)Vmax' 10.0 2.2 0.58 0.48 0.43 1.1 3.4 8.3 21.7 48.6

0.79 0.94 0.88 0.90 1.00 0.96 0.73

CDP-Glucose phosphotransferaseb

Km(PPi)

K m

PH

VALUES WITH ASSAYpHa

PH 4.75 5.15 5.60 6.08 6.50 7.00

Km(CDP)

(d)Vmax'

PH

2.2 1.2 0.96 0.94 2.1 6.1

4.52 4.98 5.50 6.00 6.50 6.92

1.14 1.24 1.36 1.28 1.04 0.60

Vme.xc

7.9 4.9 3.8 5.2 6.7 10.1

0.49 0.69 0.83 0.88 0.67 0.21

From Nordlie and Lygre (118, 1%). K , values for D-glucose (80 mM) were independent of pH. V,,, values are expressed relative to an arbitrary value of 1.00 for glucose6-P phosphohydrolase activity at pH 6.5. a

clusively-see V,,, values in Table IX) from variations in K,,,with reaction mixture pH. In contrast, the rapid drop in phosphotransferase activities above pH 6.5 appears to involve alterations in velocity terms as well as in the affinity of the enzyme for phosphoryl substrates. The chemical nature of the ionizable phosphate substrates appears to be extremely important in establishing activity of the enzyme a t various p H values. On the basis of the observations that (a) glucose-6-P phosphohydrolase (72), mannose-6-P: glucose phosphotransferase ( 1 3 4 , and mannose-6-P phosphohydrolase (13.4) activities are manifest over a broad range of pH, including both acid and alkaline values; (b) similarities of activity-pH profiles for PPI- and nucleotide-glucose phosphotransferase reactions and for hydrolase reactions involving these phosphate substrates (37, 41, 98); (c) absence of significant PPI- or nucleotide-glucose phosphotransferase activity above approximately pH 7 (37, 41) ; and (d) potent inhibition by PP, and nucleotides of glucose6-P phosphohydrolase activity a t alkaline as well as acid p H (103), it has been concluded by Nordlie et al. (134) that '(. . phosphate substrates may bind to the active enzymic site over a wide range of

.

134. R. C. Nordlie, T. L. Hanson, and P. T. Johns, Federation Proc. 28, 412 ( 1989).

576

ROBERT C. NORDLIE

pH, but enzyme-substrate complexes formed from PP,-3, ADP-3, or ATP4 (in contrast with PPi-2,ADP-2, ATP-3, glucose-6-P2, or mann0se-6-P-~ which are active substrates) are catalytically inert and do not react further to produce phosphoryl-enzyme intermediates.” The inability of the electron-rich nucleophile molecules to approach phosphorus atoms of highly electronegative binary complexes (see Section III,D,5,a and Fig. 4) formed from the enzyme and the more extensively ionized species of phosphoanhydrides extant in the alkaline pH range is believed to explain this lack of reactivity of the latter compounds at the higher pH values ( 4 3 ) . Support for this concept is found in the results of recent studies (4.2, 4.9, which indicate that carbamyl-P, which according to the above theory should be active a t alkaline as well as acid pH values, displays potent phosphotransferase ability with the enzyme even a t pH > 7. Mechanistic significance of these observations is considered further in Section III,D,5. 2. Effects of Substrate Concentration Representative K, values for a variety of substrates participating in a number of the enzyme’s reactions under various assay conditions are given in Sections III,B,2 and II1,D and in Tables VIII and IX. Detailed kinetic studies of liver (40, 41) and kidney (45) PPi-glucose phosphotransferase, mannose-6-P:glucose phosphotransferase, and nucleotideglucose phosphotransferase activities, in which initial reaction velocities were assayed as a function of varied concentrations of phosphoryl donor at several constant levels of glucose, and as functions of varied concentrations of glucose a t several constant levels of phosphoryl donor, all have produced results which, when analyzed graphically according to the method of Lineweaver and Burk (135), yielded families of linear plots intersecting in each instance at common points on the axis of abscissas, as in Figs. 6A and B. The independence of K , values from concentrations of second substrates in each case is readily apparent from these observations. Further, as discussed in detail in Section III,D,5,a, these observations have been found useful in establishing a generalized reaction mechanism (see Fig. 4) encompassing both phosphohydrolase and phosphotransferase activities of the enzyme. The experimentally observed rates of reactions with phosphate substrates present individually and in various combinations have been found in good agreement with theoretical values calculated on the basis of the assumption of the involvement of a common binding site for such compounds, as indicated in Fig. 4 (37, 46). 135. H.Lineweaver and D. J. Burk, JACS 56, 658 (1934).

22. GLUCOSE-6-PHOSPHATASE

577

FIQ.6. Effects of varied substrate concentrations on velocity v of phosphotrunsferase reactions, described in terms of conventional (136) double-reciprocal plots. In (A) has been assessed as a function of concentration of phosphoryl donor D with phosphoryl acceptor constant at concentrations A], A,, A,, or A,. In (B) v has been determined as a function of varied concentrations of phosphoryl acceptor A with phosphoryl donor held constant a t concentrations D,, D,, Ds, or D,. (Generalized from references 40, 41, and 46.)

3. Effects of Temperature The effects of temperature both on the stability of the enzyme (26, 37,

41, 46) and on the catalytic reaction itself (46, 112) have been studied. The enzyme is considerably more labile to mild heating in the absence of substrates than is microsomal acid phosphatase (26, 37, 41, 46). Glucose-6-P (26, 136), Pi (%), glucose (1361, PP, (119, 1S6), and various amino acids (136) protect to some degree against thermal inactivation as do certain metal chelators which inhibit the reaction (120). 136. S. M. Huang, Master's Thesis, University of North Dakota, 1969.

578

ROBERT C. NORDLIE

Detergents labilize the enzyme to thermal inactivation in the absence of substrates (26, 90),while activation by exposure of the enzyme to high pH does not (90).The enzyme appears stable to freezing for several months (137). Both synthetic and hydrolytic activities of the enzyme exhibit remarkably high amounts of activity even a t 0" (119). At pH 6.0, an energy of activation value of 12.3-12.5 kcal/mole has been determined for glucose-6-P phosphohydrolase, PPi-glucose phosphotransferase, and CDP-glucose phosphotransferase activities of rabbit intestinal microsomal enzyme ( 4 6 ) . Segal et al. (71) observed a very similar value (12.5 kcal/mole) for glucose-6-P phosphohydrolase activity of the rat liver enzyme. 4. Efjects of Activators and Inhibitors

A wide variety of compounds and treatments have been reported to activate microsomal glucose-6-phosphatase phosphotransferase. These are described briefly below; the possible relevance of the action of a number of these factors in the biological regulation of activities of the enzyme has been considered in Section II,C,3.

a. Activators. Activation of enzymic activities by bile acids, longchain fatty acyl-CoA esters, lysolecithin, and various other natural and synthetic detergents, high pH treatment, mechanical disintegration, and glucocorticoid therapy have been described in Section I1,C. Activation of glucose-6-P phosphohydrolase activity by o- and p-tyrosine has also been described (138,139),although experiments by the author (66,119) indicate that stabilization of the enzyme by these compounds, which are capable of metal chelation, may actually be the phenomenon involved. Alloxan has been reported to increase V,,, values of the hydrolase reaction as well as to increase K,,, for glucose-6-P (140). b. Inhibitors. A wide variety of compounds have been found to inhibit activities of the enzyme in vitro. These compounds are listed in Table X, along with pertinent literature references, notes on the kinetic nature of inhibition where known, and other relevant information. Where rather detailed studies have been made, it has been found that the actions of inhibitors on enzymic activities often are quite significantly affected by such factors as reaction pH and the presence of 137. G. T. Con and C. F. Cori, JBC 199, 661 (1962). 138. G. Feuer, L. Golberg, and K. I. Gibson, Food Cosmet. Tozicol. 4, 283 (1966). 139. G. Feuer, L. Golberg, and K. I. Gibson, BJ 93,6P (1964). 140. 9. V. Jakobason and G. Dallner, BBA 169, 380 (1968).

22.

579

GLUCOSE-6-PHOSPHATASE

TABLE X INHIBITORS OF HYDROLASE AND PHOSPHOTRANSFERASE ACTIVITIES OF GLUCOSE-~-PHOSPHATASE~ Inhibitors

References

Comments

I. Inhibition Demonstrated with Both Phosphohydrolase and Phosphotransferase Activities” Krebs’ tricarboxylic acid cycle intermediates

Glycogen Various sugars and polyols

Amino acids

Lipids Long-chain fatty acyl-CoA esters Fatty acids Lysolecithin Other phospholipids

(118)

(66) (40, 41, 1% 1866)

(119, 180, 136)

Significant inhibition of all activities below pH 7 by citrate for which a Ki value of 5.2 mM has been observed at pH 6.5. Isocitrate much less effective, and only minor effects noted with cis-aconitate, 2-malate, succinate, Q ketoglutarate, and oxalate. Malonate, tartrate, gluta am ate, glutarate, adipate, 8-hydroxy-8-methyl glutarylCoA, and 8-hydroxybutyrate also found ineffective as “instantaneous” inhibitors Slight activation by pyruvate; phosphoenolpyruvate a competitive inhibitor of phosphohydrolase activity and also a phosphoryl donor in the transferase reaction No effects noted Inhibit phosphatase activities by serving as phosphoryl group acceptors in competition with water. Ki(K,) values considerably higher than for glucose, except for D-mannoe and 2-deoxy-~-glucose kCysteine, o-tyrosine, and kleucine inhibit and also stabilize enzyme against thermal inactivation. A variety of other amino acids are also effective to some degree in the latter respect Micromolar to lO-’M levels affect hydrolytic and synthetic activities in an activity-discriminating, pH- and concentration-dependent manner Millimolar-10-2 M levels also exert activity-discriminating effects The former activates all activities, the latter are necegsary for enzymic activity and appear to maintain enzyme in a stable, active conformational state

ROBERT C. NORDLIE

TABLE X (Continued) ~~

Inhibitors Nucleo tides

References

Comments

(10,103)

Various nucleoside di- and triphosphate compounds inhibit glucose-6-P phosphohydrolase activity competitively over a broad range of pH (pH 5-8). Also function as phosphoryl donors in phosphotransferase reactions. Effects are markedly potentiated by various detergents Inhibits in a manner similar to that of nucleotides. Also inhibits phosphotransferase activity Inhibits both synthetic and hydrolytic activities competitively with respect to phosphoryl substrates. Ki = 20 mM in absence of detergents Optimal concentrations of deoxycholate, cholate, Triton X-100, and cetyltrimethylammonium bromide activate, as does urea. Activation of phosphotransferase > 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

<

<

<

155a. J. D. Lueck and R. C. Nordlie, unpublished observations (1970).

586

ROBERT C. NORDLIE

the enzymc, as experimentally observed (see Sections III,B,l and III,D,4,b). Further, rate equations ( 8 ) and ( 9 ) , derived on the basis of the proposed mechanism, predict that glucose should function as an apparent noncompetitive inhibitor of phosphohydrolase reactions, as observed, although the actual phenomenon involved is a competition between water and hexose as alternate phosphoryl group acceptors (@). Of particular interest from the physiological point of view is the fact that the K , value for glucose, as defined by this kinetic treatment [see Eq. (5b)I [which equals K , for glucose in Eqs. (8a) and (9a) since k , = k , and k , = k 8 ] ,contains a watcr concentration term in the numerator. Conceivably, in the environment of the enzyme within the phospholipid-rich membrane of the endoplasmic reticulum, water concentrations may be much less than 55.5 M , and the K , value for glucose under such conditions would be appreciably lower (10). Segal (31) and Hass and Byrne (30)have pointed out the incompatability with experimental observations of such alternate mechanisms as one not involving kinetically significant binary enzyme-phosphoryl donor complexes and mechanisms involving two distinct glucose-binding sites on the enzyme. Their argument holds also for data obtained by Arion and Nordlie (40) with PP,-glucose, mannose-6-P :glucose, and nucleotide-glucose phosphotransferase activities. b. Nature of the Active Site. Although the enzyme has not been purified to an appreciable degree, some insight relating to the chemical nature of the active site has been obtained by indirect means. A study (118) of the effects of pH on K , values for phosphate substrates and K , values for the inhibitor citrate, described in Sections III,D,2, and III,D,4,b, and Table IX, suggests that cnzyme-bound histidine may participate in the binding of phosphoryl substrates. Feldman and Butler (156) have recently published a preliminary report indicating that “incubation of rat liver microsomes with “‘P-glucose-6-P results in rapid incorporation of 32Pinto protein as N-3-phosphoryl histidine,” thus confirming Nordlie and Lygre’s suggestion (118) of the involvement of an enzyme histidine residue in the catalytic mechanism. An excess of nonradioactive glucose-6-P or PP, almost completely prevented this incorporation of label from 3’P-glucose-6-P, but cold P, was without effect (156). Parvin and Smith (4-4) have also demonstrated incorporation of 32Pfrom phosphoramidate and PP, into liver microsomal enzyme preparations and the release of this 32Pby mild acid hydrolysis. 156.

F.Feldrnan and L. G.Butler, BBRC 36, 119 (1969).

22. GLUCOSE-6-PHOSPHATASE

587

Inhibition by a variety of metal-binding agents competitive with respect to phosphoryl substrates (118-190) has suggested that an enzyme-bound divalent cation (other than MgZ+) may participate also in the binding of phosphate substrates. Observed inhibition by p-chloromercuriphenyl sulfonate and iodoacetate suggests the possibility that sulfhydryl groups may also be involved at, or near, the active enzymic site (119,120). c. A Proposed Reaction Mechanism. Proposed mechanisms for both phosphohydrolase and phosphotransferase activities of the enzyme, based on presently available information, are presented in Figs. 7 and 8. Mechanistic concepts incorporated in these diagrams are based on the following experimental observations: (i) The enzyme is multifunctional in nature, catalyzing various phosphotransferase as well as phosphohydrolase reactions (see Sections I,A,2 and III,B and Fig. 4). (ii) Kinetically significant binary enzyme-phosphoryl substrate complexes are involved, as is a phosphoryl-enzyme complex (SO, 31, 40). (iii) An imidazolium group of enzyme-bound histidine has been implicated in the active site (118). (iv) The phosphoryl-enzyme intermediate appears to involve the N-3-phosphoryl derivative of enzyme-bound histidine (4.4, 156). (v) The exchange reaction [ transphosphorylation, reaction (2a) ] between glucose-6-P and glucose catalyzed by the enzyme does not involve an incorporation of lSO from HZ1*Ointo glucose-6-P (155). (vi) Water and glucose compete as acceptors of the phosphoryl group from the phosphoryl enzyme (SO, 31, 40). (vii) Strong evidence for the involvement of protein-bound divalent cation as part of the active site has been presented (119,1i?O). (viii) The various phosphate substrates, and such inhibitors as classic metal chelators, Pi and HC03-, all compete for a common binding site (37-46, 104,ii9,120, 141). (ix) Orthophosphate inhibits, competitively with respect to phosphate substrates, both phosphohydrolase and phosphotransferase reactions (103, 104) but does not react with excess glucose in the presence of the enzyme to produce glucose-6-P by reversal of the phosphohydrolase reaction (30)* (x) Incorporation of isotopic label from szPiinto protein-bound 3-Nphosphohistidine cannot be demonstrated under conditions where incorporation of isotope from g l ~ c o s e - 6 - ~occurs ~ P (166). (xi) Thus, although PI and phosphate substrates compete for a com-

588

ROBERT C. NORDLIE

Clc-0-H

FIG.7. Proposed reaction mechanism. Details are

given in Section 111,D,5,

22.

589

GLUCOSE-6-PHOSPHATASE

Active substrates and inhibitors

0-

I

Inhibitors only

OH

I

Ad-0-P-0-P-O-

0

00I l Ad-0-P-0-P-0-P-0-

a

II

0

H’

OH

l

4

0

,

11

OH

II

H,N-C-0-P-0-

,

0

II

OH

Carbamyl-P (XVIII)

’-

I

0

00Ad-0-P-0-P-0-P-0- l

P K , = 7.0

H+

0-

l

0

( X V ) ATP‘-

0 pK. = 6.1 I II ~ c L c - 0 - P - 0 I H+ O-

(XVI) Glc-6-P’-

0

0-

7I

(XIV) ATP’

CLC - 0 - p - o -

0I Ad-O-P-O-P-0-

pK, = 7.0

(XVII) Glc-6-P2-

pT PK, = 4.9

H+

0

I1

H2N-C-0-P-0-

0

II I

0-

Carbaniyl-P (XIX)

FIG.8. Ionic species of some phosphate compounds serving &s substrates and/or inhibitors of glucose-6-phosphatase (167). See Sections III,D,4 and 5 for details. 157. pK, values ( p = 0) are from H. H. Sober, ed., “Handbook of Biochemistry.” Chem. Rubber Publ. Co., Cleveland, Ohio, 1968.

590

ROBERT C. NORDLIE

mon enzymic site, binding of the former to the enzyme does not involve formation of a P-N covalent bond, although interaction of the latter with the enzyme does. (xii) Plots of pK, for phosphate substrates against pH show two sharp inflections, each with unit changes in slopes, between p H 5.6 and 7 (118). (xiii) Hydrolase and phosphotransferase activities with hexose-P (72, 134), carbamyl-P (42, 43), or phosphoramidate (44), but not with phosphate anhydrides (PP,, nucleosidetriphosphates, and nucleosidediphosphates) as substrates (37, 41, 45, 46) , are demonstrable a t and above pH 7.5 as well as a t lower pH values. (xiv) Although not reactive a t alkaline pH, these phosphate anhydrides nevertheless do bind a t this pH to the active site of the enzyme, as demonstrated by their ability to inhibit effectively in a competitive manner hydrolytic and synthetic reactions involving glucose-6-P or mannose-6-P as substrates (103, 13.4). (xv) K , values for these phosphate anhydrides, acting as competitive inhibitors with respect to hexose-6-P substrates, are equal to K , values for these compounds functioning as phosphoryl donors a t p H values where both kinetic parameters can be evaluated (37, 38, 41, 45, 103). These observations indicate that both modes of behavior of the anhydrides depend upon their binding to a single enzymic site. (xvi) Added divalent cations inhibit both hydrolytic and synthetic activities of the enzyme (25, 26, 41, 145). Reactions involving glucose-6-P as substrate are considered mechanistically in structures (11) and (111) on the left-hand half of the upper portion of Fig. 7, while corresponding reactions involving phosphate anhydrides (RP) are depicted in structures (IIa) and (IIb) in the righthand side of the upper portion of this diagram. Glucose-6-P and R P are presented as representatives of two general groups of phosphate substrates. Such compounds as mannose-6-P, fructose-6-P, carbamyl-P, and phosphoramidate may be substituted for glucose-6-P, while various nucleosidetriphosphates and nucleosidediphosphates and PPi fit generally the mechanistic considerations given R P in Figs. 7 and 8 (157), and in the discussion below. The formations of phosphoryl-enzyme complexes from glucose-6-P and RP, respectively, are shown in structures (I), (II), (III), and (IV) and (I), ( I I a ) , (IIIa), and (IV). Water and glucose, if present, then compete for this phosphoryl-enzyme complex, leading ultimately to the release of Pi (hydrolase) or glucose-6-P (phosphotransferase), respectively, as indicated in structures (IV), (V), (VI) , and (I) and (IV), (Va), (VIa), and (I) in the bottom half of Fig. 7. The active enzymic site [see ( I ) ] is shown to contain protein-bound

22. GLUCOSE-6-PHOSPHATASE

591

divalent cation M2+and enzyme-bound histidine, the imidazolium group of which is shown in Figs. 7 and 8 ( 1 5 8 ) .Both glucose-6-P and phosphate anhydride substrates are considered to carry two negative charges. The catalytic reactions are considered to proceed as follows (capital Roman numerals correspond with those in Figs. 7 and 8) : (i) Glucose-6-P or R P binds to protein-bound divalent cation of the free enzyme (I) to produce complexes (11) and (IIa), respectively. (ii) Electrons of imidazolium nitrogen of enzyme-bound histidine make a nucleophilic attack on the phosphorus atom of phosphate substrates (111) and (IIIa). (iii) Glucose or R- dissociates leaving the phosphoryl-enzyme complex (IV). (iv) Water and glucose competitively interact with the phosphorylenzyme complex (V) and (Va) . (v) Electrons of oxygens of water or glucose make a nucleophilic attack on the P atom of the enzyme-bound N-3-phosphohistidine complex (VI) and (VIa). (vi) Orthophosphate or glucose-6-P thus produced dissociates yielding free enzyme (I). The ability of Pi to inhibit competitively with respect to phosphate substrates ( 1 0 4 , while not being incorporated into protein-bound phosphohistidine (156) or participating in glucose-6-P synthesis by reversal of the hydrolytic reaction (SO), also is explainable in terms of these mechanistic concepts. As shown in (XX), Fig. 8, binding of the Pi to enzyme-bound metal ion, without further formation of a P-N bond with imidazole N, would explain all of these experimental observations. The fairly complicated effects of varied pH on the differential patterns of participation of various phosphate compounds as substrates and as inhibitors, as described in detail above, also may be rationalized on the basis of this mechanism. Prevailing ionic species of some phosphate substrates a t various pH values are pictured in Fig. 8, structures (X)( X I X ) , along with relevant pK, values. Glucose-6-P2- (XVII), PPi2( X ), nucleosidediphosphate2- ( X I I ), nucleosidetriphosphate3- (XIV), mannose-6-P2- (see XVII), and carbamyl-P2- (XIX) are effective both as substrates and as inhibitors, for both phenomena are very prominent 158. pK. values in the range 5.6-7.0 have been reported [see Edsall (1691, Dixon and Webb (1601, and Boyer (161)l. 159. J. T. Edsall, in “Proteins, Amino Acids, and Peptides aa Ions and Dipolar Ions” (E. J. Cohn and J. T. Edsall, eds.), p. 444. Reinhold, New York, 1943. 160. M. Dixon and E. C. Webb, “Enzymes,” 2nd ed., p. 143, 1964. 161. P. D. Boyer, Science 141, 1147 (1963).

592

ROBERT

C.

NORDLIE

over the range of pH in which these ionic species predominate. I n contrast, the dissociation of an additional hydrogen ion from phosphate anhydrides to produce PP:- (XI), nuc1eosidediphosphate”- ( X I I I ), or nucleosidetriphospliate4- (XV) leads to a loss of activity of these compounds as substrates for either phosphotransferase or phosphohydrolase activity, although the ionic species thus produced remain capable of binding to the active enzymic site as indicated by their continued ability to inhibit competitively reactions involving glucose-6-P, mannose-6-P, etc. (103, 134). These observations are rationalized in terms of the more highly ionized species of these compounds binding to enzyme-bound divalent cation, for which they compete with glucose-6-P, mannose-6-P, carbamyl-P, or phosphoramidate, without, however, reacting further with enzyme-bound histidine imidazole N. As illustrated in structure (XI) in Fig. 8, nucleophilic attack of electrons of enzyme-imidazole N on the P atom of the more highly ionized species of phosphate anhydrides is believed to be precluded owing t o repulsion effects of unmasked electrons surrounding oxygen atoms of PP:-, nuc1eosidediphosphate“-, or nuc1eosidetriphosphate’-. Since hexose-6-P (XVII), carbamyl-P (XIX) (and phosphoramidate), even when totally dissociated, have a maximum of two negatively charged oxygens, which are effectively neutralized by coordination with enzyme divalent cation [see structures (111) and (IIIa), for example] , these compounds remain active even in the alkaline pH range. Added divalent cations would be expected to inhibit, as experimentally observed (95, 96, 41, 1 & ) , by chelating with phosphate substrates and thus interfering with their binding t o enzyme-bound metal as depicted in (11) and (IIa) in Fig. 8.

E. CONTROL

OF

GLUCOSE-6-PHOSPHATASE

PHOSPHOTRANSFERASE

ACTIVITIES Recent studies indicate that the various phosphohydrolase and phosphotransferase activities of glucose-6-phosphatase are affected by numerous metabolites (see Table X and Sections II,C and 111,D14). The possible significance of observed activation or inhibition by a number of these compounds in vitro relative to regulation of both types of activity of the enzyme in vivo has been considered in a number of instances. Possible modes of control of net glucose release, involving the regulation by a variety of factors, of both hydrolytic and synthetic activities of the enzyme, have been discussed in considerable detail in earlier reviews by the author (9, 1 0 ) .

593

22. GLUCOSE-6-PHOSPHATASE

The following factors and experimental observations with the enzyme all are suggestive of metabolic control a t the glucose-6-phosphatase phosphotransferase level: (i) The strategic, crossroad metabolic position of the enzyme (see Fig. l ) , which catalyzes the terminal reaction in both glycogenolytic and gluconeogenic processes. (ii) The relatively low maximal catalytic capacity of the enzyme compared with many other enzymes involved in glycolysis and gluconeo-

"

5

6

7

8

9

OH

FIG.9. Comparison of the relative rates of hepatic glucose phosphorylation by hexokinase (A),and glucokinase (W), and by ATP-glucose phosphotransferase (V and v), PPi-glucose phosphotransferase (0 and O), and carbamyl-P: glucose phosphotransferase (*)activities of glucose-6-phosphatase at various pH values. Filled symbols depict results with preparations from normal rats, while open symbols indicate results with preparations from alloxan-diabetic animals. Liver homogenates were supplemented with deoxycholate, to 0.2% (w/v), prior to assay for activities of glucose-6phosphatase. Activities are expressed for 5 mM phosphoryl donors and either 100 mM glucose (glucokinase) or 120 mM glucose (phosphotransferase activities of glucose-6-phosphatase). Glucokinase and hexokinase plots are based on experimental observations of Sharma et al. (1631, Salas e t al. (164, 166), and Fromm and Hanson (unpublished observations). Phosphotransferase activities of glucose-6-phosphatase are from the work of Nordlie and co-workers; see references 9 and 118 for additional details. The horizontal double arrow indicates the predominant range of intracellular pH values for liver.

594

ROBERT C. NORDLIE

genesis. Expressed on the basis of units of substrate converted to products per minute per gram of wet liver a t 37”, glucose-6-P phosphohydrolase has been reported to have a maximal catalytic capacity of 17, as compared with values of 15 for fructosediphosphatase, 6.7 for pyruvate carboxylase and phosphoenolpyruvate carboxykinase, which are also key enzymes involved in gluconeogenesis ; and relatively high values of 150 for phosphoglycerate kinase, 170 for glyceraldehyde-3-P dehydrogenase, and 230 for lactate dehydrogenase (162). (iii) The multifunctional nature of the enzyme, phosphotransferaw activity of which can equal or actually exceed that for hydrolysis. (iv) The very steep nature of activity-pH profiles for PP,-glucose and nucleotide-glucose phosphotransferase activities of the detergentexposed enzyme between pH 6 and 7 (see Fig. 9 ) . (v) The relatively high values for K,,, for various substrates, compared with physiological levels of these substrates in mammalian liver and kidney. For example, K , for glucose-6-P is approximately 1-2 mM compared with hepatic glucose-6-P levels of 0.05-0.13 m M (103, 166, 167); and K , for glucose equals approximately 80-90 m M compared with normal physiological values of 5 m M (9). Because of such variance in I<, values and normal physiological substrate levels, the enzyme may be especially susceptible to competitive inhibition by such metabolites as ATP [physiological levels in liver equals 4 mM (see reference IOS)] and the like (9). (vi) Responsiveness of the enzyme to such physiological manipulations as hormone administration or deprivation and nutritional variations (see Table XI and Figs. 2 and 9). (vii) Observed inhibition of all activities, competitive with respect to phosphate substrates, by P, (104), HCO,- (141), and citrate (15.2). (viii) Observed competitive inhibition, over a wide range of pH values, by PPi and various nucleosidetriphosphates and nucleosidediphosphates (103). (ix) Reduction in net glucose release via glucose-6-P phosphohydrolase activity by glucose acting as phosphoryl acceptor alternate with water (9,30, 31, 4). 162. M. C. Scrutton and M. F. Utter, Ann. Rev. Biochem. 37, 249 (1968). 163. C. Sharma, R. Manjeshwar, and S. Weinhouse, JBC 238, 3840 (1963). 164. M. Salas, E. Viiiuela, and A . Sols, JBC 238, 3535 (1963). 165. J. Salas, M. Salas, E. Viiiuela, and A. Sols, JBC 240, 1014 (1965). 166. H. G. Hers and H. de Wulf, in “Control of Glycogen Metabolism” (W. J. Whelan, ed.), p. 65. Academic Press, New York, 1968. 167. D. A. Young, ABB 114, 309 (1966).

22. GLUCOSE-6-PHOSPHATASE

595

(x) Observed activity-discriminating effects of a variety of factors which serve to alter selectively phosphohydrolase and phosphotransferase activities of the enzyme. Factors observed to exert such effects include:

a. Reaction pH. As described in detail in Section III,D,l, glucose-6-P phosphohydrolase activity of the enzyme is manifest over a broad range of pH both below and above neutrality, while PPi-glucose and nucleotideglucose phosphotransferase activities are most active below p H 7 (see Figs. 2 and 9 ) . Inhibition of glucose-6-P phosphohydrolase activity by these compounds, in contrast, persists a t alkaline as well as acid pH. Thus variations in pH within the cell may effectively alter the two types of activity discriminantly. Carbamyl-P, in contrast with PPi and nucleotides, is effective both as an inhibitor of phosphohydrolase activity and as a substrate for the phosphotransferase reaction at higher (> pH 7) as well as lower pH values (4.2, 43) (see Fig. 9 ) , an observation suggesting a further discrimination, even among phosphoryl donors, by cellular pH. The latter concept is particularly interesting from a teleological point of view. b. Long-Chain Fatty Acyl-CoA Esters. As described in Sections II,C,l and 3, these compounds a t “physiological levels” can activate transferase activity of the enzyme while a t identical levels inhibiting hydrolase activity. A marked shift in activity-pH profiles for phosphotransferase (but not phosphohydrolase) activities of the enzyme also is produced by these compounds (9, 10, 88).

c. Cetrimide in Concert with Phlorizin. As considered in detail in Section II,C,3, the cationic detergent cetrimide potentiates inhibition by phlorizin of phosphotransferase activity while this same compound of glucose-6-P enzyme. d. Detergent Potentiation of Inhibition b y Nucleosidetriphosphates and Nucleosidediphosphates and Pi of Phosphohydrolase Activity. Various synthetic and natural detergents markedly potentiate inhibition by ATP and other nucleotides of glucose-6-P phosphohydrolase activity. Inhibition by P, of phosphohydrolase (but not phosphotransferase) activity is likewise potentiated by detergents (104) (see also Sections II,C,3 and III,D,4,b). The significance of many of the above observations from the physiological point of view has been discussed in some detail elsewhere by the author (9).

596

ROBERT C. NORDLIE

IV. Metabolic Roles and Regulation, in Vivo

The metabolic roles and regulation of glucose-6-P phosphohydrolase activity have been considered in detail in reviews by Cahill et al. (6)and Ashmore and Weber ( 7 ) . More recently, possible metabolically important roles for phosphotransferase activities of this enzyme in liver, kidney, and intestine have been described in reviews by the present author (9, 10)-who also considered a variety of regulatory features based on interaction of substrates, inhibitors, and activators with the multifunctional enzyme-and by Cohn et al. (11). The prime function of this enzyme classically has been considered to be the controlled release, through hydrolysis of glucose-6-P formed by glycogenolytic and gluconeogenic processes, of free glucose (6).Net glucose liberation from gluconeogenic tissues (liver and kidney) was believed controlled through opposing actions of glucose phosphorylation via hexokinase and/or glucokinase and glucose release through glucose-6-P phosphohydrolase action (6) (see Fig. 1). A function for the enzyme in releasing glucose from meningeal glycogen for utilization by the brain during periods of extreme physical stress has also recently been suggested by Rovainen (168). The recent discovery of the dual nature of the enzyme, involving potent synthetic as well as hydrolytic activities, has served as a basis for a variety of new control mechanisms and physiological roles for the catalyst. For example, hydrolytic activity of the enzyme is limited through competitive inhibition by PP,, carbamyl-P, Pi, HC0,-, ATP, ADP, and various other phosphoanhydrides. Glucose itself, through participation as phosphoryl acceptor alternative with water (30,31, @), may regulate its own release in an autocatalytic manner (9, 10, 169). Net synthesis of glucose-6-P through phosphotransferase activity of the enzyme may also be important, a t least in certain metabolic situations. The establishment of new, higher than normal plateau levels of blood glucose in untreated diabetes, for example, may be accomplished by the replacement of insulin-dependent liver glucokinase [ K , for glucose = 1040 mM (170,171)] with phosphotransferase activity of glucose-6168. C. M. Rovainen, Science 167, 889 (1970). 169. E.F. McCraw, M. J. Peterson, and J. Ashmore, Proc. SOC.Exptl. Bid. M e d . 126, 232 (1967). 170. D. L. DiPietro, C. Sharma, and S. Weinhouse, Biochemistry 1, 455 (1962). 171. E.Viiiuela, M.Salas, and A. Sols, JBC 238, PC1175 (1963).

22. GLUCOSE-6-PHOSPHATASE

597

a)]

phosphatase [ K , for glucose = 80 mM (37, ( 4 1 ) . The latter activity increases significantly in diabetes and is further favored by hyperglycemia, dehydration, acidosis, and possibly also by increased levels of long-chain acyl CoA esters (88) characteristic of this condition. Experimental observations supporting such a role for PPi-glucose phosphotransferase activity in livers of diabetic, fasted, and fructose-refed fasted animals have been described (41, 169, 172, 173). For example, PP, produced in the UDP glucose synthetase reaction (see Fig. 1) may, in such a fashion, participate further in the conversion of glucose to glycogen (172). Scott and Jones (48) have demonstrated that bovine pancreas islet cell glucose-6-phosphatase catalyzes phosphotransferase reactions and suggested that both hydrolytic and synthetic activities of the enzyme may, under appropriate conditions, contribute to net glucose concentrations in the pancreas and hence be instrumental in regulating insulin release from /3 cells. Possible roles for hydrolytic and synthetic activities of the enzyme in glucose transport have been considered in Section I1,D. A. CONTROL OF ENZYMIC ACTIVITIES,in Vivo Both hydrolytic and synthetic activities of the enzyme are highly responsive to a variety of hormonal, dietary, and other factors, in vivo. In 1959, Ashmore and Weber ( 7 ) published a comprehensive report on the effects of such factors in the regulation of hepatic glucose-6-P phospliohydrolase activity. Weber has also summarized interesting, more recent work in several volumes of “Advances in Enzyme Regulation” (174). The present author has drawn upon the review of Ashmore and Weber (7) in preparing Table XI in which responses of the enzyme to various hormonal, dietary, and other in vivo factors are briefly summarized. I n addition, the results of certain recent studies, in which responses of both synthetic and hydrolytic activities of the enzyme to such treatments and conditions were Considered, are also included along with pertinent literature references. Of particular interest with respect to in vivo regulation of the enzyme 172. B. Friedmann, E. H. Goodman, Jr., and S. Weinhouse, Endocrinology 81, 486 (1967). 173. R. D. Hornichter and J. Brown, Diabetes 18, 257 (1969). 174. G. Weber, ed., “Advances in Enzyme Regulation,” Vols. 1, 2, 3, and 5. Pergamon Press, Oxford, 1963-1967.

