Ribonucleases Structures and Functions
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Ribonucleases Structures and Functions Edited by
Giuseppe D'Alessio Universit~ Degli Studi di Napoli Federico II Naples, Italy
James F. Riordan Harvard Medical School Boston, MA
A C A D E M I C PRESS San Diego
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Front cover photograph: Illustrates the secondary structure and residues important for catalysis and recognition of bovine pancreatic ribonuclease. C o u r t e s y of Dr. Gary L. Gilliland, University of Maryland, Center for A d v a n c e d Research in B i o t e c h n o l o g y , B i o t e c h n o l o g y Institute and National Institutes o f Standards and T e c h n o l o g y , Rockville, Maryland.
This book is printed on acid-free paper. @
Copyright 9 1997 by ACADEMIC PRESS All Rights Reserved. No part of this publication may be reproduced or transmitted in tiny h)rm or by any means, electronic or mechanical, including photocopy, recording, or any information storage and retrieval system, without permission in writing from the publisher. A c a d e m i c P r e s s , Inc. 15 East 26th Street, 15th Floor, New York, New York I()()!(), USA h t t p ://w w w. apn e t .corn Academic Press Limited 24-28 Oval Road, London NW! 7DX, UK http://www.hbuk.co.uk/ap/ Library of Congress Cataloging-in-Publication Data Ribonucleases : structures and functions / edited by Giuseppe D'Alessio, James F. Riordan p. cm. Includes index. ISBN 0-12-588945-3 (alk. paper) I. Ribonucleases--Structure-activity relationships. I. D'Alessio, Giuseppe. II. Riordan, James F., date. QP6()9.P53R53 1996 96-41839 574.87'3283--DC20 CIP
PRINTED IN THE UNITED STATES OF AMERICA 96 97 98 99 00 01 EB 9 8 7 6
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I
Contents
Contributors ............................................................................... Preface
1
......................................................................................
xi xv
Escherichia coli Ribonucleases: Paradigms for Understanding Cellular R N A Metabolism and Regulation
Allen W. Nicholson I. II. III. IV. V. VI. VII. VIII.
Introduction
.........................................................................
G a l l e r y of Escherichia Coli R i b o n u c l e a s e s . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Endoribonucleases .................................................................. Exoribonucleases
...................................................................
19
Ribonuclease Functional Roles ....................................................
23
Other Ribonucleases ............................................................... Ribonuclease Regulation and RNA Metabolic Control .......................
33 35
Conclusions and Prospects
37
........................................................
References ...........................................................................
2
2 3 3
38
Barnase and Barstar
Robert W. Hartley I. II. III. IV. V. VI. VII.
.........................................................................
51
Gene Structure ......................................................................
Introduction
54
Activity and Structure ............................................................. Barnase, Barstar, and the Folding Problem .....................................
57 70
Barnase and Barstar Homologs ...................................................
86
O t h e r P r a c t i c a l A p p l i c a t i o n s of B a r n a s e Concluding Remarks
........................................
...............................................................
References ...........................................................................
89 90 91
vi
Contents
RNase T1/RNase T2 Family RNases
3
Masachika Irie I. II.
R N a s e T1 F a m i l y RNases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . R N a s e T2 F a m i l y RNases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
101 109 124
Structure and Mechanism of Action of Cytotoxic Ribonuclease a-Sarcin Ira G. Wool I. II. III. IV. V. VI. VII. VIII. IX. X. XI. XII.
5
Discovery of c~-Sarcin and D e t e r m i n a t i o n of Basis of Its Cytotoxicity . . . . . . Substrate Specificity of ot-Sarcin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . a - S a r c i n Cleavage Site S e q u e n c e . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . S t r u c t u r e of ot-Sarcin and of R e l a t e d Aspergillus Toxins . . . . . . . . . . . . . . . . . . . . . . M e c h a n i s m by Which c~-Sarcin E n t e r s Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . ot-Sarcin R e c o g n i t i o n E l e m e n t s in 28S r R N A . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . C o n f o r m a t i o n of Sarcin D o m a i n in 28S r R N A . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Effect of M u t a t i o n s of A n a l o g of G4319 in 28S r R N A on R e c o g n i t i o n of O l i g o r i b o n u c l e o t i d e s by c~-Sarcin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . P h e n o t y p e of M u t a t i o n s in G2655 in Escherichia coli 23S r R N A . . . . . . . . . . . . R e l a t i o n s h i p of ot-Sarcin R N A Identity E l e m e n t to Selection by T o x i n of U n i q u e Site of P h o s p h o d i e s t e r B o n d Hydrolysis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . T h r e e - D i m e n s i o n a l Structure of Rcstrictocin, a H o m o l o g of c~-Sarcin. Binding to c~-Sarcin D o m a i n R N A and Catalytic M e c h a n i s m . . . . . . . . . . . . . . . . Coda . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
131 132 133 136 138 138 145 148 149 152 153 159 159
Plant Ribonucleases
Pauline A. Bariola and Pamela J. Green I. II. III. IV.