598

ROBERT C. NOFLDLIE

TABLE X I TO HORMONAL, DIETARY, RESPONSESOF ACTIVITIESOF GLUCOSE-~-PHOSPHATASE A N D OTHERFACTORS, in Vivo Treatment or factor

Response noted

Referencesn

I. Responses of Phosphohydrolase and Phosphotransferase Increase in both normal and (7,72,80, 93-96, 176) Glucocorticoids Adrenalectomy Diabetes (alloxan-induced) Adrenalectomy of diabetic animals Insulin administration Growth hormone

Acute fasting High-maltose diet High-fructose diet Developing fetus

Newborn Type I glycogenosis

adrenalectomized animals Decrease Increase

(7,93, 113) (7,41,7.9,60,go, 96, 99, 113,117,118,176, 176) Activity increased to normal (7,113)

Decrease in both normal (7,4l,95, 113) and diabetic Decrease in hypophysec(116) tomized rats Increase or no change in normal animals Increase Decrease Increase Activity very low; detectable at 18 days in rat fetus Present very early in chick embryo Marked increase a t birth Markedly decreased

11. Response of Glucose-6-P Phosphohydrolase Hypophy sectomy ACTH Glucagon Thyroxine 3',5'-Cyclic AMP or the dibutyryl derivative High-protein diet High-fat diet High-galactose diet Protein-free diet Regenerating liver Tumors Induced carcinogenesis (liver enzyme)

Decrease Increase Increase Increase Increase, fetus only Increase Increase Increase No change No change Decreased or absent Decreased

0 Reference 7 alludes to the review by Ashmore and Weber, which contains many primary references.

22.

GLUCOSE-6-PHOSPHATASE

599

have been the results of some recent studies (71,80, 93, 95, 98,99, 176) which indicate that responses of activities of the enzyme to diabetes or fasting are accentuated when assayed in the presence of supplemental detergent while alterations in responses to glucocorticoid therapy are markedly ameliorated or totally abolished by such treatment of microsoma1 preparations (see Fig. 2). Basic mechanistic differences in modes of response of activities of the enzyme to glucocorticoid therapy in contrast with experimental diabetes or fasting are thus indicated.

B. A FINALSPECULATION It previously has been pointed out that, potentially, PP,-glucose phosphotransferase and ATP-glucose phosphotransferase activities of glucose6-phosphatase are the most potent glucose phosphorylating systems which have been characterized for liver (9,10, 41, 118).Such a conclusion appears to have possible validity principally a t and below pH 7 however (see Fig. 9) because of the nature of the pH-activity profiles of the phosphatase-associated phosphotransferase activities. New and rather exciting possibilities relating to the regulation of, and regulatory role for, this enzyme have been raised by the recent observations of Lueck and Nordlie (42, 49),who have observed potent phosphotransferase activity with carbamyl-P-an established, metabolically important intermediate-over a wide range of pH values. As indicated by data in Fig. 9, this kinaselike activity markedly exceeds that of both hepatic glucokinase and hexokinase in ability to phosphorylate glucose at any pH value studied (pH 5-8). It is the author’s view that synthetic activity of this unique enzyme may well prove to be equally as significant as its more familiar phosphohydrolase activity. The generation of metabolically active pools of glucose-6-P, in certain cellular compartments (181) , for further participation in specialized biosynthetic pathways may well prove to involve phosphorylative action of this enzyme.

175. R. C. Nordlie, T. L. Hanson, and W. J. Arion, Abstr. 7th Intern. Congr. Biochem., Tokyo, 1967 Abstr. G-51. Sci. Council Japan, Tokyo, 1968. 176. P. N. Pandhi and H. Baum, Nature 218, 1324 (1967). 177. 0. Greengard, Science 183, 891 (1969). 178. W. B. Anderson, R. N. Horne, and R. C. Nordlie, Biochemistry 7, 3997 (1968). 179. 0. Greengard, BJ 115, 19 (1969). 180. P. J. Collip, S. Y. Chen, and M. Halle, BBA 187, 141 (1968). 181. D. F. Heath, BJ 110, 313 (1968).

600

ROBERT C. NORDLIE

Appendix

Information on the distribution of glucose-6-phosphatase in tissues and various taxonomic groups is given in Tables XII-XVII, which are relevant to the discussion in Section I,B,l. Because of the extremely large body of literature relating to the presence of the activity in certain tissues (for example, rat liver) an exhaustive citation of all pertinent literature references here is physically impossible. Accordingly, references on the occurrence of enzymic activity in any given tissue have arbitrarily been limited to a maximum of five. The actual number of citations coming to the author’s attention during a systematic survey of the literature since 1949 is also indicated in certain tables. TABLE XI1 REPORTED PRESENCE OF GLUCOSE-~-PHOSPHATASE IN VERTEBRATE LIVER Vertebrate source Mammals Rat Mouse Human Bovine Guinea pig Sheep Rabbit Dog Pig Ground squirrel Cat Monkey Birds Chicken Duck Pigeon Other Fish Frog Lamprey

References (16,17,19,$1, SY,38) (182-1 86) (70,116,116,168,187) (76,76,78,188, 189) (21,190-1 93) (194-198) (24, 198-2001) (74,202, 20s) (771204) (204 (198)

Total No. of reports noted 420 37 49 6 20 7 30 8 2 1 1 2 20 1 2

(214-216) (817-2800) (9211

5 6 2

182. K. H. Shull, J. Ashmore, and J. Mayer, ABB 62,210 (1956). 183. K. H. Shull, G. F. Cahill, Jr., E. L. Gadsden, and J. Mayer, JBC 222, 415 (1956). 184. J. P. Turchini and P. Malet, Ann. Histochim. 9, 331 (1964). 185. A. Zorzoli, J . Gerontol. 17, 359 (1962).

22. GLUCOSE-6-PHOSPHATASE

601

186. C. C. Griffin, V. S. Waravdekar, B. F. Trump, P. J. Goldblatt, and R. E. Stowell, Am. J . Pathol. 47, 833 (1965). 187. S. Auricchio and N. Rigillo, Biol. Neonatorum CN. S.1 2, 146 (1960). 188. J. C. Bartley, R. A. Freedland, and A. L. Black, Am. J . Vet. Res. 27, 1243 (1966). 189. E. J. H.Ford, J . Comp. Pathol. Therap. 71, 60 (1961). 190. A. M. Nemeth, JBC 208, 773 (1954). 191. M.A. Lea and D. G. Walker, BJ 85, 30P (1962). 192. M. A. Lea and D. G. Walker, BJ 91, 417 (1964). 193. E. Degkwitz, D. Luft, U. Pfeiffer, and H. Sbudinger, 2. Physwl. Chem. 349, 465 (1968). 194. S. R.Wagle and P. Nelson, BBA 121, 190 (1966). 195. E. J. H.Ford, J . Agr. Sci. 59, 67 (1962). 196. F. Raggi, D. S. Kornfeld, and M. Kleiber, Proc. SOC. Ezptl. Bwl. Med. 105, 485 (1960). 197. F.J. Ballard and I. T. Oliver, BJ 95, 191 (1965). 198. M. J. R. Dawkins, Brit. Med. Bull. 22, 27 (1966). 199. M. Flint, G. H. Lathe, T. R. Ricketts, and G. Silman, BJ 75, 1OP (1960). 200. V.S.Il’in and N. G. Stepanova, Vop Med. Khim. 10, 576 (1964). 201. J. Jonek and Z. Olkowski, Acta Histochecm. 20, 103 (1965). 202. G. Cahill, Jr., S. Zottu, and A. S. Earle, Endocrinology 60, 265 (1957). 203. S. Pontremoli and M. Orunesu, Boll. SOC.Ital. B i d . Sper. 32, 760 (1956). 204. C. R. C. Heard, Diabetes 15, 78 (1966). 205. R. F. Burlington and G. J. Klain, Comp. Biochem. Physiol. 22, 701 (1967). 206.K.I. Shanygina, Vopr. Med. Khim. 12, 258 (1966). 207. L. G. Leibson, E. M. Plisetskaya, and L. G. Ogorodnikova, Dokl. Akad. Nauk SSSR 153, 240 (1963). 208. J. K. Pollak and C. D. Shorey, Develop. Bwl.17, 536 (1968). 209. M. T.Rinaudo, Boll. SOC.Ital. Biol. Sper. 35, 2157 (1959). 210. S.S.Simbonis and R. A. McBride, Develop. Bwl. 12, 347 (1965). 211. G. S. Kilsheimer, G. R. Weber, and J. Ashmore, Proc. SOC.Ezptl. Biol. Med. 104, 515 (1960). 212. S.-M. Ting, T’ai-wan I Hsueh Hui Tsa Chih 66, 149 (1967); C A 68, 10707d (1968). 213. R. Bruno, M. T. Rinaud, and C. Giunta, Boll. SOC.Ital. Bwl. Sper. 44, 1640 (1968). 214. H.Noda, Mie. Kenritsu Daigaku Suisan Gakubu Kiyo 7, 65 and 73 (1967); C A 68, 1063Oy (1968). 215. P. M. Orkand, Am. J . Med. Technol. 28, 296 (1962). 216. P. A. Janssens, Comp. Bbchem. Physiol. 16, 317 (1965). 217. N. Sonnewhein and M. J. Kopac, J . Cellular Comp. Physiol. 45, 361 (1955). 218. V. A. Galton and 5.H. Ingbar, Endocrinology 70,622 (1962). 219. E.Frieden and H. Mathews, ABB 73, 107 (1958). 220. A. E.Ferreri and M. Goria, Uen. Comp. Endocrinol. 3, 378 (1963). 221. E. M. Plisetskaya and L. G. Ogorodnikova, Zh. Evolyutsionnoi Biokhim. i Fiziol. 3, 304 (1967).

602

ROBERT C. NORDLIE

TABLE XI11 REPORTEDPRESENCE OF GLUCOSE-~-PHOSPHATASE IN VERTEBRATE KIDNEY Vertebrate source Mammals Rat Mouse Human Guinea pig Sheep Rabbit Dog Monkey Other Chicken Frog Mud PUPPY Newt

References (46, 101, 146, 222, 223) (223-226) (226~-229) (229-232)

Total No. of reports noted 33 4

9 4

($29) (229, 230,233-236) (236-238) (239)

2 9 3

(240) (24411 (242) (243)

3 1 1 1

1

222. G. Weber and A. Cantero, Cancer Res. 15, 105 (1955). 223. A. D. Chiquoine, J. Histochem. Cytochem. 1, 429 (1953). 224. A. Zorzoli, Develop. Biol. 17, 400 (1968). 225. 0. V. Deimling, G. Baumann, and H. Noltenius, Histochemie 5, 1 (1965). 225a. S. Ritchie and D. Waugh, Am. J. Pathol. 33, 1035 (1957). 226. P. Pakdaman, A. A. Skin, and W. A. Milner, J. Urol. 87, 309 (1962). 227. H. Tanaka, Acta Pathol. Japon. 12, 177 (1962). 228. B. Ivemark, Acta Pathol. Microbwl. Scand. 45, 1 (1959). 229. M. J. R. Dawkins, Nature 191, 72 (1961). 230. M. Wachstein and M. Bradshaw, J. Histochem. Cytochem. 13, 44 (1965). 231. N. S. Shah, L. E. Fox, and S. P. Martin, Ezperientia 22, 648 (1966). 232. N. S. Shah, J. F. Monroe, L. E. Fox, and S. P. Martin, Life Sci. 6, 1733 (1967). 233. M. T. Lutsenko, Tr. Blagoveshch. Gos. Med. Inst. 8, 155 (1966); CA 66, 93224b (1967). 234. V. Ferioli and F. Fiaccadori, Ateneo Pannense 32, 20 (1961); CA 56, 4005b (1962) 235. M. Wachstein and K. Lange, Am. J . Pathol. 34, 835 (1958). 236. W. P. McCann, Proc. Soc. Exptl. Biol. Med. 124, 185 (1967). 237. M. Nagano, A. Heidland, K. Kluetsch, and H. Horchrein, Klin. Wochschr. 41, 605 (1963). 238. E. Pausescu, F. Negrea, and R. Florescu, Studii Cercetan' Fiziol. 10, 157 (1965); CA 63, 16870~(1965). 239. E. F. Deig and L. P. Gebhardt, 2.Naturjorsch. 18b, 903 (1963). 240. M. T. Rinaudo and L. Galletti, Gwrn.Biochim. 15, 383 and 396 (1966); CA 67, 30230a (1967). 241. G. T. Adunts and I. G. Aslanyan, Vopr. Biokhim., Akad. Nauk Arm. SSR 3, 133 (1963); CA 62, 3273s (1965). 242. S. R. Himmelhoch and M. J. Karnovsky, J . Bwphys. Bwchem. Cytol. 9, 893 (1961). 243. E. Ferreri and A. Peyrot, 2. Zelljorsch. Mikroskop. Anat. 56, 470 (1962); CA 61, 16491f (1964).

22.

603

GLUCOSE-6-PHOSPHATASE

TABLE XIV REPORTED PRESENCE OF GLUCOSE-~-PHOSPHATASE IN VERTEBRATE SMALLINTESTINE Vertebrate mwce

References

Total No. of reports noted

Mammals

Rat Mouse Human Guinea Pig Sheep Rabbit Monkey Other Chicken

2

244. J. M. Wrigglernorth and W. F. R. Pover, Intern. J . Radiation Biol. 1% 243 (1967). 245. K. Kozlowska, Folk Morphol. 24, 251 and 421 (1965); C A 64,10051b (1966). 246. S. Maeda, Zgaku To Seibutsugaku 31, 52 (1954); C A SO, 9468g (1956). 247. P. A. Ockerman, BBA 105, 22 (1965). 248. J. Jos, J. Frezal, J. Rey, and M. Lamy, Ann. Hktochim. 12, 165 (1967). 249. J. Jos, J. Frezal, J. Rey, and M. Lamy, Pedkt. Res. 1, 27 (1967). 250. J. B. Field, S. Epstein, and T. Egan, J. CZin. Invest. 44, 1240 (1965). 251. H. E. Williams, P. L. Johnson, L. F. Fenster, L. Laster, and J. B. Field, Metab., Clin. Exptl. 12, 235 (1963). 252. J. James and L. L. Salomon, Federation Proc. 23, 533 (1964). 253. G.Hubscher, G.R. West, and D. N. Brindley, BJ 97, 629 (1965). 254. D. Tournaire, P. Bastide, and G. Dastugue, C m p t . Rend. SOC.BWZ. 160, 1597 (1966). 255. T. H. Kent, H. R. Jervis, and J. G. Kuhns, Am. J . PathoZ. 48, 667 (1966). 256. D.L.Baxter-Grillo, Histochemie 19, 31 (1969).

604

ROBERT C. NORDLIE

TABLE XV REPORTED PRESENCE OF GLUCOSE-~-PHOSPHATASE IN VERTEBRATE PANCREAS ~

Vertebrate source Mammals Rat Mouse Human Bovine Guinea pig Rabbit Dog Cat Horse Hamster Other Duck Chicken

References

(266a-269) (2666~2, 269-262)

(267, 263) (48) (191, 192, 266a, 269,264) (269, 264-267)

(264)

Total No. of reports noted 5 11

2 2 9 8

(268)

1 1 1

(266a)

1

(269)

1 1

(266)

($70)

256a. I. B. Taljedal, BJ 114, 387 (1969). 257. P. Petkov, J. Verne, and R. Wegmann, Ezperientiu 21, 530 (1965). 258. T. A. I. Grillo, J. Endocrinol. 31, 67 (1964). 259. R. B. Cohen and H. J. Wolfe, J. Histochem. Cytochem. 11, 288 (1963). 260. B. Hellman and C. Hellerstrom, Acta Endocrinol. 39, 474 (1962). 261. S. J. H. Ashcroft and P. J. Randle, Nature 219, 858 (1968). 262. M. Nakamura and K. Yamada, Z . Zellforsch. Mikroskop. Anat. 66, 396 (1965). 263. T. A. Grillo, J. Endocrinol. 36, 151 (1966). 264. S. S. Lazarus, Proc. SOC.Exptl. Bwl. Med. 101,819 (1959). 265. S. S. Lazarus and H. Barden, J. Histochem. Cytochem. 12, 792 (1964). 266. P. E. Petkov, Ann. Histochim. 11, 79 (1966). 267. N. Ihara, Endocrinol. Japon 12, 215 (1965) ; C A 64, 1642431 (1965). 268. N. Bjorkman, C. Hellerstrom, B. Hellman, and U. Rothman, 2. Zellforsch. Mikroskop. Anat. 59, 535 (1963). 269. C . Hellerstrom, 2.Zellforsch. Mikroskop. Anat. 60,688 (1963). 270. T. A. Grillo, Folia Histochem. Cytochem. 1, 453 (1963); C A 62, 8211d ( 1965).

605

22. GLUCOSE-6-PHOSPHATASE TABLE XVI REPORTED PRESENCE OF GLUCOSE-6-PHOSPHATASE Tissue Blood Serum Erythrocytes

Leukocytes Thrombocytea Brain Cerebellum Cerebral cortex Sympathetic ganglion neurons Sertoli cells Spinal ganglion Neurohypophysis Adrenal cortex

Muscle, skeletal Muscle, heart Spleen Bone marrow Tibia1 cartilage Cartilage Heart valve tissu Mammary glands Mammillary buds Skin Esophagus and trachea Mural endocardium Periosteium and periodontium Uterus, cervix, vagina, sexual skin Endometrium Myometrium Ovary

IN

MISCELLANEOUS TISSUES"

Organism Man Rat Man Rabbit Rabbit Pig Various mammals and birds Man Man Rat, guinea pig, rabbit Rat Rat Rat Rat, guinea pig, man Rat Rat and frog Rat Rabbit Marmot Guinea pig Man Pig Rat Man Rat Rat Man Man Bovin Rat Rabbit Human Chick Human Guinea pig Guinea pig Human Human

References

ROBERT C. NORDLIE

TABLE XVI (Continued) Tissue Placenta Vas deferens Glandus inguinal8 Seminal vesicle, coagulating gland, dorsolateral prostate gland Testicles Spermatozoa Oocytes Milk fat globule membranes Adipose tissue Olfactory bulb Eye, cornea retina

Organism

References

Rat Human Crayfish Rabbit Rat

Rats, mice, guinea pigs Sea urchin Rat and mouse Rat Rat Bovine Bovine

Mammalian tissues other than those considered in Tables XII-XV. 271. E. Eggermont and H. G. Hers, Clin. Chim. Acta 5, 774 (1960). 272. J. Litwin, S. Szmigielski, and B. Zupanska, Polskie Arch. Med. Wewnetrznej 35, 69 (1965) ; C A 63, 3433g (1965). 273. M. Thomas and W. N. Aldridge, BJ 98, 94 (1966). 274. S. Sartori, L. Valagussa, L. Mainiteri, and L. Petronio, Haematologica 49, 11 (1964). 275. I . N. Vovk and B. A. Pavlov, Ukr. Bwkhim. Zh. 37, 331 (1965); C A 63, 11914 (1965). 276. T. Fossa, Boll. Sac. Ital. Biol. Sper. 39, 1197 (1963). 277. D. Micu, M. Zamfirescu-Gheorghiu, E. Mihailescu, S. Maximilian, and C. Vladescu, Studii Cercetari Med. Interna 6, 279 (1965) ; C A 63, 15434~ (1965). 278. A. Englhardt-Goelkel, Klin. Wochschr. 42, 1141 (1964). 279. F. Linneweh, G. W. Lohr, H. D. Wallel, and R. Gross, Enzymol. Biol. Clin. 2, 188 (1962). 280. R. Horinouchi, Osaka Daigaku Igaku Zasshi 11, 3729 (1959); C A 54, 15612e ( 1960). 281. H. B. Tewari and G. H. Bourne, J . Histochem. Cytochem. 11, 121 (1963). 282. H. B. Tewari and G. H. Bourne, Exptl. Cell Res. 28, 444 (1962). 283. A. Charbonnier and R. Wegmann, Congr. Intern. Assoc. SOC. Natl. Europeennes Mediten. Gastro-Enterol., 7th, Brussels, 1964 Vol. 1 p. 261 (1964); C A 61, 4847g (1964). 284. K. G. Prasannan and K. Subrahmanyam, Endocrinology 82, 1 (1968). 285. N. N. Sharma, Acta Histochem. 26, 278 (1967). 286. A. Petreseu and M. Alexianu, R e v . Roumaine Neural. 2, 321 (1965). 287. 0. Vilar, M. I. Perez del Cerro, and R. E. Mancini, Ezptl. Cell Res. 27, 158 (1962). 288. H. B. Tewari and G. H. Bourne, J . Histochem. Cytochem. 10, 42 (1962).

22. GLUCOSE-6-PHOSPHATASE

607

289. E. Legait, H. Legait, C . Burlet, and A. Burlet, Compt. Rend. SOC.BWl. 160, 1659 (1966). 290. M. Kool and A. Y. Truupyl'd, Tsitologiya 10, 1064 (1968); C A 69, 94228e (1968). 291. R. Hilf and F. F. Burnett, ABB 104, 106 (1964). 292. F. Ghiringhelli, G. Gereeli, and E. Mira, Folia Endocrinol. 15, 823 (1962); CA 59, 10515d (1963). 293. R. Wegmann and H. Khoarovchahi, Ann. Histochim. 9, 57 (1964). 294. M. Jonadet, J. P. Turchini, and P. Bastide, Compt. Rend. SOC.Bwl. 162, 1802 (1969). 295. P. J. Meusers, Histochmie 7, 50 (1966). 296. G. Mekanik, R. L. Smith, and R. M. MacLeod, Metab., Clin. Exptl. 15, 641 (1966). 297. S. Larsson, T. Nilsson, and B. Olsson, Acta Vet. Scand. 7, 47 (1966). 298. C. Van den Hende, E. Muylle, and W. Oyaert, Zentr. Veteriinuermed. 15, Part A, 135 (1968). 299. L. Carbini, A. Casula, and G. Liguori, Boll. SOC.Ital. BWl. Sper. 42, 1085 ( 1966). 300. B. Zupanska, S. Samigielski, J. Litwin, and J. Zielinski, Arch. Zmmunol. Therap. Ezptl. 14, 356 (1966). 301. G. Tota and R. Basile, Studi Sassaresi, Sez. II 43, 591 (1965) ; CA 65, 20645.e (1966). 302. C. Bona, V. Stanescu, M. Dumitrescu, G. Chyka, and V. Ionescu, Acta Histochem. 21, 98 (1965). 303. C. Bona, V. S'tanescu, and V. Ionescu, Acta Hktochem. 21, 284 (1965). 304. E. J. Eyring, C. E. Anderson, and J. Ludowieg, Arthritis Rheumat. 6, 208 (1983). 305. A. Delbrueck, Klin. Wochschr. 41, 488 (1963). 306. R. Narayanan and N. C. Ganguli, Indian J. Biochem. 2, 240 (1965). 307. K. Wallenfels, H. D. Summ, and W. Creutzfeldt, Deut. Med. Wochschr. 82, 1581 (1957). 308. S. Paek, H i s t o c h a k 13, 29 (1968). 309. Y. Nakajima and K. Kishi, Bull. Tokyo Med. Dental Univ. 14, 279 (1967); CA 68, 37319r (1968). 310. Y. Miura and W. C. Lobita, J. Invest. Dermatol. 42, 115 (1964). 311. G. Bonu and G. Zina, Dermatologica 111, 349 (1955). 312. 0. Braun-Falco, Dermatol. Wochschr. 134, 1252 (1956). 313. G. W. Hinsch, J. Morphol. 119, 327 (1966). 314. B. J. P. Becker, J. Pathol. Bacterial. 88, 541 (1964). 315. E. H. Charreau, J. A. Kofoed, and A. B. Houssay, Arch. Oral BWZ. 11, 709 (1966). 316. M. H. Burgos and G. B. Wislocki, Endocrinology 59, 93 (1956). 317. B. Goldberg and H. W. Jones, Obstet. Gynecol. 4, 426 (1954). 318. I. Ricciardi, A. Sallusto, and M. Torella, Boll. SOC.Ital. Biol. Sper. 40, 6 (1964). 319. V. Danesino, I. Ricciardi, and A. Sallusto, Arch. Obstet. G y e c o l . 67, 513 (1962). 320. J. Koudstaal, B. Bossenbroek, and M. J. Hardonk, European J. Cancer 2, 313 (1966). 321. H. A. Padykula and D. Richardson, Am. J. Anat. 112, 215 (1963).

608

ROBERT

C. NORDLIE

322. I. Ricciardi, A. Sallusto, and M. Malato, Arch. Ostet. Ginecol. 68, 35 (1963). 323. P. Curaen, J . Obstet. Gynaecol. Brit. Commonwealth 71, 388 (1964). 324. P. Curzen, J . Obstet. Gynaecol. Brit. Commonwealth 74, 385 (1967). 325. R. G. Kessel, W. R. Panje, and M. L. Decker, J . Ultrastruct. Res. 27, 319 (1969). 326. W. Kuhnel and K. Wrobel, Histochemie 8, 315 (1967). 327. J. P. Fouquet, Ann. Histochim. 9, 163 (1964). 328. P. B a d , H. G. Goslar, and E. Tonutti, Acta Histochem. 26, 343 (1967). 329. W. A. Anderson, J . Ultrastruct. Res. 25, 398 (1968). 330. A. M. Dalcq, Arch. Biol. 77, 205 (1965). 331. R. M. Dowben and D. E. Philpott, BBA 135, 1 (1967). 332. G. Weber, G. Banerjee, and J. A s h o r e , BBRC 3, 182 (1960). 333. K, Nandy and G. H. Bourne, Acta Histochem. 23, 86 (1966). 334. N. N. Sharrna, Acta Hhtochem. 27, 165 (1967). 335. M. Jonadet, P. Bastide, and G. Dastugue, Pathol. Biol., Semaine Hop. [N. S.l 15, 271 (1967). 336. E. Cameron and D. F. Cole, Exptl. Eye Res. 4, 62 (1965). 337. P. Bastide, P. Tronche, and G. Dastugue, Compt. Rend. SOC.Biol. 160, 328 ( 1966).

22.

609

GLUCOSE-6-PHOSPHATASE

TABLE XVII REPORTED PRESENCE OF GLUCOSE-~-PHOSPHATASE I N MISCELLANEOUS NONMAMMALIAN ORGANISMS Source

References

Plants Pisum arvense, cotyledons and sieve elements Cactus (Echirwpsis), stalk Panicoid and festucoid grasses, root epidermis Phuseolus vulgaris (common kidney bean), cotyledon Robinia pseudoacacia, phloem sap Pea seeds Opuntia $ma-indiea, chlorophyll-bearing parenchyma Microorganisms Escherichia wli Balantidium wli Aspergillus oryzae Aspergillus awammi, variety kawachi ‘‘Yeast cells” Fwarium oxysporum, F. semitectum, F. semitectum, F. sporotrichiella, F. gabbosum, and F. moniliforme (microsomal fraction) Toxoplasma gondii, proliferative phase Miscellaneous Hydra Sciera wprophila (a gnat), salivary gland larvae Eisenia foetida (an earthworm), nervous system

338. A. M. Flinn and D. L. Smith, Planta 75, 10 (1967). 339. M. Aiazzi, Boll. Soc. Ztal. Bwl. Sper. 38, 290 (1962). 340. C. J. Avers, Ezptl. Cell Res. 16, 692 (1959). 341. H.Wanner, Ber. Schweiz. Botan Ges. 63,201 (1953). 342. D.H.Turner and J. F. Turner, BJ 74, 486 (1960). 343. M. A. Satta and A. Sisini, Boll. SOC.Ital. Bwl. Sper. 40, 1109 (1964). 344. A. Sisini and A. Satta, Ztal. J . B w c h m . 15, 407 (1966). 345. P. Mitchell, Biol. Struct. Function, Proc. IUB/ZUBS Intern. Sump., lst, Stockholm, 1960 Vol. 2, p. 581. Academic Press, New York, 1961. 346. N. N. Sharma and G. H. Bourne, Acta Histochem. 17, 293 (1964). 347. S.Ueda, Nippon Nogei Kagaku Kaishi 38, 281 (1964); C A 63, 7268f (1965). 348. L. Kiesow and S. Doege, 2.Naturforsch. 16b, 576 (1961). 349. L. D.Varbaneta, 0. Y. F h h b a , and E. Z. Koval, Mikrobwl. Zh. (Kiev) 29, 491 (1967); C A 68,93779~(1968). 350. M. Glowinski and T. I(.Niebroj, Acta Parasitol. Polon. 13, 399 (1965); B A 48, 4894 (1967). 351. T. L. Lentz and R. J. Barrnett, J. Ezptl. Zool. 150, 103 (1962). 352. J. Y. Terner, R. M. Goodman, and D. Spiro, J . Histochem. Cytochem. 13, 168 (1965). 353. R. M. Goodman, J. Y. Terner, and H. V. Crouse, J. Cell Bwl. 23, 37A (1964).

610

ROBERT C. NORDLIE

354. R. M. Goodman, J. Y. Terner, and D. Spiro, Exptl. Cell Res. 49, 504 (1968). 355. I. Vigh-Teichmann and H. G. Goslar, Hktochemie 14, 352 (1962).

ACKNOWLEDQMENTS The author is indebted to Drs. J. K. Pollak and D. B. M. Scott for generously providing valuable information, some of it prior to publication; to Drs. E. A. Stadtman and H. A. Lardy for stimulating discussions; to his graduate students and postdoctoral associates, past and present, who carried out many of the studies discussed; and to Miss Hilda Klein, without whose diligent and persistent aid in the survey of the literature this chapter would not have been completed.

Fructose.1, 6.Dif hosfihatases S. PONTREMOLI

B . L . HORECKER

I. Introduction . . . . . . . . . . . . A. Historical Review . . . . . . . . . B . Regulation and Physiological Function . . . . C . Methods of Assay and Mechanism of Action . . . I1. Liver FDPase A . Purification and Properties . . . . . . . B. Changes in Properties Induced by Proteolysia . . C . Regulation of Catalytic Activity by AMP . . . D . Activation by Chemical Modification of SH Groups E. Activation by Disulfide Exchange . . . . . F. Molecular Structure of Rabbit Liver FDPase . . I11. Kidney FDPase . . . . . . . . . . . A Purification and Properties . . . . . . . B. Regulation of Kidney FDPase . . . . . . IV . Muscle FDPaae . . . . . . . . . . . A . Evidence for the Presence of the Enzyme in Muscle B. Purification and Properties of FDPase from Rabbit Muscle . . . . . . . . . . C. Structure and Relation to Liver and Kidney FDPase . D . Physiological Role of Muscle FDPase . . . . V. Fructosediphosphatase of Candida utilis . . . . . A . Purification and Properties . . . . . . . B. Inhibition by AMP . . . . . . . . C. Structure of the Purified Cundiala F D P w . . . D . Relation to Cundida SDPase . . . . . . VI . FDPaaes in Other Microorganisms . . . . . . . A . The Specific FDPase of Escherichiu coli . . . . B. Other Bacterial FDPases . . . . . . . C . FDPase in Slime Molds . . . . . . . D . Regulation of FDPase in Succhu~omycescerevkceae mdC.utilk. . . . . . . . . .

.

. . .

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

.

.

611

. . .

. . .

.

. .

.

.

.

612 612 613 615 616 616 618 618 621 622 626 629 629 630 632 632

. .

632 633 634 635 635 636 637 638 638 638 639 640

.

640

.

. .

. . . . . .

612

S. PONTREMOLI AND B. L. HORECKER

.

.

.

. . .

,

.

VII. FDPases in Higher Plants and Blue-Green Algae A. Purification and Properties B. Physiological Role of Plant FDPases C. Regulation of Plant FDPases .

. . . . . . . . . . . . . . . . VIII. Summary and Conclusions . . . . . . . . . A. Physiological Role of FDPases . . . . . . B. Comparative Properties of FDPases

.

. . . . . .

640 640 642 643 644 644 645

1. Introduction

A. HISTORICAL REVIEW The presence in mammalian liver of a specific phosphatase which catalyzes the hydrolysis of fructose l16-diphosphate ( 1 ) was first reported by Gomori in 1943 ( 6 ) . He succeeded in separating the enzyme from other phosphatases present in mammalian tissues and thus clarified much of the confusion which had previously existed regarding the specificity of these phosphatases. The specific fructosediphosphatase (FDPase) was shown to require a divalent cation such as Mg2+and to be inactive a t acid or neutral pH. It was present in the livers and kidneys of a number of mammalian species. Phosphatases are generally considered to be catabolic enzymes, and the role of the specific FDPase in carbohydrate metabolism was not recognized until many years after Gomori’s original discovery of the enzyme. It was shown to specifically hydrolyze the 1-phosphate group of FDP to yield fructose 6-phosphate (3, 4 ) , and Hers and Kusaka ( 5 ) clearly established its role in the metabolism of fructose in mammalian liver (Fig. 1). This pathway was confirmed by Leuthardt and his coworkers (6). A specific role of FDPase in gluconeogenesis was suggested by McGilvery and his co-workers (7)’ who found that conditions which favored gluconeogenesis resulted in increased levels of the enzyme in rabbit liver. At the time of the symposium organized by McGilvery and 1. The following abbreviations have been employed : FDNB, 2,4-fluorodinitrobenzene ; F6P, fructose 6-phosphate ; FDP, fructose 1,64iphosphate ; FDPase, fructose-l,6diphosphatase ; NEM, Nethylmaleimide ; PFK, phosphofructokinase ; PLP, pyridoxal phosphate; SDP, sedoheptulose 1,7diphosphate ; SDS, sodium dodecyl sulfate. 2. G. Gomori, JBC 148, 139 (1943). 3. J. Roche and S. Bouchilloux, Bull. SOC.Chim.B i d . 32, 739 (1950). 4. B. M. Pogell and R. W. McGilvcry, JBC 197, 293 (1952). 5. H. G. Hers and T. Kusaka, BBA 11, 427 (1953). 6. F. Leuthardt, E. Testa, and H. P. Wolf, Helv. Chim. Acta 36, 227 (1953). 7. L. C. Mokrasch, W. D. Davidson, and R. W. McGilvery, JBC 222, 179 (1956).

23,

613

FRUCTOSE- 1,6-DIPHOSPHATASES Fructose fructokinase

1/

Fructose1 -P

Fructose-6-P

It

FDPase

Fructose-l ,6-P2 Aldolase

f

Glyceraldehyde

I

+

It t-”

Glucose-6-P phosphohexose isomerase

- - - - - - -+

3r

~

1

FGlyceraldehyde 3-P

Dihydroxyacetone P + Phosphotriose isomerase

t I

I

AT<

D-Glyceraldehyde kinase

FIQ.1. Pathway of fructose metabolism in liver ( 6 ) .

Pogell :n 1961 (8),it was generally recognized that this enzyme played a key role in gluconeogenesis (9) (Fig. 2) and that it must therefore be subject to metabolic regulation in a manner complementary to the regulation of the enzyme phosphofructokinase, which catalyzes the opposing step in glycolysis. One of the most intriguing observations to emerge from the symposium held in Charlottesville in 1961 related to the p H optimum of liver FDPase. Reports from several laboratories suggested that under certain conditions, and particularly in crude extracts, FDP was hydrolyzed a t neutral pH, in contrast to the complete lack of activity below p H 8 which had been reported by Gomori ( 6 ) .

B. REGULATION AND PHYSIOLOGICAL FUNCTION

It is now generally recognized that an important site of regulation of both glycolysis and gluconeogenesis is a t the level of fructose diphosphate formation and hydrolysis (10).I n the direction of glycolysis, the activity of phosphofructokinase is inhibited by ATP and citrate, and this inhibition is reversed by AMP (11). The discovery that FDPase 8. R. W. McGilvery and B. M. Pogell, eds., “Fructose 1,6-Diphosphatase and its Role in Gluconeogenesis.” Am. Inst. Biol. Sci., Washington, D. C., 1964. 9. J. Ashmore, in “Fructose 1,6-Diphosphatase and its Role in Gluconeogenesis” (R. W. McGilvery and B. M . Pogell, eds.), p. 43. Am. Inst. Biol. Sci., Washington, D. C., 1964. 10. D. E. Atkinson and G. M. Walton, JBC 242, 2239 (1967). 11. A. J. Ramiah, J. A. Hathaway, and D. E. Atkinson, JBC 239, 3619 (1964).

614 Glucose

ATP ducokinase

S. PONTREMOLI AND B. L. HORECKER

Clucose-6-P vGlucose WPase

Fructose-6-P phosphofructokinase Fructose-l ,6-P2

It

Triose phosphate

-c Oxaloacetate

it

Phosphoenolpyruvate

y

I

FIG.2. Role of FDPase in gluconeogenesis (9).

from a number of biological sources is highly sensitive to inhibition by AMP (12-14) indicated that the direction of flow in the EmbdenMeyerhof pathway might be controlled by the ratio of AMP to adenosine triphosphate (ATP) ; high ratios would prevent glycolysis, and a t the same time permit gluconeogenesis to proceed. On the other hand, the inhibition of FDPase by AMP would protect the cell against the wasteful dephosphorylation of fructose diphosphate during glycolysis. A second possible control mechanism was suggested by the observation that modification of specific sulfhydryl groups in the protein resulted in an increased activity of the purified enzyme at neutral pH (16, 16) The final proof of the gluconeogenic function of FDPase came from observations with mutants lacking this enzyme. Bacterial strains deficient in a specific FDPase (17) were found to be incapable of growth on compounds such as glycerol, acetate, or succinate. Similar metabolic defects have recently been described in man (18, 19). Children with K. Taketa and B. M. Pogell, BBRC 12, 229 (1963). C. Gancedo, M. C . Salas, A. Giner, and A. Sols, BBRC 20, 15 (1965). K. Taketa and B. M. Pogell, JBC 240, 651 (1965). S. Pontremoli, B. Luppis, W. A. Wood, S. Traniello, and B. L. Horecker, JBC 240, 3464 (1965). 16. S. Pontremoli, B. Luppis, S. Traniello, W. A. Wood, and B. L. Horecker, JBC 240, 3469 (1965). 17. D. G. Fraenkel and B. L. Horecker, J. Bacterial. 90, 837 (1968). 18. W. C. Hulsmann and J. Fernando, in press. 19. L. Baker and A. I. Winegard, in press. 12. 13. 14. 15.

615

23. FRUCTOSE-lJ6-DIPHOSPHATASES

this disease show a tendency toward hypoglycemia accompanied by a persistent lactacidemia, which is increased after oral administration of glucose or casein.

C. METHODS OF ASSAYAND MECHANISM OF ACTION The early studies of FDPase activity employed an assay based on the release of inorganic phosphate. More recently, a coupled spectrophotometric assay was introduced in which the fructose 6-phosphate formed is converted to glucose 6-phosphate, which is allowed to reduce TPN:

+

+ +

Fructose-1,6-P~ HzO -+ fructose-6-P Pi Fructose-6-P glucose-6-P TPNH Glucose-6-P TP N+ + gluconate-6-P

+

+ H+

In the presence of excess phosphohexose isomerase and glucose-6-phosphate dehydrogenase the rate of reduction of T P N is proportional to the rate of cleavage of fructose diphosphate. For cases when small, quantities of fructose diphosphate must be used, a second spectrophotometric assay, in which fructose diphosphate is regenerated, has been proposed (20).Fructose 6-phosphate is phosphorylated with ATP and phosphofructokinase, and the adenosine diphosphate (ADP) produced is measured with phosphoenolpyruvate and lactic dehydrogenase:

+ +

+

Fructose-1,6-P2 HzO --* fructose-6-P Pi Fructose-6-P ATP -+ fructose-1,6-P~ ADP ADP P-enolpyruvate --t ATP pyruvate Pyruvate DP NH H+ ---t lactate DPN+

+ +

+

+

+

+

For studies of the specificity of FDPases, and particularly with sedoheptulose 1,7-diphosphate, the assay for inorganic phosphate is most convenient. The disappearance of SDP can also be followed spectrophotometrically using aldolase, glyceraldehyde-3-phosphate dehydrogenase, and D P N ( 6 1 ) . The hydrolysis of fructose 1,g-diphosphate occurs a t the oxygenphosphorus linkage of the substrate, leading to the formation of lSOlabeled inorganic phosphate when the hydrolysis is carried out in H2I80. No evidence for an enzyme-phosphate intermediate could be obtained (22) The form of fructose 1,6-diphosphate which is hydrolyzed appears to be the furanoside (Fig. a), based on the observation of Bencovic and

-

20. J. Mendicino and F. Vasarhely, JBC 238, 3528 (1963). 21. S. Pontremoli and E. Grazi, Bull. SOC.Chim. Biol. 42, 753 (1960). 22. S.Pontremoli, S. Traniello, B. Luppis, and W. A. Wood, JBC 240, 3459 (1965).

616

vHH

S. PONTREMOLI AND B. L. HORECKER H~OH

PI

HOjPOCH?

H

H OH

-

[POsH-

~

- HOsF’OCHl

H

+ HzPW

OH

H

FIQ.3. Hydrolysis of fructose 1,6-diphosphate by FDPase.

his co-workers (23) that methyl ~-fructofuranoside-l,6-P~, which they have synthesized, is a partial competitive inhibitor of the hydrolysis of FDP, although it is not itself hydrolyzed. In contrast, hexitol diphosphate is no more inhibitory than inorganic phosphate, which shows only weak inhibition (unpublished observations). No information is yet available as to the preference of the enzyme for one or another anomeric form of the enzyme. Of particular interest with respect to the mechanism of action of the enzyme is the finding by Bencovic et al. ( 2 3 ) that methyl fructoside 1,6-diphosphate does not protect the active site of the enzyme against acetylation by acetyl imidazole (see below) but instead increases the rate of the reaction. This suggests that the substrate induces a conformational change in the enzyme and further, since the methyl derivative is not hydrolyzed, that the hydroxyl group a t C-2 may play a role in the catalytic mechanism.

II. liver FDPase

A. PURIFICATION AND PROPERTIES 1. Purification Procedures

Since the discovery and partial purification of FDPase by Gomori ( 2 ) ,a number of purification procedures have been described ( 4 , 22, 242 7 ) . Among these, the most widely employed are based on the procedure of Pontremoli et al. ( 2 2 ) , using acetone powders from freshly collected rabbit livers. The steps include precipitation of inactive protein a t pH 4.5, fractionation with ammonium sulfate, heating to 50°, and chromatography on carboxymethyl cellulose columns, from which the enzyme S. J. Bencovic, M. M. de Maine, and J. J. Kleinschuster, ABB 139, 248 (1970). B. M. Pogell and R. W. McGilvery, JBC 208, 149 (1954). L. C. Mokrasch and R. W. McGilvery, JBC 221, 909 (1956). B. M. Pogell, BBRC 7, 225 (1962). A. Bonsignore, G. Mangiarotti, M. Mangiarotti, A. De Flora, and S. Pontremoli, JBC 238, 3151 (1963). 23. 24. 25. 26. 27.

23. FRUCTOSE-1,6-DIPHOSPHATASES

617

is specifically eluted with the substrate FDP (26). Following filtration through Sephadex G-100 the enzyme preparations, which are homogeneous a t this stage, can be crystallized from ammonium sulfate solution in the presence of MnCl,. The crystalline enzyme preparations catalyze the hydrolysis of 19.5 pmoles of FDP per minute per milligram of protein a t room temperature a t pH 9.1,and in the presence of 1 m M M n C L A modification of the column chromatographic procedures which employs back adsorption and elution has been described by Sarngadharan et al. ( 2 8 ) . However, although the specific activities of these preparations are reported to be comparable to those obtained by the procedure of Pontremoli et al., no evidence for the purity or homogeneity of the preparations was given.

2. Optimum p H and Effect of Cations Purified liver FDPase shows an absolute requirement for a divalent cation, which can be satisfied by Mn2+or somewhat higher concentrations of Mg2+.I n the presence of Mn2+the purified enzyme preparations show alkaline optima near p H 9, similar to those reported for the partially purified preparations by Gomori ( 2 ) . However, the shape of the pH activity curves was very much dependent on the nature and concentration of the activating cation. With low concentrations of Mg2+ maximum activity is also observed in the alkaline region, but at higher concentrations of this cation the p H optimum is shifted to the neutral p H range (29).The activity of the purified enzyme in the neutral pH range is also influenced by the addition of chelating agents such as histidine or glycine (30).The effects of chelating agents are similar to those reported earlier for the enzyme in crude liver extracts (31,38). Pogell et al. (33) have reported that a natural substance present in liver supernatant fractions, which appears to purify together with phosphofructokinase activity, also causes an increase in catalytic activity in the neutral pH range. The precise nature of this activator and its possible 28. M. G. Sarngadharan, A. Watanabe, and B. M. Pogell, JBC 245, 1926 (1970). 29. S. Pontremoli and E. Grazi, in “Carbohydrate Metabolism and its Disorders” (F. Dickens, P. J. Randle, and W. J. Whelan, eds.), Vol. 1, p. 259. Academic Press, New York, 1968. 30. K. Nakashima, B. L. Horecker, S. Traniello, and S. Pontremoli, ABB 139, 190 (1970). 31. R. W. McGilvery, in “Fructose 1,bDiphosphatase and its Role in Gluconeogenesis’’ (R. W. McGilvery and B. M. Pogell, eds.), p. 3. Am. Inst. Biol. Sci., Washington, D. C., 1964. 32. H. G. Hers and E. Eggermont, in “Fructose 1,6-Diphosphatase and its Role in Gluconeogenesis” (R. W. McGilvery and B. M. Pogell, eds.), p. 14. Am. Inst. Biol. Sci., Washington, D. C., 1964. 33. B. M. Pogell, A. Tanaka, and R. C.Siddons, JBC 243, 1356 (1968).

618

S. PONTREMOLI AND B.

L.

HORECKER

role in the physiological regulation of FDPase activity remain to be established.

3. Substrate Specificity The crystalline enzyme preparations were nearly equally active with F D P and the next higher homolog, SDP ( 1 5 , 2 1 , 2 7 ) ,although the enzyme has a lower affinity for the latter substrate (27). On the basis of the constant ratios of activities with these two substrates during the purification of the enzyme and from the results of competition experiments (27), it has been concluded that liver contains a single enzyme protein capable of hydrolyzing both substrates. Other phosphorylated sugars, including fructose l-phosphate, fructose 6-phosphate, and ribulose 1,5-diphosphate are not hydrolyzed, even a t much higher concentrations. The ability to hydrolyze both F D P and SDP is a common property of FDPases from mammalian tissues and is in contrast to the enzyme from Candida utilis, which is specific for FDP (see below).

B. CHANGES IN PROPERTIES INDUCED BY PROTEOLYSIS A number of investigators have reported changes in catalytic properties resulting from limited exposure to systems containing proteolytic enzymes. Gomori ( 2 ) observed a considerable increase in catalytic activity measured a t pH 9.1 when the crude liver extracts were kept a t room temperature, and similar increases were reported by Pogell ( 4 ) following incubation of partially purified preparations with particulate fractions from liver or with papain. It has also been reported that these procedures lead to decreases in enzymic activity measured at neutral pH. Similar effects have not been reported with the purified enzyme preparations] largely because these already show high activity a t alkaline pH and relatively little activity in the neutral pH range. The molecular basis for the changes, and whether they indeed result from proteolytic activity] remains to be elucidated. This will depend on the isolation of the enzyme in the form which appears to be present in crude liver extracts, and which shows the neutral, rather than the alkaline] pH optimum (%a).

C. REGULATION OF CATALYTIC ACTIVITYBY AMP 1. Efiect of pH on Inhibition by AMP

The specific inhibition of liver FDPase activity by AMP was discovered by Taketa and Pogell ( l a ) , who later provided evidence for 33a. The “native” enzyme, which shows optimum activity in the neutral pH

23. FRUCTOSE-1,6-DIPHOSPHATASES

619

the allosteric nature of this inhibition (14). An interesting feature of the inhibition by AMP, which is shared by other FDPases, is that it is much less pronounced above pH 9.0 (14, 34). I n the case of the liver enzyme, it has been established (36) that this is not a result of a lower affinity of the enzyme for the inhibitor (see below). 2. Desensitization by Papain

The allosteric model for AMP inhibition of FDPase is supported by the results of experiments in which the enzyme is desensitized to the inhibitor with little or no loss of catalytic activity. Mild digestion with papain causes almost complete loss of AMP inhibition under conditions where the catalytic activity is only slightly decreased (13). Indeed the catalytic activity measured a t alkaline pH has been observed to increase ( 3 6 ) . 3. Desensitization by Modification of Tyrosine Residues

The enzyme can also be desensitized by reagents such as acetyl imidazole, which Riordan and Vallee (37) have introduced for the specific acetylation of tyrosine hydroxyl groups in proteins. This reagent has been particularly useful in the elucidation of the role of tyrosine residues in FDPase since it has permitted the identification of four classes of such residues (38,39) based on their relative reactivity and the specific changes in catalytic properties resulting from their modification. Of the approximately 50 tyrosine residues in the protein, only 10 will react a t pH 7.5, even with a large excess of reagent. Acetylation of these tyrosine residues results in complete loss of catalytic activity. When the reagent is added stepwise, 2 of the 10 tyrosine residues are acetylated first with no detectable changes in catalytic properties. The acetylation of 4 additional tyrosine residues is accompanied by nearly complete loss of AMP inhibition and, finally, when the last 4 reactive residues are acetylated, catalytic activity is lost. The substrate and the allosteric effector provide specific protection of the tyrosines a t the catalytic and range, has recently been purified from rabbit liver [S.Traniello, S. Pontrernoli, K. Tashima, and B. L. Horecker, ABB (submitted for publication)]. 34. B. L. Horecker, S. Pontrernoli, 0. Rosen, and S. Rosen, Federation Proc. 25, 1521 (1966). 35. S. Pontremoli, E. Graai, and A. Accorsi, BBRC 33, 335 (1968). 36. G. Mangiarotti and S. Pontremoli, BBRC 12, 305 (1963). 37. J. F. Riordan, E. C. Warner, W. E. C. Wacker, and B. L. Vallee, Biochemistry 4, 1758 (1965). 38. S. Pontrernoli, E. Graai, and A. Accorsi, Biochemistry 5,3072 (1966). 39. S. Pontremoli, E. Grazi, and A. Accorsi, Biochemistry 5, 3568 (1966).

620

S. PONTREMOLI AND B. L. HORECKER

regulatory sites. In the presence of FDP, only the first 6 tyrosine residues are acetylated, and no loss of catalytic activity is observed. In the presence of AMP, again only 6 tyrosine residues are acetylated, but this time it is the catalytic activity which is lost; sensitivity to AMP is retained by the partially inactivated enzyme. The relative reactivity of these tyrosine residues toward acetyl imidazole can be correlated with the pK values established by spectrophotometric titration ( 4 0 ) . Of the 50 tyrosine residues in the native enzyme, approximately 6 are titrated with a pK value close to 8.4, 7-8 other residues show p K values of 9.0-9.2, and the remainder are not titratable below pH 10. In the presence of low concentrations of FDP, sufficient to saturate the four substrate binding sites (see below), the pK values for four of the low pK residues are shifted from 8.4 to 9.7; the other two low pK values are shifted only slightly, to 8.7. This accounts for the protection by FDP in the reaction with imidazole. Titration of tyrosine residues in the enzyme which has been acetylated in order to lose its sensitivity to AMP confirms that the acetylated residues are those with lowest pK (40). Desensitization of the enzyme to AMP inhibition can also be accomplished by treatment with diazobenzene sulfonic acid (41). In this case loss of AMP sensitivity is observed when approximately 4 monoazotyrosine residues are formed. 4. Desensitization with Pyridoxal Phosphate

In addition to the above reagents, which modify specific tyrosine residues in the protein, desensitization has been reported with pyridoxal phosphate, which forms a Schiff base derivative with lysyl residues (4.2). This reagent was first reported by Marcus and Hubert (43) to react with FDPase from swine kidney and to abolish AMP sensitivity with very little loss of catalytic activity. With liver FDPase most of the sensitivity to AMP is lost when 7-8 residues are incorporated, with concomitant loss of about 25% of the enzymic activity (4.2). The effects become irreversible when the Schiff base derivative is reduced with NaBH, and N6-pyridoxyllysine has been isolated from the reduced complex. I n the presence of AMP the sensitive lysine residues are protected, but the amount of PLP incorporated is increased (@). 40. S. Pontremoli, E. Grasi, and A. Accorsi, JBC 244, 6177 (1969). S. Pontremoli, E. Grazi, and A. Accorsi, JBC 242, 61 (1967). T. A. Krulwich, M. Enser, and B. L. Horecker, ABB 132, 331 (1969). F.Marcus and E. Hubert, JBC 243, 4923 (1968).

41. 42. 43.

23. FRUCTOSE-1,6-DIPHOSPHATASES

621

D. ACTIVATION BY CHEMICAL MODIFICATION OF SH GROUPS The crystalline liver FDPase preparations show little activity a t neutral pH, which suggests that the enzyme in situ may exist in a modified, activated form (see above). It has been suggested that the required conformational change can be induced by modification of a limited number of the 20 sulfhydryl groups in the protein (44). The original evidence for activation by modification of cysteine residues came from studies with 2,4-fluorodinitrobenzene ( 1 5 ) . Incubation of the crystalline enzyme preparations with 4 equivalents of FDNB led to a marked increase in activity in the neutral p H range, together with a small decrease in the activity assayed at alkaline pH (Fig. 4). The modified enzyme showed two broad and nearly equal activity maxima: one a t pH 7.7 and the other a t pH 9.0. When dinitrophenylation was carried out a t p H 7.5, this change in catalytic properties was associated with the modification of only 2 of the 20 cysteine residues in the protein (16). These 2 highly reactive cysteine residues were found to be completely protected against the action of FDNB by addition of FDP.

I

I 7

8

I 9

PH

FIG.4. Effect of dinitrophenylation of purified rabbit liver FDPase on the pH activity curves measured in the presence of 5 mM MgCL (16). 44. S. Pontremoli, B. Luppis, S. Traniello, M. Rippa, and B. L. Horecker, ABB 112, 7 (1965).

622

S. PONTREMOLI AND B. L. HORECKER

However, the concentrations required for protection (10-4 M ) were much higher than were required to saturate the catalytic sites and corresponded to F D P concentrations which inhibit FDPase activity (14). The results suggest that the enzyme contains two distinct sites for FDP, one the high affinity catalytic site, and the other an inhibitory site which is occupied only a t much higher concentrations of FDP. The activating effects of FDNB were observed only when the enzyme was tested with Mn2+as the divalent cation. When Mg2+was substituted for Mn2+,the dinitrophenylated enzyme was less active than the native enzyme throughout the pH range, although it still showed the biphasic pH activity curve (16). Other sulfhydryl reagents, such as p-mercuribenzoate and iodoacetamide, produced similar activation (44), except that with these compounds increases in activity were also observed a t pH 9.1 (Table I ) . With p mercuribenzoate maximum activation was observed when 2-4 sulfhydryl groups were titrated, and with excess reagent catalytic activity was almost completely abolished (44). Similar results were obtained with FDNB (15). The reactive sulfhydryl groups may be located in apolar regions of the enzyme molecule since they were not affected by N ethylmaleimide or iodoacetic acid. E. ACTIVATIONBY DISULFIDEEXCHANGE 1. With Cystamine

The first natural sulfhydryl activator of FDPase to be described was cystamine, which a t p H 7.5 was found to undergo a disulfide exchange reaction with 2 reactive cysteine residues in the protein ( 4 5 ) , leading TABLE I

EFFECT OF SULFHYDRYL REAGENTS ON FDPase Changes in catalytic activity Addition

At pH 7 . 5

At pH 9.1

FDNB (4 equivalents) p-Mercuribenzoate (4 equivalents) Iodoacetamide (10-3 M ) Iodoacetic acid (10-8 to 10+ M ) NEM (10-3 to 10-*MI

Increase (2.5-3-fold) Increase (2-fold) Increase (2-fold) No change No change

Decrease Increase (2-fold) Increase (2-fold) N o change No change

45. S. Pontremoli, S. Traniello, M. Enser, S. Shapiro, and B. L. Horecker, Proc. Natl. Acad. Sci. U.S. 58, 286 (1967).

23.

623

FRUCTOSE-1,6-DIPHOSPHATASES

to a fourfold increase in catalytic activity. Again activation was observed only when the enzyme was assayed in the presence of Mn2+;with Mg*+, activity was decreased by more than 50%. With this compound only 2 cysteine residues were incorporated even when a large excess of reagent was added. The conclusion that a mixed enzyme-cysteamine disulfide was formed was based on the fact that the reaction was reversed by reduced glutathione or cysteine (46).It is noteworthy that the disulfide exchange reaction with cystamine occurred at neutral pH and did not require a catalyst. Assuming that the disulfide exchange reaction may represent a physiological mechanism for the regulation of FDPase, Pontremoli et al. (45) have presented an interesting model for the feedback control of this system (Fig. 5 ) . Under conditions where the product F6P accumulates it would give rise to reduced TPN, which would cause the accumulation of reduced glutathione and the deactivation of FDPase. This model was shown to operate in a reconstructed system containing oxidized glutathione, TPN, and the highly purified enzymes (46). The activation of liver FDPase by a variety of sulfhydryl reagents has been examined by Little et al. ( 4 6 ) , and their results generally confirm those reported by Pontremoli and his co-workers. In the disulfide exchange reaction 5,5'-dithio-bis (2-nitrobenzoic) acid was most effective (@), and in general, the disulfides were more effective than reagents such as p-mercuribenzoate or iodoacetamide. The nature of the group introduced appears to affect the conformation of the modified enzyme. 2. With CoA and Acyl Carrier Protein Although cystamine has been reported to be a product of the physiological degradation of CoA (47) and a precursor of taurine (&), its presence in mammalian liver has not been adequately established, and Fructose-l ,6-P2 Mnz+l active FDPase

Fructose-6-P

it

Glucose-6-P

tI 1

GSH

+

TPN

_t

-

inactive FDPase

-

6-PCluconate

+

t

GSSG

TPNH

FIO.5. Scheme for feedback control of FDPase activity. 46. C. Little, T. Sanner, and A. Pihl, European J . Biochem. 8, 229 (1969). 47. G. D. Novelli, F. J. Schmetz, and N. 0. Kaplan, JBC 206, 533 (1954). 48. L. Eldjam, JBC 206, 483 (1954).

624

S. PONTREMOLI AND B. L. HORECKER

it was therefore of interest when Nakashima et al. (30,4.9) reported that similar activation of FDPase would be obtained with CoA or acyl carrier protein from E. coli. At pH 8.5 activation resulted from a disulfide interchange reaction between the protein and the oxidized coenzymes : ES-

+ COAS-SCOA-+ ES-SCOA + COAS-

With oxidized CoA, increases of nearly fivefold in the neutral FDPase activity were reported ; with acyl carrier protein or panthethine the extent of activation was somewhat smaller. Only compounds containing the terminal cysteamine residue were effective. It is of interest that with CoA maximal activation was obtained when 4 equivalents per mole of protein (one per subunit) were added. At pH 7.0 a similar activation was observed when the enzyme was incubated with reduced CoA or acyl carrier protein. At this lower pH oxidized CoA was ineffective, and the activation was inhibited by anaerobic conditions or by the addition of EDTA. It was concluded that activation a t neutral p H was an oxidative reaction catalyzed by traces of Cuz+present in the reaction mixtures: ESH

+ CoASH

---.

CUf+,O,

ESSCoA

With acyl carrier protein, ethylenediaminetetraacetate (EDTA) did not affect the activation reaction, although 0, was still required, suggesting that in this case a specific protein-protein interaction suffices to orient the SH groups in a position favorable for their oxidation. The formation of mixed enzyme-CoA or enzyme-acyl carrier protein disulfides was supported by the fact that the activation by these compounds was readily reversed by treatment with glutathione or cysteine (49).Preliminary experiments (60) with radioactive CoA or by titration of sulfhydryl groups in the protein suggest that maximum activation is associated with the incorporation of 4 equivalents of CoA per mole of protein (one per subunit). Maximum activation was observed when the enzyme was tested in the presence of low concentrations of chelating agents such as EDTA (0.5 mM) or histidine (1.0 mM) . It is noteworthy that in this case activation was observed with Mg2+ as the activating cation, as well as with Mn2+. It will be recalled that the enzyme treated with FDNB or cystamine showed activation only in the assay with Mn2+(16, 46).

3. With Hornocystine The most effective activator of liver FDPase is homocystine, which is known to be the major catabolic product in the metabolism by methio49. K. Nakashima, S. Pontremoli, and B. L. Horecker, Proc. Natl. Acad. Sci. 64, 947 (1969). 50. Y. Tashima, unpublished observations (1970).

u. S.

23. FRUCTOSE-1,6-DIPHOSPHATASES

625

nine. Exposure of FDPase to this compound results in an increase in activity of approximately tenfold when measured a t neutral pH (61). Maximum activation was observed when 4 equivalents of 35S-homocysteine were incorporated per mole of enzyme, with a corresponding decrease in the number of titratable SH groups (Fig. 6). Bound homocysteine was rapidly removed and activation reversed by addition of dithiothreitol. On activation with homocystine the pH optimum was found to shift from 8.6 to 7.3 with Mgz+ as the activating cation and from 9.1 to 8.0 with Mn2+ as the activating cation. The specific activity of the activated enzyme with MnZ+is the highest yet observed for mammalian FDPase under any assay conditions (Fig. 7 ) . 4. Disulfide Exchange as a Physiological Regulatory Mechanism

The activation of liver FDPase by formation of mixed disulfides invites speculation as to the possible role of this mechanism in t h o , and particularly as to the metabolic conditions which might result in such activation. Thus it is not likely that CoA serves as a physiological regulator since fatty acid catabolism during gluconeogenesis would leave 1000

+ DTT

I

1

I

I

I

5

I

I

I

I

I

1

I

1

I

I

15

10

I

I

I

I

I

20

I

I

I

I

I

-lo

25

Hour

FIG.6. Activation of purified rabbit liver FDPase by homocystine (61). 51. K. Nakashima, B. L. Horecker, and S. Pontremoli, ABB 141, 579 (1970).

626

S. PONTREMOLI AND

r

I

I

I

I

I

1

B. L.

HORECKER

I

50 A

Treated with 'Whomocys tine

i - \

I

I

40

\

I

\4p\

I

I

I

\

30

i 20

10 Untreated enzyme

I

I

I

1

I

I

I

FIG, 7. Effect of activation of rabbit liver FDPase by homocystine on the pH activity curves (61).(A) Assayed with MgCl,; (B) assayed with MnCL

very little CoA in the free sulfhydryl form. On the other hand, fatty acid synthesis would not be expected to occur simultaneously with gluconeogenesis in liver, and under these conditions the mammalian counterpart of acyl carrier protein would be present in the free form and would be available for the activation of FDPase. A more attractive possibility, however, is the activation of FDPase by homocystine. Gluconeogenic conditions are characterized by increased catabolism of proteins; thus, there is a strong correlation between the rates of gluconeogenesis and ureogenesis in perfused rat liver (62). The breakdown of proteins and amino acids might be expected to result in increased levels of homocystine and the activation of FDPase. It is of interest that homocysteine is the best of the physiological activators, and, as pointed out earlier, leads to increases in neutral activity which can be observed in the presence of Mg2+as well as Mn2+.It should be pointed out, however, that no evidence has yet been obtained for these activated forms of the enzyme in the irltact cell or in fresh liver extracts.

F. MOLECULAR STRUCTURE OF RABBITLIVER FDPase 1. Molecular Weight and Subunit Structure

The crystalline preparations from rabbit liver have been shown to be homogeneous on the basis of electrophoretic and ultracentrifugal 52. L. A. Menahan and 0. Wieland, BBRC 29,880 (1967).

23.

FRUCTOSE-

1,6-DIPHOSPHATASES

627

analysis. The molecular weight of the native enzyme has been estimated to be approximately 131,000 (66, 53). At pH 2 the enzyme dissociates to yield inactive molecules whose molecular weight is approximately 70,000. In the presence of mercaptoethanol a t neutral pH these halfmolecules will reassociate to yield the native enzyme with concomitant recovery of catalytic activity (64). Analysis of the quaternary structure in the presence of denaturing or dissociating agents indicates that the isolated crystalline enzyme is made up of four subunits which may not be identical. Electrophoresis on polyacrylamide gels in the presence of sodium dodecyl sulfate, or after treatment of the protein with maleic anhydride, revealed the presence of two distinct bands with migration coefficients corresponding to molecular weights of 31,000 and 39,000 (53) although the two bands were of unequal intensity. Sedimentation equilibrium experiments with the soluble maleylated subunits also indicated the presence of two species corresponding to molecular weights of 39,000 and 35,000, respectively. The presence of two nonidentical subunits was confirmed by analysis of the COOH-terminal amino acids ; hydrazinolysis yielded 0.64.8 equivalent of alanine and 0.6-1.2 equivalent of glycine (uncorrected for losses during hydrazinolysis) . 2. Binding Sites for FDP and AMP The binding of various ligands to rabbit liver FDPase has been studied in detail in Pontremoli's laboratory, using the gel filtration technique of Hummel and Dreyer (66). In the case of FDP it was established that the divalent cation was required for catalytic activity but not for binding. In addition, binding was found to be so tight that it was possible to carry out the filtration of the [ 14C]-FDP-enzyme complex without preloading the column with the substrate ( 5 6 ) .The complex emerged as a single excluded peak which yielded [14C]-FDP on acid treatment or 14C-F6P and inorganic phosphate on addition of Mn2+.At pH 7.5 the plot of complex formation vs. FDP concentration yielded a sigmoidal binding curve with 4 equivalents of F D P bound per mole of enzyme a t saturation. Calculation of the microscopic association constants confirmed the positive interaction of the binding sites with association constants ranging from 0.77 x lo7 M-' for the first equivalent to 5.8 x lo7 M-' for the last. At pH 9.1 these 53. C. L. Sia, S. Traniello, S. Pontremoli, and B. L. Horecker, ABB 132, 325 (1969). 54. S. Pontremoli, B. Luppis, S. Traniello, and A. Bargellesi, ABB 114, 24 (1966). 55. J. P. Hummel and W. J. Dreyer, BBA 63, 530 (1962). 56. S. Pontremoli, E. Grazi, and A. Accorsi, Biochemistry 7, 1655 (1988).

628

S. PONTREMOLI AND B. L. HORECKER

cooperative interactions were abolished with all 4 sites showing an association constant of 0.5 X lo7 M-I. A Hill plot of the data a t pH 7.5 resulted in an interaction coefficient of n = 1.7. In the absence of the other ligands, the binding of [ 14C]-AMP followed simple Michaelis-Menten kinetics at both pH 7.5 and pH 9.2 (35, 5 7 ) . At saturation 4 equivalents of AMP were bound with an association constant of 0.5 X lo5 M-l. The lack of AMP inhibition a t pH 9.2 (see above) is therefore not related to an inability of the enzyme to bind the inhibitor a t this pH. Below pH 7.5, however, the degree of inhibition could be correlated with the extent of binding ( 3 5 ) . In the presence of FDP the affinity for AMP a t pH 7.5 was found to increase nearly tenfold, while the binding curve changed from hyperbolic to sigmoidal ( 5 7 ) . The addition of Mg2+did not affect the binding of AMP except for the decreased binding resulting from removal of free AMP from solution as the AMP-Mg2+ complex. On the other hand, Mn2+ was found also to compete with AMP for binding sites on the enzyme as well as to induce a positive interaction between the AMP binding sites. There is some lack of agreement between these results for the binding of FDP and AMP and those reported by Pogell and his co-workers. For F D P these workers (58) obtained an association constant of 1.6 x lo5 M-l, although other data presented in the same paper, such as the failure to remove F D P by dialysis, indicated a much tighter association between the enzyme and this substrate. They were also unable to detect cooperative interactions between the binding sites. In an earlier study (59) they were also unable to detect significant AMP binding in the absence of FDP. The reasons for these differences in experimental results have not been clarified.

3. Binding Sites for the Divalent Cation Direct measurement of the binding of divalent cation has been carried out with 54Mn2t (60). At p H 7.5, four binding sites were detected with an association constant of 2 X lo5 M-l. At pH 9.2, two sets of four binding sites each were found, a high affinity set with an association constant of 1.3 x los M-l, and a low affinity set with a constant of 1.3 X lo4 M-l. Kinetic experiments confirmed that only the first set of four tight binding sites was associated with catalytic activity; the additional binding sites appeared to be related to the inhibitory effects produced a t higher concentrations of Mn2+. 57. S. Pontremoli, E. Grazi, and A. Accorsi, Biochemistry 7, 3628 (1968). 58. M. G. Sarngadharan, A. Watanabe, and B. M. Pogell, Biochemistry 8, 1411 (1969). 59. A. Watanabe, M. G. Sarngadharan, and B. M. Pogell, BBRC 30, 697 (1968). 60. S. Pontremoli, E. Grazi, and A. Accorsi, BBRC 37, 597 (1969).

23. FRUCTOSE-1,6-DIPHOSPHATASES

629

The picture that emerges indicates that of a tetrameric protein with four binding sites for each ligand, but no evidence is yet available as to the distribution of these binding sites among the various subunits. 4. Evidence for Induced Conformational Changes

We have previously reviewed some of the evidence for conformational changes induced by ligands, including the protection by substrate against dinitrophenylation of sulfhydryl groups and by both substrate and AMP against acetylation of specific tyrosine residues. The pK values for some tyrosine residues in the protein may be shifted by more than one pH unit on addition of the substrate (see above). The addition of AMP will also induce small changes in the ionization constants for the tyrosine residues with the lowest pK values (40). Evidence for conformational changes associated with the binding of ligands has also been provided by Rao et al. (61) who found that AMP markedly reduced the reactivity of SH groups, suggesting that the presence of the allosteric inhibitor changes the conformational state of the enzyme. Changes have also been observed in studies of the circular dichroism of the enzyme in the presence of ligands (62). The addition of either FDP or AMP or changes in the pH between 6.0, 7.5, and 9.1 cause small conformational changes detectable by the modification in the optical activity of a limited number of tyrosyl side chains, which is consistent with the effects of these ligands on the chemical properties of these residues. On the other hand, these ligands do not produce significant modifications of the secondary and tertiary structure of the protein, which seems to suggest a strong rigidity of its overall conformation. This conclusion is supported by the almost complete absence of changes in the circular dichroism spectra of the protein in the presence of low concentrations of SDS, although, as previously indicated, the quaternary structure was completely disrupted by this reagent. 111. Kidney FDPase

A. PURIFICATION AND PROPERTIES The presence of a specific FDPase in mammalian kidney was established by the early studies of Gomori (2). This enzyme may be identical with that found in mammalian liver since it is precipitated by antibody prepared against the purified liver enzyme and purified preparations J. G. S. Rao, S. M. Rosen, and 0. M. Rosen, Biochemktry 8, 4904 (1969). 62. A. M. Tamburro, A. Scatturin, E. Grazi, and S. Pontremoli, JBC 245, 6624 61.

(1971).

630

S. PONTREMOLI AND B. L. HORECKER

of the two enzymes behave identically in Ouchterlony double diffusion and immunoelectrophoresis ( 6 3 ) .However, the purified enzymes isolated from rabbit liver and kidney show some differences in amino acid composition and in their reaction with pyridoxal phosphate (422) which may have resulted from modification of the enzyme proteins during their isolation from these tissues. Differences in catalytic properties related to different procedures for purification have been reported for FDPase from rabbit muscle (64) (see below). The question of identity or nonidentity of liver and kidney FDPases from the same species remains to be resolved. Fructosediphosphatase has been purified from the supernatant fraction of swine kidney homogenates by a procedure which included chromatography on phosphocellulose and fractionation with ammonium sulfate (20, 6 5 ) . This preparation was free of other phosphatases, showed high affinity for fructose 6-phosphate1 and was inhibited a t concentrations M . The inhibition by higher concentrations of of substrate above substrate was greater a t lower pH; this caused the apparent pH optimum to shift from pH 8.5 to 9.5 when the substrate concentration was inM to 5 X lo-' M . The molecular weight was creased from 5 x estimated to be approximately 130,000 ( 6 5 ) . An improved procedure for the isolation of FDPase from swine kidney cortex has been described by Marcus ( 6 6 ) . Extraction was carried out in the presence of EDTA, which greatly increased the stability of the enzyme. The enzyme was purified 100-fold by precipitation a t pH 5, ammonium sulfate fractionation, chromatography on phosphocellulose, and heating to 62" a t pH 8. Fructose diphosphatase has also been purified from acetone powders of rabbit kidney (63)by acid precipitation, ammonium sulfate precipitation, heating a t pH 4.5, and chromatography on phosphocellulose, and sulfoethyl Sephadex. This preparation, which was homogeneous in gel electrophoresis, was employed for the immunological studies described earlier.

B. REGULATION OF KIDNEY FDPase Kidney FDPase preparations resemble those obtained from liver in their sensitivity to inhibition by AMP (20, 42, 43, 6 3 ) . The swine kidney enzyme was almost completely desensitized by exposure a t 30" to 22% 63. M. Enser, S. Shapiro, and B. L. Horecker, ABB 129, 377 (1969). 64. J. Fernando, M. Enser, S. Pontremoli, and B. L. Horecker, ABB 126, 599 (1968). 65. J. Mendicino, C. Beaudreau, L. L. Hsu, and R. Medicus, JBC 243, 2703 (1968). 66. F. Marcus, ABB 122, 393 (1967).

23. FRUCTOSE- l16-DIPHOSPHATASES

631

ethanol (67) or by acetylation with acetic anhydride (68). The latter treatment is reminiscent of the desensitization of rabbit liver FDPase by acetylimidazole (38,39),except that in the reaction with acetic anhydride tyrosine residues did not appear to be involved since the effects of acetylation could not be reversed by neutral hydroxylamine. The swine kidney enzyme was also desensitized, without loss of catalytic activity, by treatment with pyridoxal phosphate (43); this treatment also abolished the inhibitory effects of high concentrations of FDP. Complete desensitization was obtained when approximately 4 equivalents of pyridoxal phosphate were incorporated per mole of enzyme. The reagent appeared to react with the r-amino groups of lysine residues in the protein. Pyridoxal phosphate was found to desensitize a purified FDPase preparation from rabbit kidney (@), but in this case considerable loss of catalytic activity was observed. Like the purified enzyme from rabbit liver, rabbit kidney FDPase can be activated by a disulfide exchange reaction with CoA or acyl carrier protein, suggesting that this enzyme may also be regulated under physiological conditions by disulfide exchange reactions (30). An additional type of regulatory mechanism has been suggested by Mendicino and his co-workers (67).They observed inactivation of the enzyme in crude kidney extracts when these were incubated with ATP and cyclic 3’,5’-AMP. They suggested that this inactivation was associated with the phosphorylation of the protein and reported that it was reversed by incubation of the crude extracts in the absence of ATP. Loss of FDPase activity was also observed following incubation of kidney cortex slices with epinephrine. An extension of these studies (69) led to the conclusion that the inactivating system was present in the mitochondria1 fraction, and that A D P was more active than ATP in causing loss of FDPase activity. The rate of inactivation was reduced by the addition of tricarboxylic acid cycle intermediates or fatty acids and enhanced by 2,4-dinitrophenol1 suggesting that i t was dependent on a high ADP:ATP ratio. I n this later paper no further evidence for a phosphorylated form of the enzyme, or for a role of cyclic AMP, is presented. The interpretation of these conflicting results must await confirmation and further experimentation. They may be related t o the changes in catalytic activity by proteolytic enzymes which have been reported by Byrne (70)and Pogell and McGilvery ( 4 ) . 67. J. Mendicino, C. Beaudreau, and R. N. Bhattacharyya, ABB 116, 436 (1966). 68. F. Marcus, BBA 151, 312 (1968). 69. N. Kratowich and J. Mendicino, JBC 245, 2483 (1970). 70. W. L.Byrne, in “Fructose 1,6-Diphosphatase and its Role in Gluconeogenesis” (R. W. McGilvery and B. M. Pogell, eds.), p. 89. Am. Inst. Biol. Sci., Washington, D. C., 1964.

S. PONTREMOLI AND B.

L.

HORECKEH

IV. Muscle FDPase

A. EVIDENCE FOR THE PRESENCE OF THE ENZYME IN MUSCLE Although gluconeogenesis is generally considered to be confined to liver and kidney, evidence for the presence of a specific FDPase in muscle has been reported from a number of laboratories. Significant levels of activity are to be found in skeletal muscle of a wide variety of vertebrates including mammals, birds, and amphibia (71,72). The levels of activity in white muscle were reported to be similar to those found in liver and kidney, but the enzyme was not detected in heart muscle or in smooth muscle of several species tested. Fructose diphosphatase in crude muscle extracts has been reported to be stimulated by EDTA (72).

B. PURIFICATION AND PROPERTIES OF FDPase

FROM

RABBITMUSCLE

Fructose diphosphatase has been purified from the skeletal muscle of rabbits by two procedures (64). One procedure, involving precipitation with acid and heating a t pH 3.7, chromatography on phosphocellulose and precipitation with ammonium sulfate, yielded a preparation which showed maximum activity a t pH 7.0 a t high concentrations of Mg2+; the pH optimum shifted to pH 8.0 a t lower concentrations of the divalent cation. With Mn2+this preparation showed two pH optima, one a t 7.0 and the other at 9.2. The second procedure, which employed ammonium sulfate fractionation, heating at pH 5.0, and chromatography on hydroxylapatite and sulfoethyl Sephadex, yielded a preparation which showed only a single p H optimum a t pH 9.2 and was completely inactive at pH 7.0 in the presence of Mn2+. This preparation also showed little activity with Mg2+. The results emphasize the fact that the properties of purified FDPases may be variable, depending on the purification procedure. Purified rabbit muscle FDPase was found to catalyze the hydrolysis of SDP at the same rate as FDP, and the ratio of activities toward the two substrates remained constant throughout the purification procedure (64). Thus a single enzyme protein appears to hydrolyze both substrates. In this respect the muscle enzyme resembles other mammalian FDPases (see above). However, the enzyme from muscle was found to be much 71. M. Salas, E. Vifiuela, J. Salas, and A. Sols, BBRC 17, 150 (1964). 72. H. A. Krebs and M.Woodford, BJ 94, 436 (1966).

23.

FRUCTOSE- 1,6-DIPHOSPHATASES

633

more sensitive to inhibition by AMP, in confirmation of earlier reports for the enzyme in crude muscle extracts (71-73). The hydrolysis of FDP was more sensitive to inhibition by AMP than was the hydrolysis of SDP ( 6 4 ) .At pH 7.5, 50% inhibition of FDP hydrolysis was observed at an AMP concentration of 1.3 x M , 10 times less than was required to achieve the same inhibition of the hydrolysis of SDP. At alkaline pH higher concentrations of AMP were required to inhibit hydrolysis of both substrates. In relation to the possible role of AMP in the regulation of FDPase in muscle, Opie and Newsholme (73) have estimated that the concentration of free AMP in muscle is sufficient to effectively inhibit the enzymic activity. They found, however, that this inhibition was specifically relieved by Mnz+and suggested that this ion may participate in the regulation of FDPase activity in the intact cell. At pH 7.5 maximum rates of hydrolysis of F D P were observed a t a M, and higher concentrations were substrate concentration of 2.5 X inhibitory. At pH 9.2 the affinity for FDP was decreased, but no inhibition was observed a t higher concentrations. The K , values for SDP were found to be 0.1 mM a t pH 7.5 and 1 mM at pH 9.3 ( 6 4 ) .

C. STRUCTURE AND RELATION TO LIVERAND KIDNEY FDPase The molecular weight of muscle FDPase, based on sucrose density gradient experiments, was estimated to be 133,000, similar to that of liver FDPase (64).However, the muscle enzyme did not react with antibody prepared against purified rabbit liver FDPase ( 6 3 ) , and its amino acid composition differed significantly ( 7 4 ) . In particular, the muscle enzyme contained fewer histidine and methionine residues but was richer in tyrosine and arginine. Tryptophan is absent in all of the mammalian FDPases thus far examined, with the possible exception of FDPase from swine kidney (66). The muscle and liver enzymes could also be distinguished on the basis of their electrophoretic mobility ( 7 4 ) . Rabbit muscle FDPase resembled the enzyme from liver in its activation by fluorodinitrobenzene or p-mercuribenzoate ( 7 4 ) . Titration of 8 SH groups with the latter reagent resulted in a threefold increase in activity at pH 7.5 when the enzyme was tested in the presence of Mn2+ and slightly less activation when Mg2+was the divalent cation. A total of 14 SH groups could be titrated in the absence of denaturing agents, and all 20 of the SH groups were titrated in the presence of 5% n-propyl alcohol (v/v). Activation was also observed in disulfide exchange re73. L. H. Opie and E. A. Newsholme, BJ 104, 353 (1967). 74. J. Fernando, S. Pontremoli, and B. L. Horecker, ABB 129, 370 (1989).

634

S. PONTREMOLI AND B. L. HORECKER

actions with 5,5’-dithio-bis (2-nitrobenzoic acid) or ethyl disulfide, but cystamine was inactive, unlike the liver enzyme which was activated by this disulfide (see above). No experiments have been reported to test the activation of muscle FDPase by CoA or homocystine. Binding experiments with both FDP and AMP (75) indicate that a t saturation 4 equivalents of each ligand are bound, consistent with the four subunit structure proposed for liver FDPase. Like the liver enzyme, muscle FDPase a t pH 7.5 showed positive cooperativity in the binding of FDP, with a Hill coefficient of 1.8. The values of the microscopic association constants ranged from 0.3 to 3.6 x los M-I. At pH 9.3 the association constant was 0.6 x lo6 M - I , and cooperativity was abolished. With AMP the situation was reversed. Hyperbolic binding curves were obtained a t pH 7.5, with an association constant of 1.2 x loEM-l, and evidence for a sigmoidal binding curve was obtained a t pH 9.3, which was enhanced by the presence of AMP. The presence of F D P did not appear to alter the affinity of the enzyme for AMP. The binding studies confirmed the high affinity of the muscle enzyme for AMP as compared with liver FDPase. ROLEOF MUSCLEFDPase D. PHYSIOLOGICAL Although muscle has not been generally regarded as a gluconeogenic tissue, isolated rat diaphragm segments have been shown to actively convert radioactive pyruvate (2-14C) to glycogen ( 7 6 ) .The mechanism of glycogen formation in this tissue appears to differ from that in liver since it occurs without randomization of the radioactivity into carbon atoms 1 and 6 of hexose and without fixation of CO,. Similar observations have been reported with 2- [ “ C ] -lactate (77). Krebs and Woodford (72) have reported the activity of FDPase to be much lower than that of phosphofructokinase in muscle, but it has been estimated by Salas et al. (71) that in view of the relative mass of muscle tissue, the total FDPase activity of muscle was equivalent to that of liver. It was proposed by Krebs and Woodford (72) that the enzyme might play a role in the reconversion of a-glycerophosphate to carbohydrate since this substance was found to accumulate in significant quantities in muscle. This hypothesis has been examined in detail by 75. J. Fernando, B. L. Horecker, and S. Pontremoli, ABB 136, 515 (1970). 76. H. H. Hiatt, M. Goldstein, J. Lareau, and B. L. Horecker, JBC 231, 303 (1958). 77. L. G. Warnock, N. F. Inciardi, and W. E. Wilson, Federation Proc. 22, 298 (1963).

23. FRUCTOSE-1,6-DIPHOSPHATASES

635

Opie and Newsholme (78), who indeed found a correlation between the levels of FDPase and those of the enzymes of the a-glycerophosphatedihydroxyacetone phosphate cycle in muscles from a variety of vertebrate species having various physiological functions. The levels of FDPase were found to be much higher in white muscle than in red muscle, and the enzyme was absent in heart muscle. More recently, however, this hypothesis has been questioned by Newsholme and Crabtree (79), who proposed instead that the FDPase in muscle operates together with PFK to catalyze a cycle which enhances the sensitivity of the PFK reaction to regulation by inducing a threshhold response to changes in AMP concentration. The precise role of FDPase in muscle metabolism, and particularly the basis for the differences in FDPase content in white and red muscle, remains to be elucidated.

V. Fnrctosediphosphatase of Candida u t i h

A. PURIFICATION AND PROPERTIES The enzyme was purified from Candida utilis in 1965 by Rosen et al. (80). Dried yeast was allowed to autolyze in phosphate buffer a t pH 7.5

for 48 hr, and the enzyme was isolated in crystalline form from these autolysates by a procedure which included heating to 55" a t pH 5.0, fractionation with ammonium sulfate, and purification on phosphocellulose columns from which the enzyme was specifically eluted with malonate buffer containing 2.0 mM FDP. Crystallization was carried out by addition of ammonium sulfate in the presence of mM magnesium chloride. The Candida enzyme was more active than the mammalian FDPases; a t room temperature and pH 9.5 the crystalline protein catalyzed the hydrolysis of 83 pmoles of FDP per minute per milligram of protein. The enzyme was completely inactive with other phosphate esters, including sedoheptulose diphosphate, ribulose diphosphate, and fructose 1- or fructose 6-phosphates. Nor was the activity of the enzyme inhibited by any of these compounds. Optimum activity was observed at concentrations of FDP between 0.05 and 0.5 mM; higher concentrations of FDP (5 mM) were inhibitory. The crystalline enzyme preparation from Candida utilis showed optimum activity a t pH 9.5 with little activity below pH 8.0. I n the presence 78. L. H. Opie and E. A. Newsholme, BJ 103, 391 (1967). 79. E. A. Newsholme and B. Crabtree, FEBS Letters 7, 195 (1970). 80. 0. M. Rosen, S. M. Rosen, and B. L. Horecker, ABB 112, 411 (1965).

636

S. PONTREMOLI AND B. L. HORECKER

of 0.5 mM EDTA, however, a second pH optimum was induced a t p H 8.0 and the activity a t this pH became nearly equal to that observed at pH 9.5 (Fig. 8 ) . Similar effects were produced by histidine, KCN, and glutathione, but much higher concentrations (10 mM) of these reagents were required. Other chelating agents, including anions and intermediates of the citric acid cycle, were completely ineffective. The effect of EDTA could not be attributed to the removal of an inhibitory heavy metal ion since treatment of the enzyme with chelating resins did not have a similar effect, and the effects of EDTA were also observed in the crude extracts. It was suggested that the enzyme required a specific form of Mg2+or Mn2+chelate for activity (80).

B. INHIBITION BY AMP In common with other FDPases, the Candida enzyme was specifically inhibited by low concentrations of AMP (Fig. 9), particularly when the enzyme was assayed a t neutral p H in the presence of EDTA (80). The decreased sensitivity to AMP at the higher p H is also characteristic of other FDPases and is suggestive of a change in conformation of the

I

/

I

I

/

/

Without EDTA

i

1

PH

FIG.8. Effect of pH on FDPase from C. utilis in the presence and absence of EDTA (80).

23.

FRUCTOSE- 1,6-DIPHOSPHATASES

I

I

637 I

I

I

AMPconcentration (Mx1041

FIG.9. Effect of pH on AMP inhibition of FDPase from C. utilis.

enzyme a t this pH (see below). The enzyme was desensitized to inhibition by AMP, without loss of catalytic activity, on exposure to 1 M urea or reaction with 0.5 mM p-mercuribenzoate, but not with other sulfhydryl reagents (81). The effects of urea or p-mercuribenzoate were completely reversible. The enzyme could be irreversibly desensitized, again without loss of catalytic activity, by treatment in the presence of F D P with l-fluoro-2,4-dinitrobenzene (81) or iodine (82). Treatment with these reagents in the absence of substrate resulted in loss of catalytic activity. In each case loss of AMP sensitivity was associated with modification of 2 tyrosine residues per mole of enzyme and loss of catalytic activity with modification of 2 additional tyrosine residues (81, 8 2 ) . Desensitization by specific modification of tyrosine groups has also been reported for the enzyme from rabbit liver (see above).

C. STRUCTURE OF THE PURIFIED Candida FDPase The molecular weight of the purified enzyme was estimated from sucrose density gradient sedimentation to be approximately 100,OOO (80). At pH 4.0 it was dissociated into half-molecules, with a molecular weight of 40,000-50,000, which retained one-tenth of the activity of the native enzyme, but were much less sensitive to inhibition by AMP (81). This dissociation was prevented by the addition of the substrate, FDP. 81. 0. M. Rosen and S. M. Rosen, Pmc. Natl. Acad. Sci. U. S. 55, 1156 (1966). 82. S. M. Rosen and 0. M. Rosen, Biochemistry 6,2094 (1967).

638

S. PONTREMOLI AND B. L. HORECKER

In the presence of sodium dodecyl sulfate (SDS) , the apparent molecular weight was approximately 25,000, consistent with the presence of four subunits. The dissociation of the enzyme into half-molecules was also observed a t alkaline pH (83).At pH 9.0 this dissociation was promoted by the addition of AMP in the cold and prevented by the addition of FDP. It was proposed that dissociation a t alkaline pH was an extreme manifestation of a conformational change induced by AMP which was related to the allosteric effects observed with this effector. I n this connection it is noteworthy that the enzyme desensitized to AMP inhibition by treatment with I, or FDNB was also no longer susceptible to dissociation by AMP.

D. RELATION TO Candida SDPase In contrast to the FDPases isolated from mammalian tissues, which are active with both F D P and SDP, the enzyme in Candida utilis is completely specific for FDP. A second activity, which catalyzes the hydrolysis of SDP to S7P, has been purified from this organism. The specific SDPase differs from the FDPase in lacking the requirement for the divalent metal cation and in showing optimum activity at neutral pH. Recently, the presence of distinct FDP and SDPases in this organism has been confirmed by the separation of these enzymes in phosphocellulose chromatography and by the isolation of each enzyme in pure form ( 84) . The purified FDPase and SDPase were found to differ in molecular weight and amino acid composition. A specific SDPase, which does not act upon FDP, has been purified from baker’s yeast by Racker and Schroeder (85). This enzyme also shows no metal requirement, although it is inhibited by NaF, and, unlike the specific FDPase, it is active a t neutral pH in the absence of EDTA.

VI. FDPases in Other Microorganisms

A. THESPECIFIC FDPase

OF

Escherichia coli

The presence of specific FDPase in E . coli and the role of this enzyme in carbohydrate metabolism were established by the experiments of 83. 0. M. Rosen, P. L. Copeland, and S. M. Rosen, JBC 242, 2760 (1967). S. Pontremoli, unpublished observations

84. M. Calcagno, S. Traniello, and (1970).

85. E. Racker and E. A. R. Schroeder, ABB 74, 326 (1958).

23. FRUCTOSE- 1,6-DIPHOSPHATASES

639

Fraenkel et al. ( l 7 ) , who isolated mutant strains which had lost the ability to grow on glycerol, succinate, or acetate but grew normally on hexoses or pentoses. These organisms were shown to be deficient in a specific FDPase, which could be distinguished from the nonspecific acid hexosephosphatase present in both mutant or wild-type strains by the fact that the latter was present in the periplasmic space (86) and did not require a divalent metal cation. The properties of the specific FDPase were confirmed with a partially purified preparation (87); the E . coli enzyme was shown to be highly specific for FDP and to be active with very low concentrations of this substance. The requirement for a divalent cation was satisfied by Mg*+,which was far more effective than MnZ+; other divalent cations were either inactive or inhibitory. The partially purified enzyme showed optimum activity a t pH 7.8, with very little activity below pH 7 or above pH 9. The enzyme resembled mammalian and Candida FDPases in its sensitivity to low concentrations of AMP; it was approximately 50% inhibited a t an AMP concentration M. of 2.5 x Similar FDPase-negative mutants have been isolated by Yu et al. (88), who located the gene a t minute 84 on the E . coli map of Taylor and Thoman. The mutants isolated by Fraenkel et al. (17) mapped a t the same locus. B. OTHERBACTERIAL FDPases The first description of a bacterial FDPase was that of Fossitt and Bernstein ( 8 9 ) ,who purified the enzyme from extracts of Pseudomonas saccharophila and established the specificity of the enzyme and the stoichiometry of the reaction. Fructosediphosphatase has also been reported in Aerobacter aerogenes (go), where the enzyme is required for growth on D-fructose. Like the enzyme in E . coli, the Aerobacter FDPase exhibits optimum activity between pH 7 and 8. In this organism the obligatory pathway for fructose utilization is fructose + fructose l-phosphate + fructose l16-diphosphate. The presence of FDPase is required as a source of fructose 6-phosphate for biosynthetic pathways. Two microbial FDPases have been described which are unusual in their lack of inhibition by AMP. A species of gram-negative Acinetobacter is inhibited by ATP and citrate, but not by AMP, suggesting 86. H. C. Neu and L. A. Heppel, BBRC 17, 215 (1964). 87. D. G. Fraenkel, S. Pontremoli, and B. L. Horecker, ABB 114, 4 (1968). 88. M. T. Yu, A. R. Kaney, and K. C. Atwood, J. Bacteriol. 90, 1150 (1965). 89. D. D. Fossitt and I. A. Bernstein, J. B a c t e h l . 86, 598 (1963). 90. V. Sapico, T. E. Hanson, R. W. Walter, and R. L. Anderson, J . Bacterial. 96, 51 (1968).

640

S. PONTREMOLI AND B. L. HORECKEX

that the role of this enzyme in this organism may be catabolic rather than anabolic (91). The second is the slime mold FDPase described below.

C. FDPase

IN

SLIMEMOLDS

A highly purified FDPase from the slime mold Polysphondylium pallidum has been shown (92),to hydrolyze both F D P and SDP, a t nearly equal rates, to yield fructose 6-phosphate and sedoheptulose 7-phosphate, respectively. I n other respects the purified enzyme was remarkably similar to that isolated from Candida utilis; it was completely inactive at pH 7.5 or 8.0, and showed a pH optimum a t 9.2. I n the presence of low concentrations of EDTA a second pH optimum appeared a t pH 7.5. Unlike the Candida FDPase, however, the Polysphondylium enzyme was not inhibited by AMP a t any pH. The levels of enzyme which could be extracted from the cells did not change significantly during the various stages of differentiation, and its activity could not be related to catabolic or anabolic processes which characterize these stages. A very similar study, carried out with another species of slime mold by Baumann and Wright (93),completely confirmed the results reported earlier by Rosen (92). D. REGULATION OF FDPase AND C . utilis

IN

Saccharomyces cerevisceae

Fructosediphosphatase in S. cerevisceae appears to be an inducible enzyme, present in cells grown on lactate, ethanol, or glycerol but not in glucose-grown cells (IS). The enzyme in yeast extracts is strongly inhibited by AMP, with a value for K , of 0.08 mM. Similar increases have been reported for levels of FDPase in C . utilis when the cells were grown on glycerol, as compared with cells grown on glucose (94).

VII. FDPases in Higher Plants and Blue-Green Algae

A. PURIFICATION AND PROPERTIES Although the hydrolysis of FDP to F6P was proposed as an essential step in the reductive carbon cycle of photosynthesis (95-97), it was 91. A. J. Mukadda and E. J. Bell, BBRC 37, 340 (1969).

23. FRUCTOSE-1,6-DIPHOSPHATASES

641

not until 1958 that the presence of FDPase and SDPase activities in plant tissues was reported (85). Two enzymes capable of hydrolyzing FDP were detected in spinach leaf extracts, one with optimum activity a t pH 6.9, and the other an alkaline FDPase most active a t pH 8.5. The alkaline FDPase was purified approximately 2500-fold by a procedure which included heating to 62" a t pH 5.8, adsorption and elution from calcium phosphate gel, and fractionation with ammonium sulfate. The purified enzyme showed little activity below p H 8, and was active only with F D P ; SDP, ribulose diphosphate and monophosphate esters were not hydrolyzed. It showed an absolute requirement for a divalent cation, which was satisfied by either Mg2+or MnZ+. The neutral FDPase and SDPase activities, which were present in the crude spinach extracts, were precipitated a t lower ammonium sulfate concentration and could thus be separated from the specific alkaline FDPase. These activities appeared to be associated with the chloroplast fraction and did not require the presence of a divalent cation for activity. I n crude extracts only the alkaline FDPase activity was inhibited by antiserum prepared by immunizing rabbits with the purified alkaline FDPase. The neutral FDPase was also active with ribulose diphosphate (RuDP) (98). A similar alkaline FDPase has also been obtained in highly purified form from Euglena gran'lis (99) by heating and fractionation on DEAEcellulose. The specific activity of the best preparation was approximately one-half that reported by Racker and Schroeder for the spinach enzyme. The enzyme appeared to be specific for fructose diphosphate, although SDP and ribulose diphosphate were not tested. The enzyme also required Mg2+ and was most active a t pH 8.3; it showed very little activity a t pH 7.5 or below. More recently the properties of plant FDPases have been reinvestigated by Scala e t a2. (100).Three activities were separated from ungerminated and germinating castor beans and from mature leaves, which were similar in molecular weight (120,0oO-135,000)but which 0. M. Rosen, ABB 114, 31 (1966). P. Baumann and B. E. Wright, Biochemistry 8, 1655 (1969). S. M. Rosen, 0. M. Rosen, and B. L. Horecker, BBRC 20, 279 (1965). B. L. Horecker and A. Mehler, Ann. Rev. Biochem. 24, 207 (1955). J. A. Bassham, A. A. Benson, L. D. Kay, A. Z. Harris, A. T. Wilson, and M. Calvin, JACS 76, 1760 (1954). 97. W. Vishniac, B. L. Horecker, and S. Ochoa, Advan. Enzymol. 19, 1 (1957). 98. M. Chakravorty, H. C . Chakrabortty, and D. P. Burma, ABB 82, 21 (1959) 99. A. A. App and A. Jagendorf, BBA 85, 427 (1964). 100. J. Scala, C. Patrick, and G. Macbeth, ABB 127, 576 (1968). 92. 93. 94. 95. 96.

642

S. PONTREMOLI AND B. L. HORECKER

differed in specificity, pH optima, and sensitivity to inhibition by AMP. The properties of the three enzymes are summarized in Table 11. Only one of the three FDPases was found to be sensitive to inhibition by AMP; this was the neutral FDPase present in ungerminated castor beans. During germination a second activity appeared which was also active with RuDP. The alkaline activity was present in mature photosynthesizing leaves, and its properties resembled those of the enzyme isolated from spinach leaves by Racker and Schroeder (85). All of the enzymes from the castor bean plant were found to require either Mgz+ or Mn2+.

B. PHYSIOLOGICAL ROLEOF PLANT FDPases Racker and Schroeder (85) questioned the importance of the alkaline FDPase in photosynthesis because of its lack of activity a t neutral pH, its apparent cytoplasmic localization, and the presence of a second enzyme or enzymes which appeared to be associated with the chloroplasts and which hydrolyzed both F D P and SDP. Later work, however, has clearly established the function of this enzyme in the photosynthetic carbon cycle. Smillie has shown that the alkaline FDPase is associated with photosynthetic tissues in higher plants and Euglena (101, 102). The enzyme was also shown to be localized in the chloroplasts and to be absent in nonphotosynthetic tissue or bleached algae. It was the only FDPase detected in the photosynthetic bacterium Chromatium grown under autotrophic conditions (102). Preiss et al. (103) have pointed TABLE I1 PROPERTIES OF CASTOR BEANFDPases” PH optimum

Inhibition by AMP

FDP FDP and RuDP

7.5 6.7

Yes No

FDP

8.8

No

FDPase

Source

Specificity

I I1

Castor beans Germinating castor beans Mature leaves

I11 0

From Scala et al. (100).

101. R. M. Smillie, Nature 187, 1024 (1960). 102. R. M. Smillie, in “Fructose 1,6-Diphosphatase and its Role in Gluconeogenesis” (R. W. McGilvery and B. M. Pogell, eds.), p. 31. Am. Inst. Biol. Sci., Washington, D. C., 1964.

23. FRUCTOSE-1,6-DIPHOSPHATASES

643

out that the pH optimum of the alkaline FDPase of spinach leaves can be shifted toward the neutral range by raising the MgZt concentration; at 20 mM Mg2+optimum activity can be observed a t pH 7.4.They and other workers (104-106) confirmed the localization of FDPase in chloroplasts. The neutral and nonspecific FDPases may function in nonphotosynthetic carbohydrate metabolism in higher plants. I n the germinating castor bean acetate is utilized for the synthesis of sucrose (107, 108), and the presence of the AMP-sensitive FDPase in plant embryo tissues has been demonstrated by Bianchetti and Satirana (109). The changes in levels of this enzyme in response to changes in physiological conditions (109) support a gluconeogenic role for this enzyme. The role of the nonspecific acid FDPase in plant tissues remains unknown.

C. REGULATION OF PLANT FDPases The specific neutral FDPase present in nonphotosynthetic plant tissues resembles that isolated from animal tissues in its sensitivity to AMP. The alkaline FDPase of chloroplasts is not inhibited by AMP, but evidence has been presented which suggests that the enzyme may be inhibited by fatty acids and fatty acid esters (110). These substances also seem to inhibit the conversion of SDP to S7P in chloroplasts; the presence of this activity in chloroplasts was reported by Racker and Schroeder ( 8 5 ) , but the nature of this enzyme and its possible relation to the FDPase of chloroplasts remains obscure. An interesting observation relating to the possible control of FDP cleavage in chloroplasts has been reported by Buchanan et al. (111). A latent FDPase present in spinach chloroplast extracts was shown to be specifically activated by reduced ferrodoxin. No other cofactors were required, which appeared to distinguish the latent FDPase from the alkaline FDPase studied by other workers. 103. J. Preias, M. L. Biggs, and E. Greenberg, JBC 242, 2292 (1967). 104. M. Losada, A. V. Trebst, and D. I. Arnon, JBC 235, 832 (1960). 105. U.Heber and J. Willenbrink, BBA 82, 313 (1964). 106. J. A. Bassham, M. Kirk, and R. B. Jensen, BBA 153, 211 (1968). 107. H. L. Kornberg and H. J. Beevers, Nature 180, 35 (1957). 108. D. T. Canvin and H. J. Beevers, JBC 236, 988 (1961). 109. R. Bianchetti and M. L. Satirana, BBRC 27, 378 (1967). 110. T. A. Pedersen, M. Kirk, and J. A. Bassham, BBA 112, 189 (1966). 111. B. B. Buchanan, P. P. Kalbener, and D. I. Arnon, BBRC 29, 74 (1967).

644

S. PONTREMOLI AND B. L. HORECKER

VIII. Summary and Conclusions

A. PHYSIOLOGICAL ROLE OF FDPases The requirement of FDPase for gluconeogenesis has been firmly established by the observations with bacterial and human mutants referred to in an earlier section. However, the role of this enzyme in the regulation of carbohydrate metabolism remains to be clarified. Control of gluconeogenesis in liver appears to be exerted primarily a t the steps leading to the formation of phosphoenolpyruvate (IIZ), but evidence has accumulated which suggests that control is also exerted at the level of fructose 1,6-diphosphate. AMP, which is a specific inhibitor of FDPase in every organism examined except the slime mold, is also an activator of phosphofructokinase. High levels would thercfore stimulate glycolysis, while low concentrations would favor gluconeogenesis. However, the levels of AMP have been found not to fluctuate significantly in fed or fasted animals (IIS), and other factors must therefore contribute to the metabolic control by this substance. One such factor may be the concentration of FDP, which itself inhibits FDPase, and which greatly enhances the inhibition by AMP. An additional regulatory mechanism is suggested by the low activity of purified FDPases in the neutral pH range and by the increases in neutral FDPase activity brought about by reagents which modify sulfhydryl groups in the protein, particularly physiological agents such as CoA, acyl carrier protein, or homocystine. I n the presence of Mn2+ and chelating agents such as histidine, such modification of sulfhydryl groups shifts the pH optimum, in the case of the liver enzyme, from 9.2 to 7.5. The effects of chelating agents were observed very early and led McGilvery (SI) to suggest that these may play a role in the catalytic activity of FDPase. An interesting suggestion for the regulation of carbohydrate metabolism by FDPase is that of Newsholme and Crabtree (79), who have proposed that i t can act as a modulator of the control of phosphofructokinase by AMP in muscle. A similar suggestion for regulation of carbohydrate metabolism in liver has been advanced by Williamson and co-workers (114). 112. J. H. Exton and C. R. Park, JBC 244, 1424 (1969). 113. C. Start and E. A. Newsholme, BJ 107, 411 (1968). 114. J. R. Williamson, R. Schulz, E. T. Browning, R. G. Thurman, and M. H. Fukami, JBC 244, 5044 (1969).

23. FRUCTOSE-1,6-DIPHOSPHATASES

645

A major unsolved problem of great significance for an understanding of the regulation of FDPase activity is the neutral pH optimum for the enzyme in crude extracts, as compared to the purified enzyme preparations. There are strong indications that the protein is readily modified during purification in order to decrease the activity a t neutral pH, with a concomitant increase in the activity a t alkaline pH. It remains to be established how this observation is related to the opposite effects observed when the purified enzyme is treated with sulfhydryl reagents. Control of F D P hydrolysis may also involve increases and decreases in FDPase levels brought about by changes in diet or the effects of hormones. I n yeast FDPase levels are higher in cells grown on 2- and 3-carbon intermediates than in glucose-grown cells, suggesting that the synthesis of this enzyme may be derepressed under conditions where synthesis of hexose is required (13, 94). In mammals the administration of adrenal glucocorticoid hormones causes an increased synthesis of glucose in liver, accompanied by increases in the levels of a number of gluconeogenic enzymes, including FDPase (115, 116). These increases in activity are prevented by inhibitors of protein synthesis (116-120). Diets with low carbohydrate content (25, 1 2 l ) , fasting or alloxan diabetes (122) also cause increases in levels of FDPase in liver.

B. COMPARATIVE PROPERTIES OF FDPases At least two distinct FDPases are found in animal tissues, one in liver and kidney, and the other in white muscle. The liver and kidney enzymes show minor differences in amino acid composition and in their response to agents, such as pyridoxal phosphate (&), but these differences may be the result of modification during isolation (see above). On the other hand, the muscle enzyme is distinctly different in immunological properties as well as in amino acid composition (63, 7 4 ) . All of the mammalian FDPases are similar in having a molecular weight of approximately 135,000, and all are composed of four subunits; the 115. G. Weber, G. Banerjee, and S. B. Bronstein, JBC 236, 3106 (1961). 116. D.C. Kvam and R. E. Parks, Jr., Am. J. Physiol. 198, 21 (1960). 117. G.Weber, G.Banerjee, and S. B. Bronstein, Am. J . Physiol. 202, 137 (1962). 118. G.Weber, R. L. Singhal, N. B. Stamm, E. A. Fisher, and M. A . Mentendiek, Advan. Enzyme Regulation 2, 1 (1964). 119. E. Shrago, H.A. Lardy, R. C. Nordlie, and D. 0. Foster, JBC 238, 3188 (1963). 120. G. Weber, S. K. Srivastava, and R. L. Singhal, Life Sci. 3, 829 (1964). 121. R.A. Friedland and A. E.Harper, JBC 234, 1350 (1959). 122. G. Weber, R. L. Singhal, and S. K. Srivastava, Proc. Natl. Acad. Sci. U. S. 53, 96 (1965).

646

S. PONTREMOLI AND B. L. HORECKER

specific FDPase of plant tissues appears to have a similar structure. On the other hand, the enzyme of Candida utilis has a molecular weight of approximately 100,000, and may contain only two subunits. All of the FDPases, except that isolated from slime mold, are inhibited by AMP, and nearly all, when purified, show the characteristic alkaline pH optimum. Specific FDPases all require a divalent cation, either Mg2+or Mn2+. EDTA and other chelating agents enhance the activity of FDPase at neutral p H ; this effect is most striking with the enzyme from Candida utilis, which shows no activity in the absence of EDTA. The allosteric properties of FDPases present an interesting subject for future study. In the case of the liver enzyme the substrate shows positive cooperativity in binding, but no evidence for cooperativity in catalytic activity has been obtained. Perhaps this is because of the high affinity of the enzyme for the substrate, which prevents precise kinetic measurement a t low substrate concentration. On the other hand, the substrate has been shown to increase the affinity of the enzyme for AMP, the allosteric inhibitor. Mammalian FDPases will hydrolyze the next higher homolog, sedoheptulose diphosphate, nearly as rapidly as fructose diphosphate. In other organisms, where FDPase does not hydrolyze SDP, a second enzyme specific for SDP has been found to occur. This suggests a specific metabolic function for SDPase, which remains to be elucidated.

Bovine Pancreatic Ribonuclease FREDERIC M . RICHARDS

HAROLD W . WYCKOFF

I . Introduction . . . . . . . . . . . . . I1. Isolation and Chromatography . . . . . . . . . I11. Structure . . . . . . . . . . . . . . A . Amino Acid Sequence . . . . . . . . . B . Three-Dimensional Structure . . . . . . . . I V . Modification of Covalent Structure . . . . . . . . A . Enzymic Cleavage of the Main Chain . . . . . . B . Chemical Modification of Functional Groups . . . . C . Chemical Synthesis and S-Peptide Summary . . . . V . Molecular Properties . . . . . . . . . . . A . Physical Parameters . . . . . . . . . . B. Chain Conformation and Solvent-Induced Conformational Changes . . . . . . . . . . . . . C . Aggregation . . . . . . . . . . . . VI . Catalytic Properties . . . . . . . . . . . . A . Nature of the Reaction Catalyzed . . . . . . . B. Assays for Enzymic Activity . . . . . . . . C . Specificity in the Enzyme-Catalyzed Reaction . . . . D . Stable Complexes-Inhibition-Activation . . . . E . Steady State Kinetic Data . . . . . . . . F. Mechanism of Catalysis . . . . . . . . . G . Discussion of the Mechanism and Specificity . . . .

647 649 653 653 654 669 669 674 697 705 705 725 744 746 746 747 750 758 772 780 784

.

1 Introduction

The ribonucleases are a class of enzymes catalyzing the hydrolytic cleavage of ribonucleic acids. Although such activity can be demonstrated in almost all tissues both plant and animal. relatively few of 647

648

F. M. RICHARDS AND H.

W.

WYCKOFF

the individual enzymes have been isolated in pure form and studied in detail. This chapter is concerned almost solely with the properties of the principal component from the bovine pancreas which shows ribonuclease activity. There is more information available about this particular enzyme than about any other member of this class. The relative ease of purification, the stability, and low molecular weight have made pancreatic ribonuclease for many years one of the proteins commonly used for general studies of protein structure, for testing physical techniques, and for developing protein chemical procedures. The same properties have encouraged many groups to study the phosphodiesterase activity in attempts to get a t the general problem of the mechanism of enzymic action. Reviews of the work on this protein have appeared regularly. Most of the work prior to 1959 is summarized by Anfinsen and White (1) in the previous edition of this series. The following is a very abbreviated list of subsequent reviews: Hirs (,%’),Stein (3), Anfinsen ( 4 ), Josefsson and Lagerstedt ( 5 ) , Scheraga and Rupley ( 8 ) , Hummel and Kalnitsky ( 7 ) , Irie ( 8 ) , Stein ( 9 ) ,and Barnard (10). This review does not discuss any of the extensive work on the effects of ionizing radiation or on the immunochemistry of ribonuclease. The comparative aspects of the various ribonucleases and their possible biological roles are well covered by Barnard (lo), and a number of the related enzymes are discussed in other chapters in this volume. Egami and Nakamura (11) have reviewed the microbial ribonucleases in a separate book. I n most species the level of pancreatic ribonuclease is quite low. Its function presumably is the digestion of exogenous RNA in the diet. In ruminants there is very much more of the enzyme, and Barnard (12) has concluded that the primary purpose of pancreatic ribonuclease is digestion of the RNA of the bacteria in the rumen rather than of the dietary RNA. The reutilization of the nitrogen and phosphorus of this 1. C. B. Anfinsen and F. H. White, Jr., “The Enzymes,” 2nd ed., Vol. 5, p. 95 (1961). 2. C.H. W. Hirs, Ann. N . Y . Acad. Sci. 88, 611 (1960). 3. W. H. Stein, Brookhaven Symp. Biol. 13, 104 (1960). 4. C. B. Anfinsen, Brookhaven Symp. Biol. 15, 184 (1962). 5. L. Josefsson and S. Lagerstedt, Methods Bbchem. Anal. 9, 39 (1962). 6. H. A. Scheraga and J. A. Rupley, Advan. Enzymol. 24, 161 (1962). 7. J. P. Hummel and G. Kalnitsky, Ann. Rev. Biochem. 33, 15 (1964). 8. M. Irie, Tampakushitsu Kakusan Koso 9, 257 and 385 (1964). 9. W. H. Stein, Federation Proc. 23, 599 (1964). 10. E. A. Barnard, Ann. R e v . Biochem. 38, 677 (1969). 11. F. Egami and K. Nakamura, “Microbial Ribonucleases.” Springer, Berlin, (1969). 12. E. A. Barnard, Nature 221, 340 (1969).

24.

BOVINE PANCREATIC RIBONUCLEASE

649

large pool is presumably essential for these mammals. The release of trace metals could also be significant. Although “ferments” affecting yeast nucleic acid were recognized a t lcast as early as 1891 (IS),the work of Jones, especially his 1920 paper (14), is usually cited as the “beginning” of pancreatic ribonuclease. The modern history of the enzyme begins with its crystallization by Kunitz in 1939 (15). This by itself was enough to ensure that it came to the attention of those early in the protein crystallographic field. Although the war brought a hiatus to scientific work on this particular protein, it also brought the then existing research group a t Armour, Inc. into contact with E. J . Cohn and the blood program a t Harvard University, and with the philosophy that it was essential to have substantial amounts of pure protein if one really wanted to study it in detail. The indirect result of this exposure was the preparation by Armour in the early 1950’s of well over 1 kg of crystalline enzyme in a very high degree of purity even by today’s standards. The company offered this material at a very nominal fee to any members of the biochemical community who had a use for it. The result was an explosion of work on the enzyme which continues to this day. Although, regretfully, no one hears of Armour any more in this context, the significance of this large single preparation of known uniformity a t a time when so many techniques were being developed and compared in different laboratories cannot be overestimated.

II. Isolation and Chromatography

In view of the high stability of the enzyme most samples have been prepared by the procedure described by Kunitz (16) and modified by McDonald (17) to remove all traces of proteolytic activity. During this procedure the minced bovine pancreas is exposed to 0.25 N sulfuric acid, ammonium sulfate precipitation, 10 min a t 95”-100” and pH 3, and, finally, reprecipitation. The product can be crystallized; it was also shown later to contain a number of components all with ribonuclease activity. A practical summary of all details is given by Kunitz and McDonald (18). 13. E. Salkowski, Z . Physiol. Chem. 13, 606 (1889). 14. W. Jones, Am. J . Physiol. 52, 203 (1920). 15. M. Kunitz, Science 90, 112 (1939). 16. M. Kunitz, J . Gen. Physwl. 24, 15 (1940). 17. M. R. McDonald, J . Gen. Physiol. 32, 39 (1948). 18. M. Kunitz and M. R. McDonald, Bwchem. Prep. 3, 9 (1953).

650

F. M. RICHARDS AND H.

W.

WYCKOFF

Martin and Porter (19) described a partition chromatographic procedure and first demonstrated the presence of a t least one minor active component in the crystalline enzyme preparation. King and Craig (20) found a solvent system permitting effective countercurrent distribution of ribonuclease, ethanol: water: ammonium sulfate in the ratios 25.9: 57.6 : 16.5. The principal component of the Kunitz preparation behaved as an almost ideal solute with a partition ratio of 0.8. Albertsson has provided a liquid polymer countercurrent system based on dextrari and methyl cellulose (61). At present there are three simple and widely used chromatographic procedures : (1) Hirs et al. (22) base their method on the carboxyl ion exchange resin IRC-50 with 0 . 2 M phosphate buffer pH 6.45 as the eluting medium. The principal active component of the enzyme preparation is well retarded and is universally referred to as ribonuclease-A. Several poorly resolved faster running peaks are usually seen, the area having the highest activity and running closest to A normally being called ribonuclease B. The ratio of A to B varies with the preparation but may be as high as 10 to 1 (see Fig. 1 ) . (2) Taborsky (23) [see also Shapira ( d d ) ] has described a system based on carboxymethyl cellulose as the exchanger operated in Tris buffer a t pH 8 with a sodium chloride gradient. The excellent and adjustable resolution of this system is frequently useful. The principal peak, labeled D by Taborsky, is indistinguishable from ribonuclease-A in the IRC-50 system (see Fig. 1 ) . ( 3 ) Crestfield et al. (25) found chromatography on sulfoethyl Sephadex valuable (Fig. l c ) . Ribonuclease-A may develop heterogeneity during lyophilization and storage [see, e.g., Craig et al. ( a s ) ] . Aggregation appears to occur. A careful study of the preparation problem has been made by Crestfield et al. (25) by using chromatography on Sephadex G-75, and sulfoethyl Sephadex (3-25 as well as IRC-50. These authors recommended that RNase-A be stored as a solution in phosphate buffer a t -20", that salts be exchanged by dialysis or preequilibrated Sephadex columns, and that concentration, if necessary, be effected by ultrafiltration. If lyophiliza19. 20. 21. 22. 23. 24. 25. 26.

A. J . P. Martin and R. R. Porter, BJ 49, 215 (1951). T. P. King and L. C. Craig, JACS 80,3366 (1958). P. A. Albertsson, Nature 182, 709 (1958). C. H. W. Hirs, S. Moore, and W. H. Stein, JBC 200, 493 (1953). G. Taborsky, JBC 234, 2652 (1959). R. Shapira, Anal. Biochenz. 4, 322 (1962). A. M. Crestfield, W. H. Stein, and S. Moore, JBC 238, 618 (1963). L. T. Craig, T. P. King, and A. M. Crestfield, Biopolymers 1, 231 (1963).

24. BOVINE PANCREATIC RIBONUCLEASE

651

tion is necessary it should be carried out from dilute salt-free solution to minimize aggregate formation. The aggregates can be converted to monomers by heating to 60" for a few minutes a t neutral pH. The properties of the ribonuclease dimer are discussed below. Ribonuclease-A, as will be documented below, appears to pass all tests as a pure homogeneous protein. The origin of the minor components is not completely clear. One might expect some to arise from the strenuous acid extraction procedure [see, e.g., Dickman et aZ. (27) 1. From a commercial preparation of the enzyme with an unusually large amount of non-RNase-A material, Eaker et al. (28-30) isolated and characterized two of the components. One was identified as des-lysyl RNase and the other as des-lysyl pyroglutamyl RNase. The authors suggested that these species may arise from esterification of Glu 2 during the commercial acid-alcohol extraction procedure with subsequent tryptic cleavage followed by ester hydrolysis or pyroglutamyl ring closure. However, several of the minor components occur naturally (31) as they can be demonstrated in direct chromatography of pancreatic juice (32) or extracts of zymogen granules (3s). The normal minor components that have been isolated and studied are glycoproteins where the protein moiety appears to be indistinguishable from RNase-A but the sugar complement differs between the different components (34, 3 5 ) . Whether there is any component corresponding to the loss of a single amide residue as originally proposed by Tanford and Hauenstein (36) to explain the RNase-A:RNase-B difference is now a moot question. In one preparation of RNase-B the sugar consisted of 2 glucosamine (probably acetylated) and 6 mannose residues and was attached to Asn 34 by a P-aspartamido [-2-acetamido] 1,2-dideoxy-/3-~-glucoselinkage (37, 38). Shapira and Parker (39) have reported a peak moving ahead of RNase-A on CM-cellulose which is produced by brief heating at 100" 27. S. R. Dickman, G . A. Morrill, and K. M. Trupin, JBC 235, 169 (1960). 28. D. L. Eaker, T. P. King, and L. C. Craig, Biochemistty 4, 174 (1965). 29. D. L. Eaker, T. P. King, and L. C. Craig, Biochemktry 4, 1479 (1965). 30. D. L. Eaker, T. P. King, and L. C. Craig, Biochemistty 4, 1486 (1965). 31. W. H. Stein, Ciba Found. Symp., Chem. Struct. Proteins, p. 17, disc. 27, (1954). 32. P. J. Keller, E. Cohen, and H. Neurath, JBC 233, 344 (1958). 33. L. J. Greene, C. H. W. Hirs, and G . E. Palade, JBC 238, 2054 (1963). 34. T. H. Plummer, Jr., C. H. W. Hin, and A. L. Tench, JBC 238, 1396 (1963). 35. T. H. Plummer, Jr. and P. Kosinski, JBC 243, 5961 (1968). 36. C. Tanford and J. D. Hauenstein, BBA 19, 535 (1956). 37. T. H. Plummer, Jr. and C. H. W. Hirs, JBC 239, 2530 (1964). 38. T. H. Plummer, Jr., A. Tarentino, and F. Maley, JBC 243, 5158 (1968). 39. R. Shapira and S. Parker, BBRC 3, 200 (1960).

652

F. M. RICHARDS AND H. W. WYCKOFF

Effluent volume (ml) (b)

2.4

Sulfoethyl sephadex

I.6

0.8 0

30

60

90

24.

653

BOVINE PANCREATIC RIBONUCLEASE

at pH 3.2. Moskvitina and Budovskii (40) found a component produced from heated samples of RNase and moving ahead of RNase-B. This component, designated RNase-F by the latter authors, is quite stable in salt solutions (for example, 0.01 M borate buffer, pH 8.0) but disappears in salt-free solutions. The material appears to be a conformational isomer. The substrate specificity is altered since RNase-F cleaves polyadenylic acid a t a rate about one-eighth that of yeast RNA (41). In view of the normal commercial preparation procedures, varying amounts of RNase-F might be expected in these samples.

111. Structure

A. AMINOACIDSEQUENCE The amino acid composition of RNase is listed in Table I. This corresponds closely to the values normally measured on a 24-hr hydroTABLE I AMINOACID COMPOSITION OF RIBONUCLEASE A Amino acid

Number of Residues

Asparagine Aspartic acid

lo 15

Threonine Serine Glutamine Glutamic acid Proline

5

Amino acid Glycine Alanine

Number of Residues 3 12

Amino acid Leucine Tyrosine

10 15

Half-cystine Valine

85 9

Phenylalanine Lysine

7 12 5

Methionine Isoleucine

4

Histidine Arginine

3

Number of Residues 2 6 3 10 4 4

4

FIG.1. (a) Chromatography of ribonuclease on Amberlite IRC-50, 0.9 X 30 cm column in sodium phosphate buffer 0.2 M pH 6.47. Reproduced from Hirs et al. (2.2). (b) Chromatography of ribonuclease on carboxymethyl cellulose 0.9 X 20 cm column, in tris buffer 0.005M pH 8, NaCl gradient as shown: ( 0 )200 mg of RNase and (0) 41 mg of RNase. Reproduced from Taborsky (23). ( c ) Chromatography of ribonuclease-A on sulfoethyl Sephadex, 0.9 X 60 cm column, 0.1 M phosphate buffer, pH 6.47, 5 mg load. Reproduced from Crestfield et al. ( 2 6 ) .

654

F.

M.

RICHARDS AND

H.

W. WYCKOFF

lysate except for those residues suffering losses on hydrolysis and those that are slowly released (Ile and Val). The original work of Sanger on the sequence of insulin saw the elegant application and development of the then recently discovered paper chromatographic techniques. The work on ribonuclease started a short time later in the laboratories of Anfinsen and his associates and of Him, Stein, and Moore and their colleagues. This was the first enzyme and only the second protein whose sequence was to be determined. The work spanned and spurred the development of the amino acid analyzer, the fraction collector, the peptide column separation procedures, the refinement of the Edman procedure, in fact, most of the procedures used today except for gas chromatography and mass spectroscopy. This work appeared in a series of papers from Anfinsen et al. (@-46) and the monumental series from the Rockefeller Institute (47-54). The final summary is given by Smyth et al. (55) and is given in Table 11. More recent data on the pancreatic enzymes from certain other species are also included in Table I1 (56, 6 7 ) .

B. THREE-DIMENSIONAL STRUCTURE The initial X-ray diffraction study of ribonuclease was reported by Fankuchen in 1941 ( 5 8 ) . Since then investigations have been carried on for many years by C. H. Carlisle and his associates [see Avey et al.

. 42.

C. B. Anfinsen, M. Flavin, and J. Farnsworth, BBA 9, 468 (1952). 43. C. B. Anfinsen, R. R. Redfield, W. L. Choate, J. Page, and W. R. Carroll, JBC 207, 201 (1954). 44. R. R. Redfield, C. B. Anfinsen, and J. Cooke, JBC 221, 385 (1956). 45. A. P. Ryle and C. B. Anfinsen, BBA 24, 633 (1957). 46. J. T. Potts, A. Berger, J. Cooke, and C. B. Anfinsen, JBC, 237, 1851 (1962). 47. C. H. W. Hirs, W. H. Stein, and S. Moore, JBC 211, 941 (1954). 48. C. H. W. Hirs, W. H. Stein, S. Moore, and B. M. Fallon, JBC 221, 151 (1956). 49. C. H. W. Hirs, S. Moore, and W. H. Stein, JBC 219, 623 (1956). 50. J. L. Bailey, S. Moore, and W. H. Stein, JBC 221, 143 (1956). 51. D. H. Spackman, W. H. Stein, S. Moore, and A. M. Zamoyska, JBC 235, 648 (1960). 52. C. H. W. Hirs, S. Moore, and W. H. Stein, JBC 235, 633 (1960). 53. C. H. W. Hirs, JBC 235, 625 (1960). 54. D. G. Smyth, W. H. Stein, and S. Moore, JBC 237, 1845 (1962). 55. D. G. Smyth, W. H. Stein, and S. Moore, JBC 238, 227 (1963). 56. R. L. Jackson and C. H. W. Him, JBC 245, 637 (1970). 57. J. J. Beintema and M. Gruber, BBA 147, 612 (1967). 58. I. Fankuchen, J. Gem. Phvsiol. 24, 315 (1941).

TABLE I1 SEQUENCES OF SOME RIBONUCLEASES".)

20 Lys-Glu-Thr-Ala-Ala-Als-Lys-Phe-Glu-Arg-Gln-His-MeeAspSer-Ser-ThrSer-Als- Ala Gln Ser Pro Lys Gln Pro AspSer Ser Ser Thr Glu Gly Pro Ser Lys Gly Glu Ser Arg Ser Ser Asp LYS 25 30 35 40 Ser - Ser- Ser-Asn-Tyr-Cys-Asn-Gln-MeeMeeLys-Ser-Arg-Asn-Leu-Thr-Lys-Asp-Arg-Cys ASN Leu Ser Arg ASNMet Gln Gly Arg Gln G 1y Met Gly Ser Pro Thr Gln 45 50 55 60 Lys-Pro-Val-Asn-Thr-Phe-Val- His-Glu-Ser- Leu-Ala-AspVal-Gln-Ala-Val-Cys- Ser- Gln 1

Bovine Porcine Rat Bovine Porcine Rat Bovine Porcine Rat Bovine Porcine Rat Bovine Porcine Rat Bovine Porcine Rat Bovine Porcine Rat

5

10

15

Pro Glu Ile 65 70 75 80 Lys-Asn-Val-Ala- Cys-Lys- Am-Gly-Gln- Thr- Asn-Cys-Tyr-Gln-Ser-Tyr- Ser-Thr-Me6 Ser Ile Asn ASN His Gly Gln Thr k g ASP HisLys Ser Leu Arg 85 90 95 100 Ile- Thr- Asp-Cys-Arg-Glu- Thr-Gly-Ser- Ser- Ly s-Tyr-PreAsn-Cys-Ala-Tyr-Lys-Thr-Thr Gln Ala Ser Leu Lys Thr Asn 105 110 115 120 Gln-Ala-Asn-Lys-His-Ile-I1e- Val- Ala-Cys- Glu-Gly-Asn-Pro -Tyr-Val-Pro- Val- His- P h e Glu Gln Pro Asn Ser Glu Ile Asp 125 AspAla-Se r-Val

For the porcine and rat enzymes only the differences from the bovine enzyme are shown. The sites of carbohydrate attachment in the porcine enzyme are shown by the residues in full capital letters (Asn, 21, 34,and 76).For all the enzymes the four disuliide groups are paired in the same way, 26-84,40-95, 58-110, and 65-72. * Data for bovine from Smyth et a2. (66),porcine from Jackson and Him (66),and rat from Beintema and Gruber (67).

F 8 3

3 w * 2La P

2 s 2r P

B

(I

8

656

F. M. RICHARDS AND H. W. WYCKOFF

(59) for a recent publication] and by Harker, Kartha, Bello, and their associates. Both groups have worked with crystals grown from aqueous organic solvent mixtures. Many different crystalline forms of ribonuclease were found and reported during this work. I n 1967, Kartha e t al. (60) reported the successful interpretation of an electron density map at 2 A resolution and gave a picture of the course of the peptide chain. A separate investigation of the modified enzyme RNase-S (see Section IV,A,l) has been carried out by Wyckoff and Richards and their colleagues on crystals grown from strong salt solutions. The structure of this enzyme a t 3.5 A (61) and a t a nominal resolution of 2 A (66) has been reported in detail by Wyckoff et al. A stereopicture of the model is presented in Fig. 2. The structures of RNase-A and RNase-S have not been fully compared a t this time. However, Dickerson and Geis (63) have drawn two stereopairs of the a-carbon chains of these two enzymes in very similar orientations as shown in Fig. 3. The structures are undoubtedly very similar in most regions. A known exception will be residues 16 through 23, containing the cleaved bond in RNase-S, where the enzymes are clearly very different. It should be noted that the quality of the electron density maps differs markedly from one region to another. This may result from buildup of experimental errors in certain places; it may reflect motion of certain parts of the molecule; or it may reflect the existence of two or more slightly different conformers, with appropriate statistical weights, coexisting in the same crystal lattice. The net result of any of these effects is lower peak electron densities and consequent uncertainty in interpretation. Whether the differences between the structures of A and S-apart from the 1&23 region-are real or just, the result of errors in interpretation is not yet known. The extent to which the dilute solution structure of an enzyme differs from that in the crystalline state is still the subject of some debate 59. H. P. Avey, M. 0. Boles, C. H. Carlisle, S. A. Evans, S. J. Morris, R. A. Palmer, B. A. Woodhouse, and S. Shall, Nature 213, 557 (1967). 60. G. Kartha, J. Bello, and D. Harker, Nature 213, 862 (1967). 61. H. W. Wyckoff, K. D. Hardman, N. M. Allewell, T. Inagami, L. N. Johnson, and F. M. Richards, JBC 242, 3984 (1967). 62. H. W. Wyckoff, D. Tsernoglou, A. W. Hanson, J. R. &ox, B. Lee, and F. M. Richards, JBC 245, 306 (1970). 63. R. E. Dickerson and I. Geis, “The Structure and Action of Proteins.’’ Harper, New York, 1969.

FIG.2. Stereoscopic view of a skeletal model of RNaseS deduced from the 3.5-A resolution map and chemical sequence data. The small balls locate sulfur atoms. The large ball hanging from the top support plate shows the van der Wads size of a paraffinic hydrogen atom.

24.

BOVINE PANCREATIC RIBONUCLEASE

657

(64,65).That they are very similar in general conformation is no longer in doubt, but subtle differences may still exist whose possible significance will depend on the questions being asked. Doscher and Richards (66) demonstrated that molecules of RNase-S are catalytically active while in the crystal lattice in equilibrium with 90% saturated ammonium sulfate. Bello and Nowoswiat (67)later showed the same phenomenon for crystals of RNase-A immersed in 75% 2-methyl-2,4-pentanediol. These two studies not only indicate a strong similarity between the solution and crystal structures but also a strong similarity between the structures of the enzyme in these two very different solvents. However, it should be noted that in RNase-S a sulfate ion is almost certainly bound specifically a t the active site (62) while with RNase-A satisfactory crystals were only obtained with a t least small amounts of inorganic phosphate present, and here a specific binding of phosphate a t the active site was found (60,68). Both of these structures could be slightly different from that of the enzyme in the absence of any polyvalent anions. Thus the changes observed crystallographically on the binding of ligands may not be identical to those found in dilute solution since the starting structure may be different from that in the solution work in the absence of the polyvalent anions. Bello and Nowoswiat (69)have studied the alkylation of His 12 and 119 in RNase-A crystals. The reaction is very similar to but not identical to that found in solution. Bello and Harker (70) crystallized fully deuterated RNase-A and showed that the diffraction pattern was identical to that of the protonated form within the limits of measurement. This observation is very important for the interpretation of hydrogen exchange data. The approximate principal dimensions of the RNase-A molecule based on the X-ray structure are 38 x 28 x 22 A (60).The maximum axial 64. F. M. Richards, Ann. Rev. Biochem. 32, 269 (1963). 65. J. A. Rupley, in “Structure and Stability of Biological Macromolecules” (S.N. Timasheff and G. Fosman, eds.), p. 291ff. Marcel Dekker, New York, 1969. 66. M. S. Doscher and F. M. Richards, JBC 238, 2399 (1963). 67. J. Bello and E. F. Nowoswiat, BBA 105, 325 (1965). 68. G. Kartha, J. Bello, and D. Harkcr, in “Structural Chemistry and Molecular Biology” (A. Rich and N. Davidson, eds.), p. 29. Freeman, San Francisco, 1968. 69. J. Bello and E. F. Nowoswiat, Biochemistry 8, 628 (1969). 70. J. Bello and D. Harker, Nature 192, 756 (1961).

FIO.3. Stereodiagrams of a-carbon atom chains of RNase-A and RNase-S from the studies by X-ray diffraction on the crystalline enzymes: (a) RNase-A from the work of Kartha et al. ( 6 0 ) . (b) RNase-S from the work of Wyckoff et al. (62). The orientation of the molecules has been made as similar as possible to simplify comparison. The figures are reproduced with the kind permission of Dickerson and Geis (63).

658

F. M. RICHARDS AND H. W. WYCKOFF

ratio is thus less than 2. From measurements on an atomic model, RNaseS has a maximum dimension of about 47 A and a minimum dimension of 25 A measured to the outside of side chain atoms. These dimensions are somewhat uncertain because of the potential flexibility of side chains. Wyckoff et al. (62) have provided a preliminary coordinate list of all nonhydrogen atoms in RNase-S. Along with the list is a series of notations on the quality of the map and the fit of the atomic model to the electron density contours. The following comments concerning group accessibilities are based on this coordinate list, but detailed interpretations must be made with caution in view of the uncertainties in many parts of the structure. A method of calculating the static accessibility of each atom from this coordinate list has been devised by Lee et al. ( 7 l ) ,and these values are given in Table I11 along with a list of qualifying comments from Wyckoff et al. (62). The calculation is based on an assumed van der Waals radius for each nonhydrogen atom or group (e.g., CH, and CH,) and an assumed solvent molecule radius. The locus of possible centers of these molecules in contact with the protein defines a hydrosphere surface, and a portion of this surface is associated with each atom of the protein. A plot of the total accessibility, in A2,of each side chain is given in Fig. 4, grouped according to type of amino acid and in Fig. 5a as a function of sequence number. Calculations were also performed for each amino acid, XI in a fully extended sequence Gly-X-Gly and these values are given for comparison in Fig. 4. It can be seen from these data that the larger hydrophobic side chains are the most buried with the exception that cystine also tends to be quite inaccessible with 26, 72, 84, and 110 completely buried. All of the alanines are exposed, three of the four prolines are very exposed, three of the valines are completely buried as are Met 30, Phe 46, and Ser 90. Phenylalanine 8 is only accessible via a tunnel from the surface which is in fact occupied and blocked by one well-defined solvent molecule. The various residues of each polar amino acid have a wide range of exposure, but the larger residues tend to be most accessible with the exception of the tyrosines, which are quite variable. Residues in the active site region, 11, 12, 41, 43, 44, 45, 119, 120, 121, and 123, tend to be the extremes within each residue type; but it should be noted that the motion of His 119 to the active position proposed later (Section VI) would increase the exposure of 11, 12, 41, and 44 and decrease the exposure of 121 and 109 in particular. The hydrophilic residues and especially the hydrophilic portions of these residues are generally ex71. B. Lee and F. M. Richards, J M B 55, 379 (1971).

24.

659

BOVINE PANCREATIC RIBONUCLEASE

Car boxyl and Hydroxyl Amide

Hydrophobic

Basic

I-I I

LY

r

50

0

80 72 84

110

46 90

57 108

FIQ.4. Area of the first shell of solvent molecules in potential contact with side chain atoms of each residue. The method of calculation is indicated in the text. Side chain atom radii were assumed to be 1.8 A and solvent molecule radii 1.4 A. The upper bar for each amino acid is the value calculated for the residue in a hypothetical sequence Gly-X-Gly in a p-like structure and presumed to represent the maximum possible solvent accessibility. Residues with no detectable exposure to solvent are listed below the axis. Some of the residues in the recognition site are indicated by underlining.

posed. The larger hydrophobic groups tend to be buried and in particular there is a hydrophobic core containing 16 methyl groups, 8 methylene groups, 2 phenylalanyl and 2 niethionyl residues, 1 disulfide bond, 1 prolyl and part of 1 histidyl residue. The total hydrosphere surface of 7000 A* is however composed of 3200 AZ of C and S compared to 3800 Az of N and 0. The surface is thus 45% hydrophobic. In fact this is a lower limit since the van der Waals radius of 1.8A was used for all side

660

F. M. RICHARDS AND H. W. WYCKOFF

;150

5 a w

100

K

a W

50

9 K 3

In 0 a n

0 0.5

=

o

*

' > 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.

BOVINE PANCREATIC RIBONUCLEASE

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.

BOVINE PANCREATIC RIBONUCLEASE

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

BOVINE PANCREATIC RIBONUCLEASE

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

F. M. RICHARDS AND H . W. WYCKOFF

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.

BOVINE PANCREATIC RIBONUCLEASE

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

F. M. RICHARDS AND H . W. WYCKOFF

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.

BOVINE PANCREATIC RIBONUCLEASE

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

BOVINE PANCREATIC RIBONUCLEASE

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.

BOVINE PANCREATIC RIBONUCLEASE

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

F. M. RICHARDS AND H. W. WYCKOFF

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 <100 100 86 71 97 NRa NR NR NR

C>P > p)

(U

NR NR 0 NR NR <0.4

0

100 0 60

145 100 112 25 140 90 140 55 NR 110 NR 130 129 60 0.4 3 0.2 100

No. of buried TYr residues

Ref.

3 3 3 NR 3 NR NR NR NR NR 3

NR 2 2 '

NR NR NR NR NR NR NR NR NR NR NR NR NR

Abbreviations: DNP, dinitrophenyl; SNP, sulfonyloxynitrophenyl; TFA, trifluoroacetyl; and others as explained in Table V footnote. In the polyamino acid derivatives the subscript is the total number of residues added to the protein molecule. These residues are distributed among the number of amino groups listed in column 3. I n column 3, analytical number of modified residues is given except where an amino acid prefix occurs; the number then refers to the position in the amino acid sequence. See Goldstein (119), less active than RNase-A, but active.

24.

BOVINE PANCREATIC RIBONUCLEASE

679

a. O-Methylisourea and Allcylimidates. These reagents are very specific for amino groups and react in alkaline solution. The products are guanidine or amidine derivatives which maintain the positive charge of the original amino groups although the pK values are higher and several additional atoms have been added. Guanidination of all 10 c-amino groups with O-methylisourea causes loss of all enzymic activity toward RNA (103).The product has a sedimentation constant and alkaline spectral titration behavior which indicate no marked conformational change. Extensive guanidination can occur with no loss of activity. Only the last one or two groups to be modified are related to the activity loss. Allewell (120)found a residual 1% activity toward C > p after exhaustive reaction of RNase-S. The product was crystallized in a new space group. Reaction with methyl acetimidate produced a fully modified and totally inactive derivative (104). The physical properties were all very similar to RNase-A. The acetimido groups have an advantage over the guanidino groups in that they can be removed in aqueous ammonia. The regenerated protein had full enzymic activity. b. Lysine-Protecting or E n d Group Reagents. I n addition to the methyl acetimidate discussed above, a number of reagents primarily of use in primary structure work have been tried on ribonuclease. Thus cyanate (121) rapidly inactivated the enzyme. When, stoichiometrically, one lysine had been converted to homocitrulline the C > p activity had already been reduced to 50%. Isolation of homogeneous derivatives was not reported. All amino groups can be trifluoroacetylated with ethyl thioltrifluoroacetate (105). The product has no enzymic activity toward RNA. The modifying groups can be removed with 1 M piperidine a t 0".Interestingly, the resulting protein had little activity, was sensitive to trypsin, and had altered spectral properties. All of these properties were normalized after reduction and reoxidation of the disulfide bonds. Cyanoethyl derivatives can be prepared with acrylonitrile. Both monoand disubstituted amino groups can be produced. Reaction a t p H 9.2,2", with 0.4M acrylonitrile for 7 days modified all lysine residues with no apparent change in any of the other amino acids (106). The lysine was almost entirely accounted for as dicarboxyethyllysine in acid hydrolysates of the derivative. The a-amino group had apparently also reacted. The fully substituted derivative had no RNA activity. The physical properties were very similar to those of native RNase-A. 120. N. Allewell, Ph.D. Dissertation, Yale University, 1969. 121. G . R. Stark, W. A. Stein, and S. Moore, JBC 235, 3177 (1960).

680

F. M. RICHARDS AND H. W. WYCKOFF

Diketene will add to protein amino groups to give acetoacetyl derivatives (12.2).Aliphatic and phenolic OH groups may also be acetoacetylated. The latter can be removed in carbonate buffer pH 9.5 without losing the nitrogen bound groups. The amino groups can be smoothly regenerated with hydroxylamine a t pH 7.

c. Nitrofluorobenzene Reagents. Dinitrofluorobenzene is routinely used for reaction with amino and other functional groups in primary structure work. A very important series of studies has been carried out by Hirs and his associates with this and similar reagents acting on ribonuclease under nonoptimal conditions. At pH 10 all amino groups can be dinitrophenylated (108), but the peculiar relative reactivities of certain specific residues are enhanced a t p H 8. Limited reaction and careful chromatographic fractionation have permitted the isolation of well-characterized derivatives. The a-amino group a t position 1 has a reactivity comparable to that of a simple dipeptide and reduces the catalytic activity of the enzyme to 60%. The c-amino group a t position 41 reacts about 70 times faster than that of a lysine residue in a simple dipeptide. Doscher reported that modification of this group causes a reduction in activity to no more than 0.01% of RNase-A ( 1 2 3 ) .The reaction also increases the reactivity of Lys 7. Reaction a t 41 appears to be obligatory for reaction at 7. No derivative of RNase-A has been obtained with reaction solely a t Lys 7. The reaction a t Lys 41 is prevented by substances which are themselves strong competitive inhibitors of the enzyme. A detailed study of the kinetics of the Lys 41 reaction by Murdock et al. (1.24) indicates that the PI(, of Lys 41 is 8.8 + 0.1.This value is about 1.4 pH units lower than that for the usual r-amino groups and would account for the apparent enhanced reactivity a t lower pH. The second-order rate constant corrected t o the free base form is within a factor of 2 of that derived from data on simple peptides. A strongly cationic center was inferred as the reason for this low pK,. I n the X-ray structure a number of cationic residues are, in fact, close by, especially Arg 39. Studies with the reagent 4-sulfonyloxy-2-nitrofluorobenzenea t 28" again showed major reaction a t Lys 41 but a t a rate about equal to that at the &-amino group (109, 110). No reaction was observed a t Lys 7. The pK, of the other 9 lysine residues was 10.1 in agreement with titration data. I n the thermal transition region above 50" the behavior of Lys 41 approached that of the other normal lysine residues. d. N-Carboxyanhydrides. Peptide chains of varying length can be 122. A. Marsotto, P. Pajetta, L. Galzigna, and E. Scoffone, BBA 154, 450 (1968). 123. M. Doscher, manuscript in preparation. 124. A. L. Murdock, K. L. Grist, and C. H. W. Hirs, ABB 114, 375 (1966).

24.

BOVINE PANCREATIC RIBONUCLEASE

681

"grown" from the amino groups by treatment with the N-carboxyanhydride of an a-amino acid. The €-amino groups are then replaced with a-amino groups in altered positions. Alanine chains of varying length have been attached to RNase (111) in the presence of phosphate buffer pH 8, 5", 25% dioxane. Samples with an average chain length of 7 residues still showed very high catalytic activity toward RNA (Anfinsen assay). About 3 residues of lysine appeared to be relatively resistant to alanination by this procedure. They have been identified as 7, 37, and 41 (125).These residues become more susceptible to reaction in bicarbonate buffer. Substitution a t Lys 41 appears to be associated with the activity loss. Although many preparations retain some catalytic activity toward both high and low molecular weight substrates, the activity parameters do not parallel those of unmodified RNase when studied in detail as a function of pH and ionic strength (112).The variations are also dependent on the nature of the substrate which implies changes in specificity. The data are not complete enough to permit more than qualitative comparisons. Peptide chains involving the following amino acids have been added to the amino groups : lysine, ornithine. arginine, histidine, tyrosine, phenylalanine, valine, and glycine. Most of the derivatives showed activity, but, as described above, they differed from RNase-A in the dependence of activity on ionic strength and pH. For the derivatives involving side chains with a positive charge, the activity toward RNA a t low ionic strength decreased rapidly as the extent of substitution increased. The interesting interpretation (114) is that tight complexes of RNA and the peptide chains form a t a distance from the original enzyme surface preventing access to the active site. This apparent inhibition is relieved a t high ionic strength.

e. N-Acetyl Homocysteine Thiolactone. This reagent will react with amino groups and result in the introduction of SH groups into a protein. Treatment of RNase with this reagent produced a variety of products when the reaction was carried out a t p H 8, 25", with a substantial molar excess of reagent (126). A number of the products were enzymically active. Much greater selectivity was obtained a t pH 7.5 with molar ratios of 1-4 and in the presence of silver ion (127, 128). Two monosubstituted products were obtained, one active, one inactive. The inactive derivative by inference is assumed to be substituted on Lys 41. 125. J. P. Cooke, C. B. Anfinsen, and M. Sela, JBC 238, 2034 (1963). 126. F. H. White, Jr. and A. Sandoval, Biochemistry 1, 938 (1962). 127. S.Shall and E. A. Barnard, Nature 213, 562 (1967). 128. S. Shall and E. A. Barnard, JMB 41, 237 (1969).

682

F. M. RICHARDS AND H.

W.

WYCKOFF

f. Haloacetates. In alkaline solution lysine residues can be alkylated in the presence of iodo- or bromoacetate ions (118). Both mono- and dicarboxymethyl derivatives can be formed. Some of the characterized derivatives are listed a t the end of Table VI. Number 25, c-CM-Lys-41RNase, shows a very low activity when measured by a step 2 assay employing C > p as substrate. This same compound is active in the depolymerization of 5 s RNA (119) but the evidence presented to show that it is not the result of contamination with native RNase-A can be interpreted to suggest the opposite.

3. Met hionine

The sulfur atom of methionine residues may be modified by formation of sulfonium salts or by oxidation to sulfoxides or the sulfone. The cyanosulfonium salt is not particularly useful for chemical modification studies because of the tendency for cyclization and chain cleavage (129). This fact, of course, makes it very useful in sequence work. Normally, the methionine residues of RNase can only be modified after denaturation of the protein, i.e., in acid pH, urea, detergents, etc. On treatment with iodoacetate or hydrogen peroxide, derivatives with more than one sulfonium or sulfoxide group did not form active enzymes on removal of the denaturing agent (130) [see, however, Jori et al. (131)].There was an indication of some active monosubstituted derivatives (130, 132). With methyl iodide as the alkylating reagent, Link and Stark (133) prepared a monosubstituted sulfonium salt. Methionine 29 was strongly indicated as the principal site of modification. The activity of the derivative toward C>p was the same as RNase-A, and the methylation reaction was not affected by competitive inhibitors such as 2’(3’) -UMP. Glick e t al. (134) and Goren and Barnard (135, 136) have shown that in the alkylation reaction with bromoacetate ion the principal alkylation a t histidine occurs as described below but that in addition Met 30 is alkylated. This modification has no effect on enzymic activity and is not affected by the histidine alkylations. Phosphate and sulfate do not inhibit the reaction but pyrophosphate and 2’-CMP do. From the X-ray structure work Met 29 is the only methionine residue whose sulfur atom appears to be directly accessible to solvent. In that 129. 130. 131. 132. 133. 134. 135. 136.

E. Gross and B. Witkop, JBC 237, 1856 (1962). R. P. Neumann, S. Moore, and W. H. Stein, Biochemistry 1, 68, (1962). G. Jori, G. Galizaao, A. Marzotto, and E. Scoffone, BBA 154, 1 (1968). G. R. Stark and W. H. Stein, JBC 239, 3755 (1964). T. P. Link and G. R. Stark, JBC 243, 1082 (1968). D. M. Glick, H. J. Goren, and E. A. Barnard, BJ 102, 7 C (1967). H. J. Goren and E. A. Barnard, Biochemistry 9, 959 (1970). H. J. Goren and E. A. Barnard, Biochemistty 9, 974 (1970).

24.

BOVINE PANCREATIC RIBONUCLEASE

683

study one of the heavy atom derivatives prepared from ethylenediamine platinum I1 dichloride, or PtCl,, has a Pt-S bond to this residue (61,137). The sulfur of Met 30 is about as completely buried as it can be according to the X-ray structure. Reaction a t this site implies considerable flexibility if an opening large enough to admit the bromoacetate ion is to occur. This is especially true, if the C M sulfonium salt produced by the reaction and the accompanying structural change do not affect the enzymic activity and thus, by implication, the geometry of the active site. The problems presented by these modifications and others are considered later. Dye-sensitized photooxidation of methionine occurs selectively in strong formic or acetic acid media (131). All four methionine residues in RNase-A appear to be converted to sulfoxides under these conditions. The product still showed 13% of the initial activity in the Kunitz RNA assay a t pH 5 [see, however, Neumann et al. ( I S O ) ] . Apparently an oxygen atom can be accommodated next to each Met sulfur atom in a structure closely resembling that of the native enzyme. The 3 “buried” tyrosine residues appeared to have been largely normalized. It would be interesting to know if the binding of inhibitors returned these to the buried condition. Methionine 13 in S-peptide has been modified by oxidation to the sulfone or by conversion to a sulfonium salt with either iodoacetic acid or iodoacetamide (138). There is a dramatic lowering of the peptide-protein binding constant for all of the derivatives, but the complexes when formed appear to have nearly normal catalytic activity. The X-ray structure does not appear to permit the normal sulfur location with the sulfonium salts. Sterically, Met 13 can be moved by rotation about the carbon a-carbon ,8 bond so that the residue sticks out into the solvent. This can be done without any major change in the conformation of the rest of the peptide. Thus the active site could be maintained undisturbed while the contribution of Met 13 to the S-peptide: S-protein association would be lacking. In the treatment of RNase-A with cyanogen bromide, chain cleavage occurs a t Met 13. The peptide comprising residues 1-13, where 13 is now homoserine or homoserine lactone rather than methionine, is designated C-peptide (139). This derivative added to S-protein a t a molar ratio of 600: 1 gave between 50 and 80% of the maximum activity. 137. H. W. Wyckoff, M. Doscher, D. Tsernoglou, T. Inagami, L. N. Johnson, K. D. Hardrnan, N. M. Allewell, D. M. Kelly, and F. M. Richards, J M B 27, 563 (1967). 138. P.J. Vithayathil and F. M. Richards, JBC 235, 2343 (1960). 139. J. M.Parks, M. B. Barancik, and F. Wold, JACS 85, 3519 (1963).

684

F. M. RICHARDS AND H . W. WYCKOFF

4. Tyrosine An extensive study of the iodination of the tyrosine residues has been carried out by Scheraga and his colleagues. These studies were prompted by the spectral and titration abnormalities which indicate a division of the 6 residues into 3 “accessible” and 3 ‘(buried” in RNase-A. Iodination of RNase-A with 12 moles of I, per mole of protein a t pH 9.4 and 0” for 24 hr converted 4 of the 6 tyrosyl residues to 3,5-diiodotyrosyl residues (140). Spectral titration indicated that the 2 uniodinated residues were buried. The Kunitz RNA activity of the samples varied from 10 to 40% of RNase-A. Similar results were obtained by Donovan (141) ; the iodinating agent was the triiodide ion, and a sample of protein with about 3 diiodotyrosyl residues was obtained. A tri (iodotyrosyl) derivative was also obtained by Covelli and Wolff (142). The pK, of these 3 modified residues a t room temperature was 7.8 in 0.2 M KCl. The 3 unmodified tyrosyl groups still appear buried with very high apparent pK values. The activity of this preparation against RNA a t pH 7.5 was the same as RNase-A. Tentative identification by Donovan (141) gave 115 and 92 as modified residues, and 97 as unmodified; the other three were uncertain. Cha and Scheraga (140, 14s) identified residues 73, 76, 92, and 115 as iodinated and 25 and 97 as buried. Under slightly different conditions a t pH 6.7, Woody e t al. (144) showed that 92 would not react while 73, 76, and 115 were still iodinated. Thus the spectrally abnormal tyrosine residues are assumed to be 25,92, and 97. The inhibitor 3’-CMP was shown not to influence the iodination of Tyr 115 (145) as appears eminently reasonable from its now known position a t a considerable distance from the active site. However, the reversible reoxidation of reduced RNase to give an active enzyme appears to require that Tyr 115 be no more than monoiodinated ( 1 4 6 ~since ) derivative diiodinated a t 73, 76, and 115 that had about 75% activity before reduction regained none on reoxidation. I n a recent study of L-RNase, the derivative modified a t 73, 76, and 115, Hammes and Walz (146) showed that the kinetic parameters for CpA cleavage were essentially identical to those of the 140. C. Y. Cha and H. A . Scheraga, JBC 238, 2958 (1963). 141. L. G. Donovan, BBA 78, 474 (1963). 142. I. Covelli and J. Wolff, JBC 241, 4444 (1966). 143. C. Y. Cha and H. A. Scheraga, JBC 238, 2965 (1963). 144. R. W. Woody, M. E. Friedman, and H. A . Scheraga, Biochemistry 5, 2034 (1966). 145. M. E. Friedman and H. A. Scheraga, BBA 128, 576 (1966). 146. G. G. Hammes and F. G. Walz, BBA 198, 604 (1970). 146a. M. E. Friedman, H. A. Scheraga, and R. F. Goldberger, Biochemistry 5, 3770 (1966).

24.

BOVINE PANCREATIC RIBONUCLEASE

685

native enzyme. The turnover number for C > p hydrolysis was lowered to about 60%. Iodination of PIR (147') showed 1 residue buried, T y r 25, and all others iodinated a t least to the monoiodotyrosyl form. Pepsin-inactivated RNase also has only one abnormal tyrosyl by titration which is thus assumed to be 25. Iodination of RNase-S is very similar to RNase-A in the early stages (148). Extensive iodination leads to dissociation of the protein and peptide components. Direct iodination of S-protein indicated that all 6 tyrosyl residues were accessible, in this sense comparable to urea-denatured RNase-A. Substantial structural changes must be involved for both S-protein and PIR if Tyr 97, in particular, is to become susceptible to attack (see Section IV,B,3). Cyanuric fluoride has been used to modify tyrosine residues, substituting the phenolic hydroxyl group. A maximum of 3 residues in RNase was found to react a t pH 10.9 and 25" ( 1 4 8 a ) . However, some mystery surrounds this number, as with other estimates of accessibility, since alkaline-denatured material where all tyrosine residues are available still showed the reaction of only 3 residues with cyanuric fluoride. However, similar observations have been made on iodination in 8 M urea (14.2). At pH 9.3, Takenaka et al. (149) found that only 2 residues reacted and that 115 was not one of them. Two more reacted after alkali denaturation. Two were resistant under all conditions tested. No enzymic activity data were reported. 5. His tidine a. Photooxidation. The first definitive indication of the importance of histidine residues for the catalytic activity of RNase came with the study of Weil and Seibles (150)on the photooxidation of the enzyme in the presence of methylene blue. All activity was lost when 3 residues of histidine had been oxidized with little or no change in any other amino acids. The correlation of modification and activity loss (RNA assay) indicated that certainly 1 and perhaps 2 residues were involved in the catalytic activity. Viscosity and optical rotation measurements pointed to little or no conformational change. Methionine can be oxidized to the sulfoxide in this system. I n an investigation of the effect of pH on the photooxidation reaction, no 147. H. Fujioka and H. A. Scheraga, Biochemistry 4, 2206 (1965). 148. R. M. Cowgill, BBA 120, 189 (1966). 148a. M. J. Gorbunoff, Biochemistry 6, 1606 (1967). 140. 0. Takenakn, H. Horinishi, and K. Shibata, J . Biochem. (Tokyo) 62, 501 ( 1967). 150. L. Weil and T. S. Seibles, ABB 54, 388 (1955).

686

F. M. RICHARDS AND H .

W.

WYCKOFF

histidine loss was observed a t pH 3.8 while a t p H 6.8,80% of the histidine was oxidized. Under these same conditions 17 and 50%, respectively, of the total methionine were converted to the sulfoxide (151).The sulfoxide formation is reversible, by reduction with mercaptans for example. In the RNase-S system, early studies showed loss of potential activity of S-peptide and S-protein separately on photooxidation (152).Histidine 12 was thus immediately implicated in the activity. Subsequently, a more careful study showed that loss of His 119 resulted in loss of activity but oxidation of 105 had no effect on activity ( 1 5 3 ) .Histidine 48 was apparently not accessible to photooxidation. Photooxidation of His 12 in Speptide markedly reduced the binding constant to S-protein.

b . Haloacetic Acids. I n 1958, Barnard and W. D. Stein (154) reported the reaction of bromoacetic acid with a histidine residue and the concomitant loss of all catalytic activity. This was the start of intensive investigations of this reaction in several laboratories which was to last at least the next decade. Barnard and Stein in full papers (156, 156) documented the production of a unique species and showed that alkylation had occurred on His 119. The investigations of W. H. Stein and Moore and their colleagues were first reported in 1959 ( 1 5 7 ) . The inactivation of RNase by iodoacetate was studied. A maximum in the rate of activity loss was noted a t pH 5.5. Reaction with a methionine residue was found a t pH 2.8; a t pH 8.5-10 lysine residues were modified, but a t pH 5.5-6.0 only histidine appeared to be involved. The specific reaction required the structure of the native enzyme. Reaction with histidine was not observed under a variety of denaturing conditions (158). Iodoacetamide did not cause activity loss, or only very slow loss, or alkylate His 119 in the native enzyme a t pH 5.5. The negative charge on the carboxyl group of the iodoacetate ion was apparently essential. I n a more detailed study, Crestfield et al. (169) found that the reaction with iodoacetate a t pH 5.5 produced two monosubstituted derivatives. The major inactive product was l-carboxymethyl-His-119-RNase, 151. B. R. DasGupta and D. A. Boroff, BBA 97, 157 (1965). 152. F. M. Richards, Proc. Natl. Acad. Sci. U.S. 44, 162 (1958). 153. U. W. Kenkare and F. M. Richards, JBC 241, 3197 (1966). 154. E. A. Barnard and W. D. Stein, Biochem. SOC.Commun., 378th Meet. pp. 7-8 (1958). 155. E. A. Barnard and W. D. Stein, J M B 1, 339 (1960). 156. W. D. Stein and E. A. Barnard, JMB 1, 350 (1960). 157. H. G. Gundlach, W. H. Stein, and S. Moore, JBC 234, 1754 (1959). 158. G. R. Stark, W. H. Stein, and S. Moore, JBC 236, 436 (1961). 159. A. M. Crestfield, W. H. Stein, and S. Moore, JBC 238, 2413 (1963).

24.

687

BOVINE PANCREATIC RIBONUCLEASE

while the minor product in about one-eighth the yield was 3-carboxymethyl-His-12-RNase. The two products appeared to form simultaneously and to be mutually exclusive (160). No disubstituted material was found. To explain the result the authors postulated that His 119 and 12 were about 5 A apart and that one residue served to orient the reagent for attack the other. Both monosubstituted derivatives were inactive against RNA or C > p. Heinrikson et al. (161) have extended the study of the reaction a t pH 5.5 to a series of halo acids, investigating the effect of chain length, D and L antipodes, and the position of the halogen substituent. All of the alkylated products showed enzymic activities of less than 0.5%. A summary of the rates of reaction of the various compounds with both His 12 and 119 and free histidine are shown in Table VII. All of the reagents react more rapidly with the protein than with free histidine. TABLE VII SECOND-ORDER RATECONSTANTS FOR ALKYLATIONOF RNase-A GHISTIDINE AT pH 5.5 AND 25'09~

AND

Ribonuclease Reagent

a t His 119

at His 12

Overall

7.3 20.5 0.17 0.19 1.88 4.16 0.18 1.61 3.60 0.05

58.4 205.0 2.30 0.85 2.97 6.00 0.68 2.40 4.71 0.81 0.89 911 6.33

GHistidine

~

Iodoacetate Bromoacetate Chloroacetate L-a-Bromopropionate n,La-Bromopropionate n-a-Bromopropionate La-Bromo-n-butyrate D,M-Bromo-n-bu tyrate D-a-BromMt-butyrate D ,ta-Bromovalerate D,M-Bromocaproate 8-Bromopyruvate 8-Bromopropionate

51.1 184.5 2.13 0.66 1.09 1.84 0.50 0.79 1.11 0.76 0.89 91 1 6.33

0.086 0.0027 0.0027 0.0028

0.0008 0.0023

Taken from Heinrikson et al. (161). Reactions were carried out in the dark in 0.10 M sodium acetate buffer a t pH 5.50. With the enzyme the ionic strength of the solutions varied from 0.11 to 0.13, depending upon the concentration of reagent used with histidine the ionic strength was 0.32. 6

160. A. M. Crestfield, W. H. Stein, and S. Moore, JBC 238, 2421 (1963). 161. R. L. Heinrikson, W. H. Stein, A. M. Crestfield, and S. Moore, JBC 240, 2921 (1965).

688

F. M. RICHARDS AND H . W. WYCKOFF

However, the relative rates with His 12 and 119 vary widely depending on the structure of the reagent. The D antipodes generally favor reaction at His 12, the L compounds a t His 119. No convincing correlation of all these facts with the X-ray structure has yet been made. In view of the fact that sulfate or phosphate ions seriously interfere with the alkylation reaction (160), the X-ray structures determined in the presence of these ions may be misleading. However, Bello and Nowoswiat have shown (69) that with RNase-A in crystalline form the alkylation with bromoacetic acid is comparable, but not identical, to that found in dilute solution. The uncharged reagent, iodoacetamide, will react with RNase-A albeit slowly (162). One of the principal products is 3-CAM-His-12-RNase. The pH dependence of the rate again shows a maximum near 5. No evidence could be found for reaction a t His 119. A fully active product was obtained containing an alkylated methionine residue, but the location of this residue was not determined. The C > p activity of the His 12 derivative was 1.3% that of RNase-A. The sample used for this measurement had been highly purified, and contamination with RNase-A a t the 1% level was highly unlikely. Alkylation a t pH 8.5 shows reduced rates of reaction a t the histidine residues but significant substitution a t lysine, particularly Lys 41 (118). The histidine reactions show the same general stereospecificity as found at pH 5.5. The inactive Lys 41 derivatives (25, 26, and 27 of Table VI) show alkylation patterns of His 12 and 119 a t pH 5.5 which are similar to those of RNase-A although with some differences in detail. When Lys 1 and 7 are acetylated in RNase-S the alkylation pattern with iodoacetic acid is not affected. When P I R is used the alkylation of His 119 is nearly abolished but that a t His 12 is accelerated (163). The probable interaction of Asp 121 with His 119 may be important in the alkylation reactions observed in the native enzyme and the various lysine derivatives. In P I R this interaction has, of course, been removed. When RNase-S was treated with iodoacetate a t pH 6, both inactivation and histidine modification occurred (164). The modified histidine was in S-protein and was assumed to be His 119 since the sole product on analysis was l-CM-His. In the absence of S-peptide only methionine modification occurred in S-protein. The loss of potential activity probably resulted from the reaction of the second of the two modifiable Met residues. The location of these residues in the sequence was not established. 162. R. G. Fruchter and A. M. Crestfield, JBC 242, 5807 (1967). 163. M. C. Lin, W. H. Stein, and S. Moore, JBC 243, 6167 (1968). 164. P. J. Vithayathil and F. M. Richards, JBC 236, 1386 (1961).

24.

BOVINE^ PANCREATIC RIBONUCLEASE

689

S-Protein was used to show that 1-CM-His-119-RNase-A still had the NH,-terminal “tail” closely associated with the rest of the molecule as no activity was regenerated on mixing. Prior denaturation of the derivative produced full activity when mixed with S-protein.

c. lodination. Covelli and Wolff (14.2) have shown that conditions leading to the iodination of 3 tyrosyl residues in RNase-A also produce one iodohistidine residue, presumably His 119. This reaction is prevented by competitive inhibitors such as 2’(3’) -CMP while the tyrosyl iodination is not significantly effected. In 8 M urea only 5 of the 6 tyrosine residues and 2 of the 4 histidines were iodinated. This observation may indicate some residual structure in this solvent. Moniodination of His 12 in S-peptide not only resulted in complete loss of potential activity but markedly reduced the binding constant to S-protein. d. Diazonium-1H-Tetrazole. Horinishi et al. (165) reported that with sufficient diazonium-1H-tetrazole ( D H T ) all 4 histidines react. Some were protected from modification by pyrimidine nucleotides. The curves of extent of reaction as a function of concentration of D H T show rather sharp breaks and plateaus. The authors attributed these to reaction a t specific residues although no explanation for this unique and unexpected behavior is given, nor did they report the isolation or identification of any individual products. (Table VIII) . 6. Arginine Arginine residues are very resistant to most of the usual reagents used for chemical modification. King (166) has reported the conversion of these residues to 6-N- (2-pyrimidinyl) ornithine by treatment with malonaldehyde in 10 N HCI. Essentially complete modification of the 4 arginine residues in RNase was obtained. Peptide bond cleavage and disulfide interchange also occurred, however, and no conclusions are warranted on the relation of the properties of the product to the actual arginine modification. A more promising approach is the use of phenylglyoxal as reported by Takahashi (16’7). Two reagent molecules combine with one guanidino group a t neutral pH and room temperature. The reaction appears to be reversible in the absence of excess reagent a t 37”.Modification of 165. H. Horinishi, 0. Takenaka, and K. Shibeta, ABB 113, 371 (1966). 166. T. P. King, Biochemistry 5, 3454 (1966). 167. K. Takahashi, JBC 243, 6171 (1968).

690

F. M. RICHARDS AND H. W. WYCKOFF

TABLE VIII SUMMARY OF THIOETHER, PHENOL,AND IMIDAZOLE DERIVATIVES ~~~

~

Activity (%) of RNase-A No.

Derivative or abbreviation

Modified residues

RNA

C >p (U > p)

No. of buried Tyr residues

100 100 NR

NR NR 2

NR

3

3

<0.5

NR

1.3

NR

Ref.

RNase-A 1 2 3

(CH3)l (CM)i (Diiodo)r

4

(Diiodo)t

5

l-CM-His-119

6

3-CM-His-12

NR Met 29 Met 30 100 1040 Tyr 73, 76, 92, 115 Tyr 73, 76, 100 115 His 119 0 (Met 30?) His 12 NR

7

3-CAM-His-12

His 12

NR PIR

8

(Mono and diiodo)

All Tyr except 25

0

0

1

(147)

2-3 Arg residues in RNase-A causes a marked loss of activity. The half-time for the loss is about 5 min a t pH 8 and slower in more acid solution. Arginines 39 and 85, the most accessible arginines, are the principal sites of modification, and the activity loss appears to correlate with the reaction a t Arg-39. Arginines 10 and 33 seem to be quite unreactive toward the reagent. (Deamination of peptide a-amino groups occurs with this reagent.)

7. Cystine-Disulfide Groups Over the years it has been established that the usual samples of RNase have 4 disulfide groups and no free SH groups. In the middle 1950’s Ledoux claimed that there were SH groups present [one of the latest papers ( 1 6 8 ) ] . Much of his work could not be repeated a t that time, and claims for the importance of SH to activity do appear to be wrong. However, more recent work has shown that some reduction of the SS groups can occur with little or no loss of activity and criticisms of some parts of the earlier work may be unjustified. 168. L. Ledoux, BBA 23, 121 (1957).

24.

BOVINE PANCREATIC

RIBONUCLEASE

691

a. Oxidation. Oxidation with performic acid leads smoothly to the quantitative conversion of the 8 sulfur atoms in the 4 disulfide groups to 8 sulfonic acid groups (169). The four methionine residues are simultaneously converted to sulfones. The product, Ox-RNase, has no enzymic activity and appears to approximate very closely to a random coil in dilute solution ( 1 6 9 ~ ) . b. Reduction and Protection. Quantitative reduction of the SS groups only occurs under denaturing conditions. The solvent commonly used has been 8 M urea. Complete reduction in 8 M urea to give 8 SH groups can be obtained with thioglycollate (170) with caution (171), with sodium borohydride (172),with sulfite (173), and with mercaptoethanol (174). Electroreduction can also be effected a t the dropping mercury electrode (176). The reduced protein is easily reoxidized by air. For stability the SH groups must be protected. Irreversible alkylation is easily achieved with iodoacetate to give S-carboxymethylcysteine derivatives (170, 172, 174). If ethyleneimine is used an S-aminoethylated derivative is obtained (176). These new groups will serve as cleavage sites for trypsin. Reversible blocking of SH groups can be obtained through the formation of unsymmetrical disulfides with a variety of sulfenyl halides, for example, 4-nitrophenylsulfenyl chloride (177), SS cleavage with trisodium phosphorothioate (178), or protection with mercury compounds such as p-mercury benzoic acid (179). I n each case the free SH groups can be regenerated with mercaptans. Early studies of partial reduction (180, 181) strongly indicated that species with a t least 1 or 2 SS bonds reduced did retain some catalytic activity. More recently, Neumann et al. (182) have been able to prepare 169. C. H. W. Him, JBC 219, 611 (1956). 169a. W. F. Harrington and M. Sela, BBA 31, 427 (1959). 170. M. Sela, F. H. White, Jr., and C. B. Anfinsen, BBA 31, 417 (1959). 171. F. H. White, Jr., JBC 235, 383 (1960). 172. 5. Moore, R. D. Cole, H. G. Gundlach, and W. H. Stein, Proc. 4th Intern. Congr. Biochem., Vienna, 1968 Vol. 8, p. 52. Pergamon Press, Oxford, 1960. 173. J. L. Bailey and R. D. Cole, JBC 234, 1733 (1959). 174. A. M. Crestfield, 5. Moore, and W. H. Stein, JBC 238, 622 (1963). 175. R. Cecil and P. D. J. Weitzman, BJ 93, 1 (1964). 176. B. V. Plapp, M. A. Raftery, and R. D. Cole, JBC 242, 265 (1967). 177. A. Fontana, E. Scoffone, and C. A. Benassi, Biochemitry 7, 980 (1968). 178. H. Neumann, R. F. Goldberger, and M. Sela, JBC 239, 1536 (1964). 179. C. B. Anfinsen and E. Haber, JBC 236, 1361 (1961). 180. M. Sela, F. H. White, Jr., and C. B. Anfinsen, Science 125, 691 (1957). 181. H. Resnick, J. R. Carter, and G. Kalnitsky, JBC 234, 1711 (1959). 182. H. Neumann, I. Z. Steinberg, J. R. Brown, R. F. Goldberger, and M. Sela, European J . Biochem. 3, 171 (1967).

692

F. M. RICHARDS AND H. W. WYCKOFF

a unique species with 4 phosphorothioate groups attached to the 4 sulfur atoms of the disulfide groups of residues 65-72 and 58-110. This derivative has full activity toward RNA, is even more active than the native enzyme toward C > p, still has 3 abnormal tyrosine residues, and is indistinguishable from RNase-A in reaction with specific antisera. This is an extraordinary observation when one examines the X-ray structure. The phosphorothioate group represents the addition of 5 atoms in a dense cluster with a negative charge to each original S atom in the SS bond. The changes required to accommodate these groups are very large, especially at the 58-110 bond which is not as exposed as 65-72. How these changes can be accommodated without serious disruption of the structure of the enzyme is not yet clear. The 65-72 octapeptide loop does permit considerable flexibility, but the same is not obviously true of the environment of SS bond 58-110, and the denine binding site includes the 65 sulfur atom. Confirmation of this possibility comes from the work of Steinberg and Sperling. From the completely reduced protein they have produced a derivative containing 1 mercury atom bridging each of the 4 pairs of sulfur atoms (183). The resulting molecule had two abnormal tyrosine residues and reacted with antiserum to RNase. In a more limited modification 1 mercury atom was introduced specifically a t the 65-72 SS group (184). This derivative was fully active and nearly identical to RNase-A. Ganther and Corcoran (186) have reported the introduction of selenium into SS bonds to produce compounds of the type R-S-Se-&R. With reduced RNase a t pH 2, 2 moles of selenium can be introduced per protein molecule. Other derivatives have differing amounts of selenium. The monomeric derivatives have some properties similar to RNaseA but only a few per cent of the original enzymic activity. The location of the selenium atoms has not been reported. Reports on the action of reduced glutathione give results quite different from the above. The bulky mercaptan does not reduce RNase at all at room temperature. As the thermal transition temperature is approached the reduction appears to be all or none with no evidence of partially reduced intermediates (186). In a study of the recovery of activity after denaturation in 8 M urea, Kim and Paik (187)reported 183. I. 2. Steinberg, and R. Sperling, Conform. Biopolymers, Papers Intern. Symp., Univ. Madras, 1967 Vol. 1, 215. Academic Preea, New York, 1967. 184. R. Sperling, Y, Burnstein, and I. 2. Steinberg, Biochemistry 8, 3810 (1969). 185. H. E. Ganther and C.Corcoran, Biochemistry 8, 2557 (1969). 186. B. E. Davidson and F. J. R. Hird, BJ 104,480 (1907). 187. S. Kim and W. I(.Pa&, BJ 106, 707 (1968).

24.

693

BOVINE PANCREATIC RIBONUCLEASE

that reduced glutathione inhibits this procedure by reduction of a single

S S bond. I t is not yet known which bond is reduced. Well defined disulfide derivatives are summarized in Table IX. c. Reoridation and Refolding. One of the most important general observations made in the ribonuclease system is that the fully reduced, random coil polypeptide can be reoxidized in air to give the native enzyme with full enzymic activity. The crucial importance of this fact in consideration of protein biosynthesis is well known (188). The reaction has been studied extensively in a number of laboratories, but the initial observations were made by Anfinsen and his colleagues who also have done the bulk of the subsequent work. There are two basic parts to the problem: (1) to demonstrate that the fully reduced material can be denatured to the point where no residual structure exists, and (2) to demonstrate that the reformed enzyme is identical to the starting material before reduction. Part 1 requires the demonstration that reduced RNase, or some derivative from which it can be directly derived (179), is a true random coil with no time-independent secondary or tertiary structure. The physical methods appear to indicate this, but the evidence by necessity is negative. The methods fail to give positive evidence for the existence of special structures, thus within the relevant limits of accuracy it is conTABLE IX SUMMARY OF CYSTINE DERIVATIVES ~

~~

Activity (%) of RNase-A No.

Derivative or abbreviation

Modified groups

RNA

C>p (U > p)

Physical state

Ref. ~~

1

2 3

4

5

6 7

Ox-RNase, sulfonic acid and sulfone Red-RNase free SH CM-Red-RNase (8carboxymethy1)s 8-PSRNase (phosphorothioate)s CPd-RNase (phosphorothioate), Hg-RNase (Hg)l-RNase

All Cys All Met All c y s All c y s

0

0

Random coil (169,

0 0

0 0

Random coil

All Cys

0

0

100

220

Cys 65, 72, 58, 110 Cya, 65-72 All Cys

169a)

(169a) (188)

“Native”

(18.8)

(1844) (183)

188. C. B. Anfinsen, Basic Probl. Neoplastic Disease, Symp, New York, 196.8 p. 112. Columbia Univ. Press, New York, 1962.

694

F. M. RICHARDS AND H. W. WYCKOFF

cluded that such structures do not exist (189).The last vestiges of doubt on this point were certainly removed by the total synthesis of the enzyme by two separate methods (190, 191). The final step in each case is the reoxidation to get the active enzyme. Part 2 again relies on a battery of comparative tests to establish the correspondence of the reduced-reoxidized product and the native enzyme (192). Chromatographic behavior, enzymic activity toward RNA, C > p U > p, UV spectra, ORD, viscosity, and reaction to antisera to RNase-A were all identical with or very similar to those of the starting enzyme, while the reduced material was markedly altered in all of these properties for which tests were possible. It was possible to crystallize the reoxidized material. The crystals were identical in space group, lattice constants, and general intensity distribution of the X-ray reflections within the normal limits of variation to the crystals of RNase-A from the same solvent (193). Recent circular dichroism studies show small differences near 240 nm in reoxidized material (194, 195) interpreted as arising from change in the chirality of one disulfide or the environment of one tyrosine residue. The kinetics of the reoxidation process both in terms of enzymic activity and the various physical parameters point to the incorrect pairing of SS groups, probably intermolecular, as the first stage in the process. This is followed by disulfide exchange in a resorting process with the normal monomeric enzyme as the final product (196, 197). Kauzmann (198) has pointed out that for a protein with 8 distinguishable cysteine residues forming 4 SS bonds there are 105 possible isomers. The relative probability of the correct structure is thus low. The fact that high yields, approaching loo%, can be obtained indicates that the structure of the native enzyme is the thermodynamically stable form, or a t least, the most stable of the kinetically accessible forms. 189. F. H. White, Jr., JBC 236, 1353 (1961). 190. B. Gutte and R. B. Merrifield, JACS 91, 501 (1969). 191. R. Hirschmann, R. F. Nutt, D. F. Veber, R. A. Vitali, S. L. Varga, T. A. Jacob, F. W. Holly, and R. G . Denkewalter, JACS 91, 507 (1969). 192. F. H. White, Jr., JBC 236, 1353 (1961). 193. J. Bello, D. Harker, and E. de Jarnette, JBC 236, 1358 (1961). 194. T. Isemura, K. Yutani, A. Yutani, and A. Imanishi, J . Biochem. (Tokvo) 64, 411 (1968). 195. M. N. Pflumm and S. Beychok, JBC 244, 3982 (1969). 196. C. B. Anfinsen, E. Haber, M. Sela, and F. H. White, Jr., Proc. Natl. Acad. Sci. U . S. 47, 1309 (1961). 197. C . J. Epstein, R. F. Goldberger, D. M. Young, and C. B. Anfinsen, ABB Suppl. 1, 223 (1962). 198. W. Kauamann, Proc. Symp. Falmouth, Mass., 1968 p. 93.

24. BOVINE

PANCREATIC RIBONUCLEASE

695

A variety of different derivatives of RNase have been studied with respect to structure and to recovery of activity on reoxidation: polyalanyl and polytyrosyl (199) and succinyl, phthalyl, butyryl, caproyl, poly-L-valyl derivatives (116) all showed substantial recovery of activity. The introduction of two 5-dimethylamino-1-naphthalene sulfonyl groups markedly reduced the recovered activity (200).Denaturing agents such as 8 M urea and 4 M guanidine produce mispairing as do uncharged phenolic compounds (201). All of these experiments together suggest that surface charge distribution is not very important in producing the unique folded structure but that certain nonpolar interactions, especially those involving ring compounds, may be critical. Scoffone et al. (202) found the CD spectrum of 4-PS-RNase to be very similar to native RNase-A and different from the fully reduced enzyme red-RNase. Thus SS bonds 26-84 and 40-95 which are present in 4-PS-RNase may be crucial for the correct final folding and may be close to the folding nucleation region. A rearranging enzyme has been found in microsomes which is a general catalyst for disulfide interchange reactions required in the reoxidation process (203-205). This enzyme may be very important biologically but is used here solely as a reagent for the study of RNase and will not be considered in detail. Reoxidation of reduced S-protein results in potential activity recovery but only about 20% of that seen with RNase-A ($06).I n the presence of S-peptide the reoxidation of S-protein leads to much higher levels of activity. The addition of the rearranging enzyme to a solution of Sprotein results in the immediate drop in potential activity to about 20% of the starting value (207).If S-peptide is added to this mixture the activity rapidly increases and approaches 90-100%. If the SS bonds were totally random in reoxidized S-protein the activity should have been much less than 20%. There is a bias in favor of a n approximately correct structure; therefore, the actual distribution of S-S bonds in reoxidized S-protein would be extremely interesting and might shed 199. C. B. Anfinsen, M. Sels, and J. P. Cooke, JBC 237,1825 (1962). 200. F. H. White, Jr., JBC 239, 1032 (1964). 201. E. Haber and C. B. Anfinsen, JBC 237, 1839 (1962). 202. E. Scoffone, F. Marchiori, R. Rocchi, A. Scatturin, and A. M. Tamburro, Proc. 10th European Peptide Symp. (in press). North-Holland Publ., Amsterdam, 1971. 203. R. F. Goldberger, C. J. Epstein, and C. B. Anfinsen, JBC 239, 1406 (1964). 204. P. Venetianer and F. B. Staub, Acta PhysMl. Acad. Sci. Hung. 27, 303 (1965). 205. P. Venetianer, Nature 211, 643 (1966). 206. E. Haber and C. B. A n h e n , JBC 236,422 (1961). 207. I. Kato and C. B. Anfinsen, JBC 244, 1004 (1969).

696

F.

M . RICHARDS AND H. W. WYCKOFF

considerable light on the nucleation process. The intermolecular aggregation that can occur would have to be avoided in such a study.

8. Serine and Threonine Aliphatic hydroxyl groups cannot normally be selectively modified except in certain special cases such as the serine proteinases. In anhydrous formic acid, the N,O-acyl migration that occurs in strong sulfuric or phosphoric acid apparently does not occur. Instead there is formylation of the serine and threonine residues (208). Enzymically inactive aggregates are produced, but the reaction is reversed in aqueous solution a t neutral pH and the activity returns. Josefsson reported the introduction of 29 formyl groups in RNase (209) as compared to the total of 25 Ser and Thr residues. This identification of reaction sites is not clear, however, since the number of formyl groups introduced into lysozyme far exceeded the Ser-Thr total.

9. Intramolecular Cross-linking Marfey et al. (210,211) have treated RNase with the bifunctional reagent, 1,5-difluoro-2,4-dinitrobenzene.The products were separated chromatographically, and three fractions of modified monomeric material were obtained showing absorption maxima near 343 and 428 nm, the wavelengths of maximum absorption of N,N'-dinitrophenylene-bis-lysine. The enzymic activities, C > p, of these fractions is reported to range from 15 to 49% that of RNase-A. The fraction obtained in largest amount, with 15% activity, was examined in detail. The cross-link found involved lysine residues 7 and 41. Inorganic phosphate had been carefully removed from the starting RNase sample so that reaction a t Lys 41 was not unexpected. The chemical and activity studies were actually done on separate preparations, and the column separations were not clean. Thus the activity of this specific cross-linked material is uncertain. Hartman and Wold (212) have used a bifunctional imido ester, dimethyl adipimidate, as a cross-linking reagent for an enzyme sample containing phosphate ion to protect Lys 41. The monomeric component of the reaction mixture was apparently itself a mixture of species which was not further fractionated. The total fraction had slightly reduced RNA activity but the same V , and a decreased K , for the activity 208. J. L. Rabinowitz, Acta Chem. Scand. 13, 1463 (1959). 209. L. Josefsson, Acta Chem. Scand. 19, 2421 (1966). 210. P. S. Marfey, H. Nowak, M. Uziel, and D. A. Yphantis, JBC 240, 3264 (1965). 211. P. S. Marfey, M. Uziel, and J. Little, JBC 240, 3270 (1965). 212. F. C. Hartman and F. Wold, Biochemistry 6, 2439 (1967).

24.

BOVINE PANCREATIC RIBONUCLEASE

697

against C > p. From the analysis of peptides from this fraction, crosslinks were firmly established between Lys 31 and 37 and between Lys 7 and 37. Because of the unresolved mixture, the cause of the changed kinetic parameters cannot be specified. Ozawa (213) has reported the probable formation of an intramolecular cross-link with a diisocyanate. A complete investigation of the product was not made. It is interesting that all of the cross-links found in the above studies appear to be sterically permitted on the basis of the X-ray structure, although some movement of the lysine side chains from their reported positions is required. The reason for almost total loss of activity when monofunctional reagents react a t Lys 41 and the possible maintenance of substantial activity when the Lys 7-41 dinitrophenylene cross-link is formed is not clear. Preliminary evidence from the X-ray structure study of the D N P Lys-41-RNase-S (120) indicates small movements of many parts of the molecule in comparison with RNase-S, but these motions have not yet been analyzed in detail. They are substantially reversed by 3’-CMP binding. If they should be responsible for the activity loss, then the cross-link may prevent or reverse these changes. 10. Other Reagents

A large number of reagents have been tried on ribonuclease, and the enzyme serves frequently as a test protein. Some of the reagents not described above are listed in Table X (214-222). In general the products of the reactions either have not been well characterized or the sites of modification in the sequence have not been established.

C. CHEMICAL SYNTHESIS AND S-PEPTIDESUMMARY The dramatic event in 1969 was the total synthesis of RNase-A and the completion of the synthesis of RNase-S. Gutte and Merrifield (190) used the solid phase method to synthesize 85 mg of a protein that had H. Ozawa, J. Biochem. (Tokyo) 62, 419 (1967). I. M. Klotz and R. E. Heiney, ABB 96, 605 (1962). S. R. Dickman, R. B. Kropf, and C. M . Proctor, JBC 210, 491 (1954). I. M. Klotz, E. C. Stellwagcn, and U. H. Stryker, BBA 86, 122 (1964). W. F. Goebel, P. K. Olitsky, and A. C. Saenz, J. E x p t l . Med. 87, 445 (1948). C.-L. Chou, Y. K. Sun, K.-C. Hsu, and Y.-T. Du, Sheng W u Hsueh YU Sheng W u W u Li Hsueh Pa0 3, 169 (1963). 219. A. V. Luisada-Opper and H. Sobotka, Immunochemistry 2, 127 (1965). 220. G. Taborsky, JBC 234, 2915 (1959). 221. G. H. Beaven and W. B. Gratzer, BBA 168, 456 (1968). 222. W. B. Melchior and D. Fahrney, Biochemistry 9, 251 (1970). 213. 214. 215. 216. 217. 218.

F.

698

M. RICHARDS AND H. W. WYCKOFF

TABLE X ADDITIONAL REAGENTS TESTED ON RIBONUCLEASE ~

No. 1 2

Reagent S-Acetylmercaptosuccinic anhydride Xanthy drol

4

Dimethylamino naphthalene sulfonyl chloride Periodic acid

5 6 7

p-Nitrophenacyl bromide p-Diazobenzoic acid 1,3-Diphosphoimidazole

8

Tetrani trometharie Ethoxyformic anhydride

3

9

Comments Modifies Lys residues and introduces SH groups 5.5 groups introduced; residual activity <2y0

Ref.

(8161

pH 5.1, 25", activity loss complete in 24 hr Activity loss Monophosphorylated products isolated; one with low RNA activity; Lys probably modified 3 Tyr residues modified Modifies His residues and amino groups at pH 4

the enzymic specificity of RNase-A and that closely resembled natural RNase-A in Michaelis constant on RNA, amino acid composition, electrophoretic and chromatographic behavior, and peptide maps of tryptic digests. The purified synthetic material had specific activities of 13% on RNA and 24% on C > 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.

BOVINE PANCREATIC RIBONUCLEASE

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.

BOVINE PANCREATIC RIBONUCLEASE

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

BOVINE PANCREATIC RIBONUCLEASE

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

F. M. RICHARDS AND H. W. WYCKOFF

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.

BOVINE PANCREATIC RIBONUCLEASE

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.

BOVINE PANCREATIC RIBONUCLEASE

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 <-amino groups 10.2, the imidazole groups 6.5, the a-amino and a-carboxyl groups could not be independently established but were assumed to have values of 7.8 and 3.75, respectively; the 8and 8-carboxyl groups were divided into two groups with values of 4.0 and 4.7. All of these values are within the ranges normally found for these various classes of functional groups. The effects of changing ionic strength were adequately accounted for by the single electrostatic factor o of the smeared charge model; numerical values of 0.112, 0.093, and 0.061 were found appropriate for ionic strengths 0.01, 0.03, and 0.15 M, respectively. In a study of the acid titration region a t high salt concentration, Bull and Breese (276, 276) concluded that a wide range of intrinsic p K values for the carboxyl groups were required varying from 3.3 to 5.0. They also divided the imidazole region into four individual groups with values of 5.8, 6.6, 6.7, and 6.8. The acid-induced transition as monitored by change in UV absorbance indicates one or two groups titrating in the pH range of 1.5-3.0 (277). Direct measurement a t these low pH values by electrometric titration is very difficult, and the most acidic group in the protein may have been missed in the earlier studies. In recent studies employing NMR it was possible to identify signals from the C2 protons of each histidine separately (see Table XIV). The thermodynamic parameters are also provided for each histidine in 271. 272. 273. 274. 275. 276. 277.

M. A. Rosemeyer and E. M. Shooter, BJ 69, 28P (1968). I. D. Raacke and C. H. Li, BBA 14, 290 (1954). I. D. Raacke, ABB 62, 184 (1966). C. Tanford, J. D. Hauenstein, and D. G. Rands, JACS 77, 6409 (1955). H. B. Bull and K. Breese, ABB 110, 331 (1905). H. B. Bull and K. Breese, ABB 117, 106 (1966). J. Hermans, Jr. and H. A. Scheraga, JACS 83, 3293 (1961).

712

F. M. RICHARDS AND H. W. WYCKOFF

TABLE XIV IONIZATION CONSTANTS FOR INDIVIDUAL HISTIDINE RESIDUES~ Acetate buffer 0.2 Mc

RNase-A inb NaC1, I = Residue No. 12 48 105 119

0.1 M

0.2 M

0.4M

5.6

6.1

6.3

6.4 5.1

6.5 5.8

6.6 6.1

-

-

-

RNase-A RNase-S 6.2 (6.4) 6.7 5.8

6.7 (6.4) 6.7 6.3

l-CM-His119-RNase

3-CM-His12-RNase

6.7 (6.4) 6.7 6.9

6.7d (6.4) 6.7 7.6

Ligands may have a large effect on the values for His 12 and 119. Commercial samples assumed free of phosphate ion were used. Temperature 30" (280). Temperature 32" (278, 2881). It is not clear from the X-ray structure how N1 of His 12 can have a pK. value so nearly normal. (1

RNase-A (278). Histidine 105 is normal, but, by a sharp change in AHioniZation, His 12 and 119 give evidence for a thermal transition a t

30"40".This transition is not seen in UV or ORD data but is seen in chromatographic behavior (279). The normal transition observed in absorbance, ORD, and viscosity occurs a t about 60". 11. Hydrogen Exchange A long series of studies of the rates of exchange of protons have followed the introduction of this technique by Linderstrom-Lang (282). Experimentally this has been approached by the original cryosublimation procedure, the use of rapid gel filtration separations, or spectroscopically in the infrared region. Initially deuterium and more recently tritium have been used to follow either the "in" or "out" exchange. Observed rates slower than those seen in model compounds have been assumed to reflect the secondary and tertiary structure of a protein and the stabilities of these structures. This work has been the subject of an extensive review by Hvidt and Nielsen (282a). Some of the reported 278. G. C. K. Roberts, D. H. Meadows, and 0. Jardetsky, Biochemistry 8, 2053 (1969). 279. C. H. W. Hirs, Brookhaven Symp. Biol. 15, 154 (1962). 280. H. Ruterjans and H. Witzel, European J . Biochem. 9, 118 (1969). 281. D. H. Meadows, 0. Jardetsky, R. M. Epand, H. H. Ruterjans, and H. A. Scheraga, Proc. Natl. Acad. Sci. U.S. 60, 766 (1968). 282. R. Linderstrom-Lang, Chem. SOC. (London), Spec. Publ. 1-20 (1955). 282a. A. Hvidt and S. 0. Nielsen, Advan. Protein Chem. 21, 287 (1966).

24.

713

BOVINE PANCREATIC RIBONUCLEASE

exchange data are presented in Table XV (683-290).The variability has been ascribed to differences in samples and to differences in the pH and temperature history of the preparation actually studied. Part of the difficulty centers on the significance of the conformational transitions. TABLE XV EXCHANGE SUMMARY HYDROGEN (Total exchangeable protons a t pH 4.8 = 245; total main chain exchangeable protons = 119) ~~

Number of protons by exchange class Reference: Methoda: Isotope:

(28s)

(284)

(286)

A D

B D

A D

(286) A D

(287)

(288)

(289)

B D

A T

C T

pH and temperature Class

Exchange half-time

4.7 (0')

(<37")"

4.7 (0")

4.8 (<38")

4.5 4.2 ( ~ 2 5 " ) ~(0")

4.7 (4")

~~~

(Film)

I I1 I11 IV

Very fast 5-10min 2-6 hr Veryslow

Class

Exchange half-time

2.4

3.9

6.0

9.1

12

I I1 I11 IV

Veryfast 5-10min 2-22 hr Very slow

153 153 67 39

162 162 50 49

197 197 40 23

210 210 26 7

213 213 7 2

(52) 15 45

40

50 33

75 25 45

80 30 14

45 70

40 73

PHb

a

Here A refers to cryosublimation, B to infrared absorption, and C to gel filtration.

* Data of Leach and Hill (2.990).

+

Amide amino only. Peptide NH only. 283. 284. 285. 286. 287. 288. 289. 290.

A. Hvidt, BBA 18, 306 (1955). G. H. Haggis, BBA 23, 494 (1957). A. Stracher, Compt. Rend. Trav. Lab. Carlsberg 31, 468 (1960). C. L. Schildkraut and H. A. Scheraga, JACS 82, 58 (1960). E. R. Blout, C. deLoze, and A. S. Asadourian, JACS 83, 1895 (1961). S. J. Leach and P. H. Springell, Australian J . Chem. 15, 350 (1962). S. W. Englander, Biochemistry 2, 789 (1963). S. J. Leach and J. Hill, Biochemistry 2, 807 (1963).

714

F. M. RICHARDS AND H . W. WYCKOFF

The cryosublimation procedures are all “exchange out” processes which assume that complete “in exchange” has previously been obtained. Any nonexchanged protons will not be seen and will automatically be classed as “very fast” whereas in reality had the in exchange been complete these protons might have appeared as “very slow.” Complete clarification would appear to require optimally a combination of infrared and tritium-Sephadex procedures to generate confidence in identification of groups and stoichiometry a t each stage. If and when total resolution of the NMR spectrum becomes possible the complete specification of the proton exchange process will follow. The very slowly exchanging hydrogens reported by Schildkraut and Scheraga (286), a “hard core” of 20, were examined by Hermans and Scheraga (2991) in the infrared. They were identified as NH, rather than OH atoms, and thus were considered to be involved in main chain hydrogen bonds. The X-ray structure of RNase-S shows 53 main chain hydrogen bonds. In addition there are 20 main chain NH and 9 side chain polar atoms which appear to be inaccessible in the static structure. This total of 82 represents roughly the maximum number of slowly exchangeable hydrogen atoms to be expected and should be compared with the 113 in Class I11 and IV found by Englander (Table XV). The rationale here is that any group with an exchange rate as slow as hours would probably appear totally inaccessible in an examination of the resting molecule. The difficulties and dangers in the analysis of simultaneous exponential processes are discussed by Laiken and Printz (292). For ordinary experimental error any actual situation can be approximated by about three classes with significantly different rates. 12. Ultraviolet Absorption Spectra Ribonuclease contains no tryptophan. The absorption near 280 nm is almost entirely resulting from the 6 tyrosine residues. The ionization of tyrosine produces a marked shift to longer wavelengths in the absorption spectrum. The ionization can be monitored near 295 nm. Shugar (293) was the first to point out the abnormal behavior of 3 of the tyrosine residues on alkaline titration. Three titrate normally with apparent p K values near 10, but three do not titrate until much more alkaline pH values have been reached and irreversible alkaline denaturation has set in. Some typical spectra and difference spectra are 291. J. Hermans, Jr. and H. A. Scheraga, JACB 82, 5166 (1960). 292. S. L. Laiken and M. P. Prints,Biochemistry 9, 1647 (1970). 293. D. Shugar, BJ 5% 142 (1952).

24.

715

BOVINE PANCREATIC RIBONUCLEASE

shown in Fig. 9 and an alkaline spectral titration in Fig. 10. The molar extinction coefficient a t 278 nm a t neutral pH and room temperature is 9700 cm’/mole (994) or a solution of the enzyme a t a concentration of 1 mg/ml will have an optical density of 0.71. The change in molar extinction a t 295 nm on ionization of 1 tyrosyl residue is about 2630 (274) compared to 2300 for free tyrosine. Alteration in the position of the “step” in the alkaline titration curve can be interpreted in terms of the number of normal tyrosine residues showing this absorbance change on ionization. The general effects to be expected on altering the polarizability of

C””’.“”’ .-..

1

-1

12.0

2

Wovelength

(1)

(b) FIG.9. (a) Ultraviolet absorption spectra of L-tyrosine ethyl ester HC1 (TEE) in (1) 0.2 M KCl-H,O and (3) 0.2 M KC1-ethylene glycol, and of RNase in (2) 0.2 M KCl-H,O, and (4) 0.2 M KClethylene glycol. Reproduced from Sage and Singer (294). (b) Difference spectra a t pH 6.5 relative to the spectrum of RNase. The PIR; changes represent decreases in absorbance a t the wavelength shown: (-) final peptic digest of RNase or P I R ; (--.**-) RNase in 8 M urea; (---I RNase in 8 M urea 0.003 M phosphate buffer. Reproduced from Sela et al. (,W?). ( . a * )

+

294. H. J. Sage and S. J. Singer, Biochemistry 1, 305 (1962).

716

F. M. RICHARDS AND H. W. WYCKOFF

PH

FIG.10. Alkaline spectral titrations. Molar absorbance change at 295 nm vs. pH: (0) data from Bigelow (297), line from Tanford e t al. ( 2 7 4 ) . ( f ) PIR and ( 0 ) Ox-RNase. Reproduced from Bigelow (297).

the environment of the various chromophoric groups found in proteins were described by Yanari and Bovey (295). A change in the environment of the buried tyrosine residues caused by a variety of denaturing conditions causes a shift in the wavelength of maximum absorption from 277.5 to 276.0 nm. This shift produces a difference spectrum with a principal minimum at 287 nm and a secondary minimum a t 280 nni. The 287-nm band can be conveniently monitored as a measure of structural change. From studies at acid pH, Scheraga concluded that each of the abnormal tyrosine residues was close to a carboxyl group (298, 299). The identification of the residue pairs was approached through an extensive comparison of derivatives of the enzyme. Reviewing the evidence, Li e t al. (300) suggested the following pairs: Asp l&Tyr 25, Asp 3&Tyr 92, and Asp 83-Tyr 97. 295. S. Yanari and F. A. Bovey, JBC 235, 2818 (1960). 296. M. Sela, C. B. Anfinsen, and W. F. Harrington, BBA 24, 229 (1957). 297. C. C. Bigelow, C o m p t . Rend. Trav. Lab. Carlsberg 31, 305 (1960). 298. H. A. Scheraga, BBA 23, 196 (1957). 299. H. A. Scheraga, JACS 82, 3847 (1960). 300. L.-K. Li, J. P. Riehm, and H. A. Scheraga, Biochemistry 5, 2043 (1966).

24.

717

BOVINE PANCREATIC RIBONUCLEASE

The X-ray structure does show that Asp 14 and Tyr 25 are hydrogen bonded to each other; Asp 38 is near Tyr 92 but the OH of the latter is H-bonded to the main chain of Lys 37 not the COOH group; Asp 83 is not even very near 97. It is not clear why titration of Asp 83 should affect Tyr 97. On the other hand, Asp 86 is much closer than 83 to Tyr 97, and Asp 111 is as close to Tyr 115, although neither of these potential interactions has been implicated in the abnormal behavior of the tyrosine residues. The procedure of solvent perturbation spectroscopy developed by Laskowski and his colleagues is a sensitive measure of surface topology, and it should in the near future permit detailed correlations with the X-ray structure. A range of perturbants of the spectrum a t 287 nm have been investigated with RNase under a variety of conditions (301). Although rough agreement was obtained with the division into 3 buried and 3 free, a much better fit to the data was obtained by assuming 2 residues fully accessible, 2 partially accessible, and 2 buried. (One should note that the accessibilities in the sense of solvent perturbation and accessibilities in the sense of pK value obtained by titration are not necessarily directly connected.) Reduced carboxymethylated RNase in 8 M urea yields an average accessibility of the tyrosine residues that is about 85% of that of the free amino acids. Ribonuclease, unreduced, in 8 M urea gives an average of about 70% of the free amino acids. The 57%. Thus the variation among the different perturbants is about difference is significant and shows the more compact structure and steric restrictions imposed by the SS bridges in an otherwise random structure. The technique of thermal perturbation spectra introduced by Bello (302) yields 3.6 exposed tyrosines in native RNase at room temperature, an estimate agreeing well with the data quoted above. Perturbation of the spectrum of RNase-A by dioxane was almost exactly one-half that of Ox-RNase or of that expected for 6 free tyrosyl residues (303).This solvent was not included in the series reported by Herskovits and Laskowski (301).The results appear to indicate a 3 : 3 division but could, of course, also fit the 2:2:2 division if the appropriate characteristics are attributed to the free, partially accessible, and buried groups. Ribonuclease-S also appears to have 3 buried tyrosyl residues and S-protein only 2 (304).Presumably Tyr 25 is “normalized” when S-peptide, and thus the Asp 14 interaction, is removed.

+

301. T. T. Herskovits and M. Laskowski, Jr., JBC 243, 2123 (1968) 302. J. Bello, Biochemistry 8, 4542 (1970b). 303. C. C. Bigelow and T. A. Krenitsky, BBA 88, 130 (1964). 304. L. M. Sherwood and J. T. Potts, Jr., JBC 240, 3806 (1965).

718

F. M. RICHARDS AND H. W. WYCKOFF

13. Fluorescence

The fluorescence of ribonuclease solutions has been studied extensively by Cowgill. The absence of tryptophan permits the tyrosine fluorescence to be observed. The tyrosine fluorescence of RNase is very low in comparison with the maximum expected from its tyrosine content. All methods of denaturing RNase lead to an increase in fluorescence. Transitions, as indicated by the pattern of fluorescence change vs. denaturant concentration, are about the same as those indicated by other physical techniques [see, e.g., Gally and Edelman (305)l. Cowgill (306) has shown that phenols tend to combine with organic acid amides in nonpolar solvents presumably by hydrogen bond formation. The association is accompanied by a loss of fluorescence and a red shift of the UV absorption spectrum. Peptides containing tyrosine normally increase their fluorescence on passing from water to organic solvents. Thus, if the buried tyrosine residues of RNase are nonfluorescent, they are likely to be hydrogen bonded to carbonyl oxygens. The X-ray structure indicates that Tyr 25, 92, and 97 are all hydrogen bonded in this manner. Disulfide groups are found to strongly quench the fluorescence of phenols in model compounds (307). Proximity is important because of the inverse sixth power distance dependence. However, there may be an important orientation factor as well. Reduction of the model compounds to the sulfhydryl form resulted in a fluorescence increase, but some quenching from the SH groups also occurred. The thioether sulfur of methionine, on the other hand, had no effect. The current qualitative explanation for the fluorescence of native RNase is: (1) the 3 buried tyrosyl residues show only slight residual fluorescence because of hydrogen bonding and, 2 of the 3 are close to SS bridges 26-84 and 40-95; (2) the “free” tyrosyl residues are fluorescent but they are substantially quenched by SS bridges 58-110 and 65-72. Based on 6 free tyrosines the total fluorescence would be very low as observed. This conclusion is borne out by other evidence. Fluorimetric alkaline titration gives a simple curve with a pK of 10.3 similar to the first part of the usual spectral titration. The phenolate ion is nonfluorescent. By itself the observed effect could represent quenching by nonfluorescent free tyrosines of fluorescent buried ones. This is particularly unlikely in view of the action of dioxane. The fluorescence of simple tyrosine peptides increases linearly with volume per cent of dioxane in the solvent. 305. J. A. Gally and G. M. Edelman, Biopolymers, Symp. 1, 367 (1964). 306. R. W. Cowgill, BBRC 16, 332 (1964). 307. R. W. Cowgill, BBA 140, 37 (1967).

24,

719

BOVINE PANCREATIC RIBONUCLEASE

At 40% dioxane the fluorescence has increased about 50% over the value in water. Ribonuclease-A shows exactly the same effect in what is essentially a fluorimetric solvent perturbation experiment (308). All other physical evidence indicates that no structural change in RNase-A has occurred in this solvent a t neutral pH and room temperature. There appears to be no explanation other than the alteration of fluorescence of accessible tyrosyl residues. With RNase-S 40% dioxane appears to cause the dissociation of Sprotein and S-peptide (308). The fluorescence change as a function of dioxane concentration shows a transition with a midpoint a t about 28% dioxane. At 40% the transition is complete. All tyrosine residues appear to be exposed and all enzymic activity is lost. The quantum efficiencies of various samples in several solvents are shown in Table XVI. Data on quenching by the formation of iodotyrosyl residues (309) or through iodide ion as a solvent component (310) fit the above picture for the structure of the native enzyme. 14. Optical Rotatory Dispersion and Circular Dichroism

The optical rotatory dispersion spectrum of RNase has been measured many times. The details of its shape and the accessible wavelength range vary with the authors and with time as the instruments improve. The general outlines are now well recognized and some typical spectra TABLE XVI AVERAGE QUANTUM EFFICIENCY OF FLUORESCENCE AT 27"" Quantum efficiency (%) Solvent:

Hz0

H20

8M urea

SDSa CDEAc 2mM 2mM

Dioxane

pH :

0.1 7

0.1 1.5

6

0.01 7.5

8

40% 0.1 7

RNase-A RNase-S S-Protein

1.4 1.5 2.0

1.9 [2.5] [2.6]

3.0 [3.3] [3.4]

3.6 13.51 [3.81

4.2 4.3 4.1

1.4 3.3 3.6

z (M):

a

-

Taken from Cowgill (308).

* Sodium dodecyl sulfate.

Cetyldimethylethylammoniumbromide. 308. R. M. Cowgill, BBA 120, 196 (1966). W. Cowgill, BBA 94, 74 (1965). 310. E. A. Brushtein, Biojisika 13, 718 (1968).

309. R.

0.01

0-Mercaptoethanol 0.05 8.5 5.8 [5.91 IS. 21

720

F. M. RICHARDS A N D H . W . WYCKOFF

199 205 0 1 ,

'1

1

,

I

,

a

IQOW

SOOC [""]A

C

-5wc

0 . 2 M PO,,

-----

A (nm)

A (nm)

(0)

(b) 0.2 M PO,,

pH 6.49

pH 6.40

_____

RNoss-A, 2 5 O C RNore-S-protein, 25' C RNare-S-protein, 3OC

RNore-A, 25' C RNore-S, 2 5 O C RNase-S, 3 O C

2000

I 1900

2000

2100

2200

I 2300 2 4 0 0 2500 1900 2000

I

2100

2200

2300

2400

2500

k ( X )

(C)

FIG.11 (a)-(c).

are shown in Fig. 11 (811-313). [ I t should be noted that the far UV curve does not agree well in shape or magnitude with that reported by Tamburro et al. (314) alt,hough these authors and Jirgensons (311) draw essentially the same conclusions about the structural implications.] 311. B. Jirgensons. JACS 89, 5979 (1967).

24.

721

BOVINE PANCREATIC RIBONUCLEASE

I -pH 7.21, Phosphate4i.

.; 3

bornte

',.

.I

<

2 --- pH 8.92, Borate 3 - - - - p H 9 . 6 7 , Borate 4 - -. pH 11.0, KCI 5 ""."" pH 12.7, K C I , KOH

1. :\,

%
*,\

I

2200

2400

I

I

I

2600

I

I

2800

I

3000

I

3200

X ( B ,

(d)

n o . 11. (a) Far ultraviolet rotatory dispersion of ribonuclease. Corrected mean residue specific rotation vs. wavelength [ m ' l ~= [al~M/100 13/(n2 2 ) l where (Y = specific rotation, M mean residue weight, and n = solvent refractive index. Bars give maximal deviation at peaks. Reproduced from Jirgensons (311). (b) Near ultraviolet rotatory dispersion of 0.4870 pancreatic ribonuclease in a 1-mm cell, in (a) 0.15 M phosphate buffer at pH 6.2; (b) 0.15 M glycine-NaOH buffer a t pH 11.5; ( c ) 0.1 N HCl; (d) 1.5% sodium dodecyl sulfate. Reproduced from Glazer and Simmons (31.2). (c) Far ultraviolet circular dichroic spectra of RNase-A, RNase-S, and S-protein at 25" and 3". Reproduced from Pflumm and Beychok ( 8 3 ) . (d) Near ultraviolet circular dichroic spectra of RNase-A as a function of pH. Reproduced from Pflumm and Beychok (313).

+

The rotationally active absorption band of the peptide group produces an ORD peak a t about 199 nm for the helix conformation and a t about 205 nm for the extended fl structure. The negative trough near 230 nm is associated with a weaker transition of the peptide link. The Cotton effects observed in the 27CL290-nm region are a result of the side chain phenolic groups of the tyrosine residues. The conclusions drawn from the curves of the native enzyme in comparison with known structure in the a-helical, @ and random coil forms are that: (1) RNase contains a significant amount of f i structure, not quantified. helix is low. Jirgenson's (2) The content (net) of the right-hand estimate (311) from the 199-nm ORD peak is 13%, remarkably close (Y

312. A. N. Glazer and N. S. Simmons, JACS 87, 3991 (1965). 313. M. N. Pflumm and S. Beychok, JBC 244, 3973 (1969). 314. A. M. Tamburro, A. Scalturin, and L. Moroder, BBA 154, 583 (1968).

722

F. M. RICHARDS AND H.

W.

WYCKOFP

to the earlier figure of 13% from Zimmerman and Schellman (315) and to the X-ray figure of about 15%. Other numbers from different parts of the ORD spectrum vary but all are less than 40%. The difficulties associated with detailed interpretation are the absence of a full range of model compounds and the overlap of the bands. The portions of the structure that are called random or unordered are not a t all random in the native enzyme; they merely are not one of the standard structures [see, e.g.. Zimmerman and Schellman (315)1. I n a more recent study using circular dichroism, Pflumm and Beychok (313) have fitted the observed curve for RNase-A (see Figs. l l c and d ) weighted mixtures of the characteristic bands for helix from p o l y - ~ , structure from poly-L-lysine. The data are comglutamic acid and G patible with 11.5% helix and 33% ,8 conformation. Ribonuclease-S and RNase-A have almost identical C D spectra from 198 to 300 nm. The spectrum of S-protein is markedly different from the other two. The displacement of the ORD trough from the expected 233 nm position to 227 nm has been troublesome for interpretation. I n a recent paper Schellman and Lowe (316) have concluded that this may be expected for short helices or ones that are badly distorted. Both of these criteria apply to the helices in the RNase structure and may account for the position of the curve. The spectrum between 300 and 600 nm generally has been interpreted with the empirical Moffitt equation (317) and the famous coefficient of its second term b,. Some of these values are recorded in Table XVII (318-320). The value of b, has been converted to percent helix on a purely empirical basis with zero equivalent to no helix and about -630 to 100% helix (321) (see Section V,B,3). By this approach also RNase is inferred to have a low helix content. Regardless of the precise interpretation of the ORD spectrum, changes in the spectrum can be used to reflect changes in conformation. These changes are related to secondary structures in the sense that only the immediate environment of a given chromophore influences its contribution to the observed spectrum. The side chain Cotton effect in the 270-290-nm region clearly arises from the tyrosine residues. This effect disappears on denaturation. However, whether the effect is owing to the surface, accessible tyrosines, or 315. 316. 317. 318. 319. 320. 321.

S. B. Zimmerman and J. A. Schellman, JACS 84, 2259 (1962). J. A. Schellman and M. J. Lowe, JACS 90, 1070 (1968). W. Moffitt and J. T. Yang, Proc. Natl. Acad. Sci. U. S. 42, 596 (1956). T. T. Herskovits, JBC 240, 639 (1965). B. Jirgensons, JBC 238, 2716 (1963). P. S. Sarfare and C. C. Bigelow, Can. J. Biochem. 45, 651 (1967). P. Urnes and P. Doty, Advan. Protein Chem. 16, 401 (1961).

24.

VALUESOF bo No.

723

BOVINE PANCREATIC RIBONUCLEASE

FROM

TABLE XVII VISIBLE AND NEARULTRAVIOLET ORD SPECTRA

Sample and conditions

bo

Ref.

- 95 - 125 - 105 -55 0 0 -25 - 115 -90

(318) (319) (3-W

11

Ribonuclease Water, pH 3.4, I = 0.03 M Water, pH 5.6, 0.02M SDS Water, 2.5 M LiBr Water, 5.5 M LiBr 8 M urea, pH 3.5,I = 0.03M 8 M urea, pH 6.6 8 M urea, pH 5.6,0.02M SDS Ethylene glycol, 0.01 M HCl Ethylene glycol, 0.2 M KCl Ethylene glycol, 0.2 M LiCl Chloroethanol

12 13

Ox-RNase Water, Water, 5 M LiBr

1 2 3 4 5 6

7 8 9 10

- 130 - 385 - 50 -50

(SW (318) (319) (319) (318) (318) (318) (318)

(3.w (390)

buried ones is not clear. The first vote for surface (312) was countered (322) and then rebutted (323) with no single explanation of all the effects yet presented. 15. Nuclear Magnetic Resonance and Electron Paramagnetic Resonance

The NMR spectrum of ribonuclease was first reported by Saunders et al. ( 3 2 4 ) . This was also the first NMR spectrum of any protein to be recorded. An interpretation based on the observed spectra of free amino acids was provided immediately by Jardetsky and Jardetsky ( 3 2 5 ) .The major problem in the application of this very powerful technique to the study of proteins is the large number of individual resonances and line broadening resulting from slow rotation even in a small protein and the resultant overlap of the peaks. Most of the absorption is contained in broad unresolved bands even a t the highest resolution presently attainable. The behavior of this area can be examined in a qualitative sense, but individual protons can only be studied in the very high or low field regions outside the main absorption area. Kowalsky indicated the possibility of following reversible conformational changes 322. R. T.Simpson and B. L. Vallee, Biochemistry 5, 2531 (1966). 323. N. S. Simmons and A. N. Glazer, JACS 89, 5040 (1967). 324. M. Saunders, A. Wishnia, and J. G. Kirkwood, JACS 79, 3289 (1957). 325. 0. Jardetsky and C . D. Jardetsky, JACS 79, 5322 (1957).

724

F. M. RICHARDS AND H. W. W Y C K O F F

by NMR (326) and Mandel (327) has given the spectra of RNase and Ox-RNase obtained on a 100-MHz machine, The marked narrowing of the lines and increase in resolution in Ox-RNase is a reflection of the increased motional freedom, on the NMR time scale, in the random coil. The decrease in line width on passing through the thermal transition at 60" is easily measurable, but the width on the high temperature side is still considerably larger than in Ox-RNase. The motional freedom introduced by the thermal transition is thus well short of that of a random coil. Bradbury and Scheraga (328) have studied the imidazole ionization in a series of compounds and made the first study of the individual histidine residues in RNase. The C2 protons are moved well down field just outside the aromatic resonance area. I n a 100-MHz machine Meadows et al. (329) were able to clearly resolve all 4 histidine peaks. By studying the influence of inhibitors and by using several RNase derivatives, 1-CM-His-119-RNase, RNase-S, S-protein, and S-peptide, Meadows et al. (281) were able to assign each of the four peaks to a particular histidine in the sequence. From studies on the pH dependence of the chemical shift, pK values were obtained for each residue (see Section II,C,lO). The behavior of the peak assigned to His 48 as a function of temperature has led Roberts et al. (678) to infer an isomerixation involving this residue, possibly a slow sampling of two states which would explain the line broadening. This may be related to the isomerization seen by French and Hammes (330) in temperature jump kinetic studies. Line broadening for His 119 on lowering the temperature to 10" indicates a similar situation may apply to this residue. The X-ray evidence definitely favors multiple positions for His 119 but provides no direct evidence on His 48 (62). Riiterjans and Witzel (280) have carefully measured the chemical shift of the C2 proton of the His residues as a function of pH a t low ionic strength. The data for His 12 and 119 cannot be fit by ionization curves for simple monobasic acid. The curves are clearly biphasic and indicate a close coupling of two ionizable groups with similar pK values. The authors site this as evidence for a direct interaction between the two imidazole rings but this is not necessary. The phenolic and amino groups of tyrosine show such ionization coupling. These authors also 326. A. Kowalsky, JBC 237, 1807 (1962). 327. M. Mandel, Proc. Natl. Acad. Sci. U. S.52, 736 (1964). 328. J. H. Bradbury and H. A. Scheraga, JACS 88, 4240 (1966). 329. D.H.Meadows, J. L. Markley, J. S. Cohen, and 0. Jardetsky, Proc. Natl. Acad. Sci. U. S. 58, 1307 (1967). 330. T.C. French and G. G. Harnmes, JACS 87, 4669 (1965).

24.

BOVINE PANCREATIC RIBONUCLEASE

725

showed the dramatic influence of ionic strength on the apparent pK values of His 12 and 119 compared with the small effect on His 105 and free imidazole. They noted the strongly positively charged environment of 12 and 119 which is amply shown in the X-ray structure. It is not clear that sole emphasis should be placed on Lys 41 since Lys 7, Lys 66, Arg 10, and Arg 39 are all close-by. Ribonuclease will not normally give a paramagnetic resonance signal. A variety of covalently attached spin labels based on the nitroxide function have been investigated by Smith (331). By choice of reagent and reaction conditions substitution was obtained on various residues. Unfortunately, the reaction mixtures were not fractionated or carefully defined chemically. Electron paramagnetic resonance (EPR) spectra were measured on the reaction mixtures after removal of excess reagent. Although the spectra reflect only those molecules having a spin label, correlation with other types of measurements is difficult. Optical spectra, for example, give an average figure for the total protein present. Nonetheless a number of interesting observations are reported, largely qualitative in nature, concerning the effects, on motion of the spin label, of solvent change, and of specific ligands. The potential of the system fo_r providing sensitive probes of molecular motion in definable parts of the structure is clear. Roberts et al. (332) have synthesized a spin label which apparently goes to the phosphate binding site in a noncovalent interaction. The NMR peaks of His 12 and 119 were broadened on binding this reagent. The other two histidine peaks were not affected.

B. CHAINCONFORMATION AND SOLVENT-INDUCED CONFORMATIONAL CHANGES The conformation of the peptide chain will affect all hydrodynamic spectral and optical rotatory properties and will influence the chemical behavior of specific functional groups. The relationships between these measurable variables and the actual conformations remain elusive, but changes in the parameters may be used to indicate changes in conformation. Unfortunately, the magnitude of the structural changes to be inferred are not always clear. [See the general review of protein denaturation by Tanford (333).] Viscosity and sedimentation are commonly used to estimate large 331. I. C . P.Smith, Biochemistry 7 , 745 (1968). 332. G. C. K. Roberts, J. Hannah, and 0. Jardetsky, Science 165, 504 (1969). 333. C. Tanford, Advan. Protein Chem. 24, 121 (1968).

726

F . M. RICHARDS AND H. W . WYCKOFF

overall changes which would appear in the frictional coefficient or hydrodynamic volume. Membrane escape rates, by dialysis, are probably a more sensitive estimate of small changes in Stokes radius. The spectroscopic methods such as rotatory dispersion, circular dichroism, fluorescence, nuclear magnetic resonance, and ultraviolet absorption all provide detailed, if not always interpretable, information on the structure and environment of specific regions of the macromolecule. The two limiting structural forms are (1) the native enzyme as it exists a t room temperature and pressure a t neutral pH in dilute salt solutions and (2) the chain as a random coil with all SS bond restraints removed and no residual time-independent structure. I n the discussion which follows, the structure of the native enzyme will be assumed to be close to that of the X-ray structure with the caveats mentioned in Section II1,B. Following Bigelow ( S S 4 ) the reference state used for dilute solution studies will be 0.1 M KCl, 25", pH 3-9. The structure does not change detectably in this range of conditions. Tanford et al. (267) have presented convincing evidence, using viscosity data, that most proteins exist as random coils in 6 M guanidine hydrochloride containing 0.1 M mercaptoethanol. They also provided criteria for deciding when this is not true. Titration of RNase in 6 M guanidine HCl with no reducing agent shows an electrostatic factor of essentially zero and all groups with normal p K values except for three tyrosines whose pK values are still 0.4 unit too high ( 3 5 5 ) . I n general there may be a very large number of intermediate states between these two forms, and the set of these which is sampled could easily depend on the conditions chosen to induce the structural change and on the observational method employed. Such structural changes can be brought about by changes in pH, temperature, pressure, ionic strength, organic solvents, urea, guanidinium salts, detergents, and specific ligands. The influences of all of these conditions and reagents can be expected to be interrelated. 1. The Thermal and Acid Transitions

A variety of physical parameters are found to change rather sharply in a narrow range of temperature. The midpoint of such a change, the transition temperature, varies markedly with changes in solvent composition such as pH, ionic strength, and organic solvents. The effect of pH on the thermal transition is shown in Fig. 12 from the work of 334. C. C. Bigelow, J M B 8, 696 (1964). 335. Y. Nozaki and C. Tanford, JACS 89, 742 (1967).

24.

BOVINE PANCREATIC RIBONUCLEASE

727

Brandts and Hunt (336) and of Hermans and Scheraga (337). Figure 12c show the comparison of absorption and rotation estimates of the extent of the transition. In this case two different measurements appear to be influenced to the same extent by the change in structure. The change in fluorescence also appears to follow the absorption change (308). This agreement is not proof that a single step process is involved; that is, a process involving only two states, native and denatured. I n fact, the nonlinear van't Hoff plots that were derived from these data were taken :is evidence of a complex process [see also Hermans and Scheraga (277) 1. Scott and Scheraga (338) found two separable, first-order processes occurring in the kinetics of the absorption change on acidification and took this as confirmation of a t least one intermediate state. Brandts (339), in a careful study of the transition a t equilibrium, pointed out that the curved van't Hoff plots can also be accounted for on the assumption of a large, temperature-dependent change in heat capacity between the native and unfolded forms. He concluded that the process could well be two state in character. He did not, however, explain why two rate processes should be observed. The studies of Van Holde and Baldwin (256) using sedimentation and viscosity also indicate at least two steps since the sedimentation and viscosity changes are not parallel. The measured sedimentation constant of RNase-A in 0.15 M KC1-phthalate buffer pH 2.8 changed from about 1.72 a t 20" to 1.38 a t 60". For the same solution the intrinsic viscosity changed from 3.3 to 5.9 ml/g over the same temperature range. Figure 13 sliows these data plotted in the form of fractional conversion vs. temperature. Klee (340) used proteolysis as a probe of structure a t pH 8 in 0.1 M NaCl and observed transitions in accessibility to a series of hydrolytic enzymes in the range of 50"-60" (see Fig. 14). Different enzymes did not show the same transition temperature. It is interesting that the trypsinindicated transition is just over 50" and that the proteolysis occurs near residue 38 ( 7 7 ) .A spectral transition in DNP-Lys-41-RNase also occurs near this temperature and below the main thermal transition (see Section V,B,5c). X-ray studies have shown the D N P group to be near residue 38. There is thus a collection of studies pointing to relative thermal instability in that part of the molecule and the almost certain existence of scvcral of conformational states in this transition. The transition a t pH 2.2 and 56" does not result in a fully unfolded molecule since residual 336. J. F. Brandts and L. Hunt, JACS 89, 4826 (1967). 337. J. Hermans, Jr. and H. A. Scheraga, JACS 83, 3283 (1961). 338. R. A. Scott and H. A. Scheraga, JACS 85, 3866 (1963). 339. J. F. Bmndts, JACS 87, 2759 (1965). 340. W. A . Klee, Biochemktry 6, 3736 (1967).

Temperature

(OC)

W

P

U

u

I

Temperature (OC)

Temperature

728

(OC)

24.

729

BOVINE PANCREATIC RIBONUCLEASE

T

(OC)

FIG. 13. Trmperaturr depcmdence of tlir fraction of RNase-A in thr denatured form as estimated by (0) sedimentation and ( 0 ) viscosity. Solvent 0.15 M KCl pH 2.8 phthalate buffer. The size of the circles is an approximate estimate of experimental error. Reproduced from Holcomb and Van Holde (&561).

structure can be shown on guanidine hydrochloride denaturation (341). Also, the intrinsic viscosity ($51)is not as high as that seen in 8 M urea, for example. The same transition may also be cxaniined as a function of pH at constant temperature (277). Some typical curves are shown in Fig. 15. These spectrophotometric titration curves are functions also of ionic strength as seen in Fig. 16. The transition temperature is not significantly 341. K. C. Aune, A. Salahuddin, ( 1967).

M. H. Zarlengo, and C. Tanford, JBC 242, 4486

FIQ. 12. (a) The thermal transition of RNase-A (0.3-0.4 mg/ml) a t various pH values. The solvent was either dilute HC1 or 0.04 M glycine buffer for the two most alkaline curves. The change in molar absorbancy a t 287 nm is shown as a function of temperature. From Brandts and Hunt (336). (b) The thermal transition as monitored by the change in optical rotation a t 436 nm. The protein concentration was 1.9 m g / d in 0.16 M KCl. From Hermans and Scheraga (557).( c ) Comparison of the data from (b) and from a set comparable to (a) where both the absorbance and rotation values have been normalized to fractional conversion to the denatured form. From Hermans and Scheraga (337).

730

F. M . RICHARDS AND H. W. WYCKOFF

I

I

30

40

4

1

50

60

Temperature

FIG. 14. The rate of hydrolysis of RNase relative to that of Ox-RNase as a aminopeptidase, ( A ) trypsin, function of temperature. The proteases used were (0) (0) chymotrypsin, and ( 0 )carboxypeptidase. Reproduced from Klee (340).

affected by ionic strength but the slope of the transition, and thus its cooperativity, certainly is. Most studies have concentrated on those conditions where reversible transitions can be demonstrated. However, a t neutral pH the thermal transition temperature is high enough to introduce difficulties. Ribonuclease kept a t 95” a t pH 7 for 20 min is irreversibly denatured both in its spectral properties and enzymic activity (337). Tramer and Shugar showed that RNase inactivated a t pH 7.8 by heating for 30 min has LLnormalized”all of its tyrosine residues as far as alkaline spectrophotometric titration is concerned. However, the magnitude of the acid difference spectrum is unaffected although the midpoint has shifted from pH 2 to 3. Further complexities in these transitions were found by the work of

0 c N m

O

a

PH

FIG. 15. Spectrophotometric titration curves for RNase, 1.9 mg/ml, in 0.08 M KC1. The temperature for each titration is given in the figure. Reproduced from Hermans and Scheraga (577).

24.

731

BOVINE PANCREATIC RIBONUCLEASE

1000

e 500

0

'

250 0

I

I

I

I

I

2

3

4

PH

FIG.16. Spectrophotometric titration curves for RNase at 25O as a function of ionic strength provided by KCl; a, 0.10 M ; b, 0.17 M ; c, 0.35 M ; and d, 1.10 M. Inset: Values of - A ~ Z U ~ ~ ,observed ,, at pH 1 at tt function of ionic strength. Reproduced from Bigelow and Krenitsky (303).

Vithayathil et al. (3.49).In 0.5 M HC1 a t 30" RNase-A undergoes structural alterations which can be detected chromatographically a t neutral pH. However, all the products are equally active enzymically, and no reaction would have been detected by assay. At pH 11.0 even more involved structural changes take place quite rapidly. Irreversible alkaline denaturation takes place a t higher pH and is very rapid a t 13. Here the activity loss is accompanied by marked spectral changes indicating reactions such as J3 elimination a t cystine or serine residues (9.49).A temperature-induced isomerization a t neutral p H has been reported by French and Hammes. This is discussed in a later section on nucleotide binding. 2. Urea Transition

High concentrations of urea produce an increase in viscosity and changes in absorption and rotation spectra. The transition as a function of urea concentration is steep, implying a cooperative phenomenon involv342. P. J. Vithayathil, A. S. Achaxya, and B. N. Manjula, Conform. Bwpolymers, Papers Intern. Sump., Univ. Madras, 1967 Vol. 1, p. 291. Academic Press, New York, 1967. 343. Z. Bohak, JBC 239, 2878 (1964).

732

F. M. RICHARDS AND H . W. WYCKOFF

ESTIM.4TES

OF

TABLE XVIII THERMAL TRANSITION TEMPI~~R.ITUR&S IN

v \KIOUS

SOLVliNTS

~ _ _ _ _ _ _ _ _

T, No. 1 2 2a 3 3a 3b 4 5 6 7 8 9 10 11 12 13 14 15 16 17

Method

Solvent

(")

- -

H20, Z 0, pH 4.2 DZO, Z 0, p H 4.5 DZO, I 0, pH 4 3 Acetate buffer, Z = 0.065, pH 5.0 3 0 4 0 0.2 AI NaCl 0.2 M phosphate bnfyer p1-I 6 . 0 38 61 5 0.12 A l KCl, 0.01 A1 phosphate pH 6.,5-6.8, 25' 57 4 10% methanol 55 4 1Oyoethanol 54 4 1O'jo isopropanol 50 4 10% propanol 51 4 10')o isobiitanol HzO, pI1 2.3, 110 bLifIe1 35 40 10 4070 glycol 42 10 25@Ioglycerol 42 10 255Zo erythritol 49 10 40y0 sorbitol 10 3 11l 2,3-britanediol 37 10 3 2cl 1,4-butanediol 34 64 0.1 A l K phosphate 5.5 1.1 mold purine 39 66 66 1 66

N N

Ref.

Tritium exchange ( i i i 24 hr) NMR variation of 7', AEXI n m

RNA-activity assay

NhlIt-C2 of llis 12 a i d 111) C hroma togra phy AEW o m

+ + + + + + + + + + + +

ing a large number of urea molecules. The transition concentration is a function of both pH and temperature. At least at neutral pH and low temperature the urea-induced structural change is reversible. Nelson and Hummel (351) made a careful investigation of the urea transition near pH 7. Their absorbance difference spectra clearly show changes in the environment of phenylalanine as well as tyrosine residues. Thc optical rotation changes occur in the same range of urea concentrations. The loss of enzymic activity ( U > p ) is biphasic. At low urea con344. D. J. Blcars and s. s. Danyluk, BBA 147, 404 (1967). 345. J. Hermans, J r . and H. A. Schcraga, BBA 36, 534 (1959). 346. G. Kalnitsky and H. Resnick, JBC 234, 1714 (1959). 347. G. C. I<. Roberts, D. H. Meadows, and 0. Jardetsky, Biochemistry 7, 2053 (1968). 348. E. E. Schrier and H. A . Schcraga, BBA 64, 406 (1962). 349. S. Y. Gerisma, JBC 243, 959 (1968). 350. E. 0. Akinrimisi and P. 0. P. Ts'o, Biochemistry 3, 619 (1964). 351. C. A. Nclson and J. P. Hummel, JBC 237, 1567 (1962).

24.

BOVINE PANCREATIC RIBONUCLEASE

733

ccntrations there is a hyperbolic loss probably representing competitive inhibition followed by a steeper fall in the transition region, approaching zero at high urea concentration [see also Barnard (352)l. The kinetics of both the denaturation and renaturation were first order, but the former was markedly pH and temperature dependent while the latter was not. The rate of urea denaturation was inhibited by a variety of anions known to bind to the enzyme: in decreasing order of effectiveness, pyrophosphate, 2'-C1\/IP1phosphate, citrate, tartrate, and sulfate ( 3 5 3 ) .This inhibition was greater a t pH 5.6 than 7.3. The binding constants were the same as those estimated by inhibition of the enzymic reaction. As with the pH and temperature effects, the anions had no demonstrable effect on the rate of renaturation. In a detailed study of thc cquilibria involved in the urea transition, Tanford (354) showed that a two-state process could not explain the RNase data and that the cooperative units, whose unfolding was reflected in the measurements, must be much less than the total molecule. Each unit was probably not more than one-third of the total molecule. This conclusion was based on Tanford's theory and the data of Nelson and Hummel (351) and of Foss and Schellman (355). The midpoint of the transition at room temperature in 0.1 M KCl and neutral pH is about 6 M urea. Barnard (356) found a midpoint a t 6 . 7 M urea at pH 7 and 2Ti"C.By fluorescence the midpoint was about 6.5 M urea (308).Between 12 and 16 molecules of urea per molecule of protein appeared to be involved in the transition (356), 12 being the kinetic order of the unfolding reaction and 16 being the value derived from the slope of the equilibrium curve. Again evidence for multiple states was presented. 3. Organic Solvents

a. Ethglene Glycol. Sage and Singer (294) were able to dissolve RNase in a variety of anhydrous organic solvents. On return to water almost complete enzymic activity was recovered from solution in ethylene glycol, fonnamide, dimethylformamide, and about 50% activity from dimethylsulfoxidc. I n pure ethylene glycol 0 . 2 M in KCl a t 25" the value of b, was -92". Spectroscopic alkaline titration in this solvent showed that all 6 tyrosinc residues appeared to titrate with a single pK value. 352. E. A. Barnard, J M B 10, 263 (1964). 353. C. A. Nelson, J. P. Hummel, C. A. Swenson, and L. Friedman, JBC 237, 1575 (1962). 354. C. Tanford, JACS 86, 2050 (1964). 355. J. G . Foss and J. A. Schellman, J . Phys. Chem. 63, 2007 (1959). 356. E. A . Barnard, J M B 10, 235 (1964).

734

F. M. RICHARDS AND H.

W.

WYCKOFF

I n a more recent study, Bello (357) has pointed out that this does not necessarily indicate denaturation in this solvent, and, in fact, that high degrees of order in both glycol and glycerol are maintained a t least up to 90% polyol. The transition temperature is lowered so that in pure glycol it is near 25". Under these conditions the protein can be expected to be very sensitive to other parameters such as the concentration of another solvent component. The acid transition (303) is affected slightly or not a t all by aqueous ethylene glycol concentrations up to 4M, in marked contrast to dioxane, for example.

b. 2-Chloroethanol. Weber and Tanford (358) have shown that RNase is soluble in the entire range of 2-chloroethanol: water mixtures a t pH 2. ionic strength 0.03. The intrinsic viscosity rose from 3.3 to 8.0 ml/g at 80 mole-% water, and then decreased to 6.5 at 0% H,O. The Noffitt constant b,, became more negative through the entire range with a final value of -380 in pure 2-chloroethanol. The protein appears to be monomeric at all stages. The results are interpreted as a disruption of the native structure followed by a reordering into a new helical structure to the extent permitted by the SS bonds. c. Diozane. The midpoint of the acid transition is shifted to higher pH values with increasing dioxane concentration. The change is about 1 pH unit at 4 M dioxane. The solvent perturbation effect of dioxane at neutral pH is that to be expected for 3 accessible tyrosine residues (see Section V,A,12). At neutral pH and dioxane concentrations up to 4 M there does not seem to be any denaturing effect. However, there must be an incipient loosening of the structure since in 40% dioxane S-protein and S-peptide are dissociated (Section V,A,13).

d. Phenol. No change in specific rotation a t the D line was observed in 80% phenol. The intrinsic viscosity increased to 11.2 but returned to the normal value on transferring the enzyme back to water. A summary of some other observations on the thermal transition is given in Table SVIII. The blierific effects of alkyl c1i:iin length 011 a variety of denaturants are Jiown iii Table SIX.These results are compatible with stabilization of the denatured RNaw hy the liydrophobic parts of the denaturants (359). It is clear (360) that the aliphatic 357. J. Bello, Biochemistry 8, 4535 (1970a). 358. R. E. Weber and C. Tanford, JACS 81, 3255 (1959). 359. E. E. Schrier, R. T. Ingwall, and H. A. Scheraga, J . Phys. Chem. 69, 298 (1965). 360. E. E. Schrier and L. D. Mackey, J. Phys. Chem. 72, 733 (1968).

24.

735

BOVINE PANCREATIC RIBONUCLEASE

TABLE XIX EFFECTON TllANSITION TEMPERATURES OF ALKYL CHAIN LENGTH OF VARIOUS DENATURANTSO+ ATm

Alkyl chain

ROHb

RNH3Clb

R3NBr/4b

RCOONab

CH3 CHaCHz CIIaCHzCHz CII3CHzCHzCHz

-1.6 -3.1 -7.1 -13.1

0

-1.1 -3.5 -6.5

0 -1.2 -3.1 -6.5

+O.G -1.3 -3.2

-

Data from Schrier and Mackey (360) and von Hippel and Wong (361). concentration in 0.12 M KC1. 0.01 M phosphate buffer pH 6.7 at 25". c The contribution of the polar group in any series has been consideredadditive and has been subtracted to permit direct comparison of alkyl group effects.

* All compounds tested at 1 M

carbon next to a group with a formal charge is less effective in this respect than those farther along the chain (361). 4. Effects of Added Electrolytes

A summary of the transition temperatures of RNase in the presence of a variety of electrolytes is given in Table XX (361, 362). The effects of the usual salts follow the Hofmeister series. The remarkable difference in denaturing power of the various guanidinium salts is quite general. The sulfate actually tends to stabilize slightly while the thiocyanate is a more powerful denaturant than the chloride. Irreversible denaturation is markedly inhibited by spermine (363).There is no clear correlation between stabilizing effects and the complex inhibition curves obtained by Wold (see Section VI,E,2).

a. Sodium Dodecyl Sulfate. At sodium dodecyl sulfate (SDS) concentrations below 4 m M a t pH 6.1 RNase is precipitated. It is soluble a t n m (-740) higher concentrations (364). The maximum change in occurs a t 8 mM. No further change occurs up to 0.1 M . This is assumed to correspond to the normalization of tyrosine residue B, 2 tyrosine residues being still buried. Stark et al. (158) found that 0 . 2 M SDS did not make the methionine residues accessible to alkylation although the unique 361. 362. 363. 364.

P. H. von Hippel and K. Y. Wong, JBC 240, 3909 (1965). A. Ginsburg and W. R . Carroll, Biochemistry 4, 2159 (1965). J. Goldstein, BBA 181, 345 (1969). C. C. Bigelow and M. Sonenberg, Biochemistry 1, 197 (1962).

736

F. M. RICHARDS A N D H . W. WYCKOFF

TABLE XX EFFECT OF ELECTROLYTES O N THE TRANSITION TEMPERATURE Salt

Transition temp.

Salt

+

0.013 M cacodylate buffer pH 7.0 Solvent 0.15 M KCl the indicated electrolyte a t a concentration of 1 Ma Water 61.5 K phosphate (pH 6.6) Urea 58.5 (NHa) 80, (Guan)sSOQ 61.5 KC1, NaCl Guan acetate 57 LiCl Guan chloride 52 NaBr Guan 8CN 28 LiBr CaCls KSCN

Chloride Phosphate Sulfate a

b

Transition temp.

+ 78 73 62.5 61.5 59 57 51.5 47

Solvent pH 2.1, ionic strength 0.018 ill (cation not specified but presumably K ) c 29.9 31.9 42.7

From von Hippel and Wong ( 3 3 1 ) Guan stands for guanidinium ion. From Ginsburg and Carroll (361).

histidine reaction was destroyed. The side chain Cotton effect near 280 nm in the normal ORD spectrum of RNase is completely removed by 0.05 M SDS (312). Ribonuclease-A is inactive (C>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

BOVINE PANCREATIC RIBONUCLEASE

(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

F. M. RICHARDS AND H.

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.

BOVINE PANCREATIC RIBONUCLEASE

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.

BOVINE PANCREATIC RIBONUCLEASE

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

BOVINE PANCREATIC RIBONUCLEASE

-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

F. M. RICHARDS AND H. W. WYCKOFF

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

F. M. RICHARDS AND H. W. WYCKOFF

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.

BOVINE PANCREATIC RIBONUCLEASE

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

F. M. RICHARDS AND H. W. WYCKOFF

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.

BOVINE PANCREATIC RIBONUCLEASE

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

F. M. RICHARDS AND H. W. WYCKOFF

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

BOVINE PANCREATIC RIBONUCLEASE

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

F. M. RICHARDS AND H . W. WYCKOFF

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

BOVINE PANCREATIC RIBONUCLEASE

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

F. M. RICHARDS AND H. W. WYCKOFF

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.

BOVINE PANCREATIC RIBONUCLEASE

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

F. M. RICHARDS AND H. W. WYCKOFF

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.

BOVINE PANCREATIC RIBONUCLEASE

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

F. M. RICHARDS AND H. W. WYCKOFF

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.

BOVINE PANCREATIC RIBONUCLEASE

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

This Page Intentionally Left Blank

24.

BOVINE PANCREATIC RIBONUCLEASE

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 <7.0 6.4 6.2

-20 -25 0 0 0

6.6 6.2

+10

$10

Aromatic shift

C(5)-H

C(6)-H

C(1’)-H

-

-

Yes

-22 2-20 - 17 - 18 - 18

-

-6 -4 -5 -5 -5

+30 Yesc +30 Yes

-3

- 12 -3 -3 -3

No No

a Downfield chemical shift changes are given as negative numbers and are in cycles per second. I n all cases except cytidine and 5’-methyl-2’-deoxycytidine, inhibitor concentrations were sufficient to saturate the enzyme between pH 5 and 7. [Reproduced from Meadows et al. (466).] * Here A8 stands for change in chemical shift of the (C2)-H peak of the fully protonated histidine. Because of broadening of the peak, this shift could not always be followed over the whole concentration range.

EiU

24.

BOVINE PANCREATIC RIBONUCLEASE

765

seen in His 12 and 119 on ligand association. No effect on His 105 is observed while the effects on His 48 are small, complicated, and apparently resulting from a conformational change in the protein. From the NMR data the cytidine ring appears to bind in the same way in all the nucleotides studied. The phosphate group binds in the same way with 3’-CMP and 2’-CMP but not 5’-CMP. The C2 proton of His 119 interacts directly with the phosphate while that of His 12 does not, and both residues appear to be protonated. The X-ray data are in general agreement with these conclusions. If the P atom in 3’-CMP and 2’-CMP is to be in the same place with respect to the protein, some difference in rotation about the glycosidic bond is required between the 2‘ and 3‘ isomers. This effect is seen in both the NMR and X-ray results. The NMR data are compatible with the assumption of strong binding of the phosphate dianion form of 3’-CMP with the fall in binding a t pH values below 5.6 attributed to protonation of the phosphate group while the fall on the alkaline side results from deprotonation of the histidine residues. Again the best fit is obtained if it is assumed that strong binding of 3’-CMP requires both His 12 and 119 to be protonated in the complex. The environment of His 48 is affected by the 3’-CMP binding, but this residue is not in contact with the nucleotide. On examination of difference spectra, Anderson et al. (455) concluded that protonation of the cytosine ring in the pH range 5.5-4.0 did not affect the nucleotide binding (Table XXV). These same data cannot be used to establish definitely the ionic form of the phosphate group of the various ligands when complexed to RNase. The binding study does suggest that 2’-CMP and 3’-CMP bind slightly differently, that the free enzyme has two groups with pK values of 5 and 6.5 involved in 3’-CMP binding, and that three groups with pK values of 5, 5, and 6.5 are involved in 2’-CMP binding. Note again that these values are expected to be markedly ionic strength dependent. Hammes (466) has summarized some of the extensive studies from his laboratory on the interaction of a variety of nucleotides with RNaseA as seen by relaxation kinetic measurements. The bimolecular and isomerization steps that occur with each of the nucleotides are very much faster than the rate determining steps separating the different substances. Thus the kinetic parameters for the interaction of each nucleotide can be established separately and then combined with steady state kinetic data to provide a detailed kinetic picture. The bimolecular steps are recognized by the concentration dependence of the relaxation time and the isomerization steps by the lack of a concentration dependence . 466. G. G . Hammes, Advan. Protein Chem. 23, 1 (1968).

766

F. M. RICHARDS AND H . W. WYCKOFF

The enzyme itself undergoes a pH-dependent isomerization summarized as follows (467): ki

EH

E’H

E+

E‘+H 71

At 25”, ionic strength 0.2 M , k, = 780 sec-l, k-, = 2470 sec-l, pK, = 6.1, pK,’ > 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.

BOVINE PANCREATIC RIBONUCLEASE

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

BOVINE PANCREATIC RIBONUCLEASE

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

F. M. RICHARDS AND H. W. WYCKOFF

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

F. M. RICHARDS AND H.

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

BOVINE PANCREATIC RIBONUCLEASE

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

F. M. RICHARDS AND H. W. WYCKOFF

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

F. M. RICHARDS AND H. W. WYCKOFF

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.

BOVINE PANCREATIC RIBONUCLEASE

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

F. M. RICHARDS AND H. W. WYCKOFF

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

BOVINE PANCREATIC RIBONUCLEASE

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

F. M. RICHARDS AND H. W. WYCKOFF

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

BOVINE PANCREATIC RIBONUCLEASE

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

F. M. RICHARDS AND H.

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

BOVINE PANCREATIC RIBONUCLEASE

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

F. M. RICHARDS AND H. W. WYCKOFF

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.

BOVINE PANCREATIC RIBONUCLEASE

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

F. M. RICHARDS AND H. W. WYCKOFF

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.

BOVINE PANCREATIC RIBONUCLEASE

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

F. M. RICHARDS AND H. W. WYCKOFF

(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.

BOVINE PANCREATIC RIBONUCLEASE

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

F. M. RICHARDS AND H.

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.

BOVINE PANCREATIC RIBONUCLEASE

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

F. M. RICHARDS AND H.

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.

BOVINE PANCREATIC RIBONUCLEASE

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.

BOVINE PANCREATIC RIBONUCLEASE

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

F. M. RICHARDS AND H. W. WYCKOFF

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

BOVINE PANCREATIC RIBONUCLEASE

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.

BOVINE PANCREATIC RIBONUCLEASE

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

F. M. RICHARDS AND H. W. WYCKOFF

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.

BOVINE PANCREATIC RIBONUCLEASE

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

F. M. RICHARDS AND H. W. WYCKOFF

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