6
Introduction ......................................................................... Classes of Plant RNases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . R e g u l a t i o n and F u n c t i o n s of Plant R N a s e s . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Conclusions and F u t u r e Prospects . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
163 164 173 182 183
S-RNases and Other Plant Extracellular Ribonucleases
Simon K. Parry, Ying-hong Liu, Adrienne E. Clarke, and Ed Newbigin I. II. III. IV.
Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Self-Incompatibility in the S o l a n a c e a e . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . S-RNases f r o m the Rosaceae . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . E x t r a c e l l u l a r R i b o n u c l e a s e s A s s o c i a t e d with Stress . . . . . . . . . . . . . . . . . . . . . . . . . . . .
192 192 202 203
Contents V. VI.
7
Seed R N a s e s a n d O t h e r R N a s e s of U n k n o w n F u n c t i o n a n d L o c a t i o n . . . . . . P l a n t E x t r a c e l l u l a r R N a s e s : E n z y m e s in S e a r c h of a S u b s t r a t e . . . . . . . . . . . . . . References ...........................................................................
vii 205 206 208
Evolutionary Reconstructions in the Ribonuclease Family
Steven A. Benner, Mauro L Ciglic, Monika Haugg, Thomas M. Jermann, Jochen G. Opitz, Sun-Ai Raillard-Yoon, Josef Soucek, Joseph Stackhouse, Nathalie Trabesinger-Riif, Katrin Trautwein, and Todd R. Zankel I. II. III. IV. V. VI.
8
Introduction ......................................................................... P r o t e i n E n g i n e e r i n g to U n d e r s t a n d E v o l u t i o n . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . R e c o n s t r u c t i n g E v o l u t i o n of B i o m o l e c u l a r B e h a v i o r in R N a s e Superfamily .......................................................................... R e p a i r of D a m a g e d P s e u d o g e n e s by G e n e C o n v e r s i o n : A M e c h a n i s m for O b t a i n i n g B i o m o l e c u l a r F u n c t i o n in P r o t e i n s . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . P h y s i o l o g i c a l F u n c t i o n of S e m i n a l R N a s e . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Conclusions .......................................................................... References ...........................................................................
214 215 225 232 237 238 239
Evolution of Vertebrate Ribonucleases:
Ribonuclease A Superfamily Jaap J. Beintema, Heleen J. Breukelman, Antonella Carsana, and Adriana Furia I. II.
The Ribonuclease Superfamily ................................................... M a m m a l i a n R i b o n u c l e a s e s 1: P a n c r e a t i c T y p e . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References ...........................................................................
245 252 265
Pancreatic Ribonucleases
9
Claudi M. Cuchillo, Maria Vilanova, and M. VictOria Nogu~s I. II. III. IV. V. VI.
10
Introduction ......................................................................... R e a c t i o n C a t a l y z e d by P a n c r e a t i c R i b o n u c l e a s e s . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Specificity of R e a c t i o n a n d Catalytic M e c h a n i s m . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . S u b s t r a t e B i n d i n g Subsites: S t r u c t u r e a n d F u n c t i o n . . . . . . . . . . . . . . . . . . . . . . . . . . . T h e C a r b o h y d r a t e Moiety: S t r u c t u r e a n d F u n c t i o n . . . . . . . . . . . . . . . . . . . . . . . . . . . . F o l d i n g / U n f o l d i n g of R i b o n u c l e a s e A . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References ...........................................................................
272 272 276 284 287 290 297
Crystallographic Studies of Ribonuclease Complexes
Gary L. Gilliland I. II. III.
Introduction ......................................................................... Phosphate/Sulfate-Free RNase .................................................... P h o s p h a t e / S u l f a t e B i n d i n g Site . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
306 308 310
viii IV. V. VI. VII. VIII. IX. X.
11
Contents Substrate Analog-RNase Complexes ............................................ Transition State A n a l o g - R N a s e Complex . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Product-RNase Complexes ....................................................... Nonproductive Ligand-RNase Complexes ...................................... Other Ligand-RNase Complexes ................................................ Semisynthetic RNases .............................................................. Conclusions .......................................................................... References ...........................................................................
311 317 320 323 324 329 333 337
N M R Solution Structures of Ribonuclease A and Its
Complexes with Mono- and Dinucleotides Carlos Gonz6lez, Jorge Santoro, and Manuel Rico I. II. III. IV.
12
Introduction ......................................................................... S t r u c t u r e of B o v i n e P a n c r e a t i c R i b o n u c l e a s e A in A q u e o u s S o l u t i o n . . . . . . L i g a n d B i n d i n g Studies: C o m p l e x e s with M o n o - a n d D i n u c l e o t i d e s . . . . . . . . . S t r u c t u r e s of O t h e r A n i m a l R i b o n u c l e a s e s in S o l u t i o n . . . . . . . . . . . . . . . . . . . . . . . . References ...........................................................................
343 345 358 373 378
Seminal Ribonuclease" The Importance of Diversity
Giuseppe D'Alessio, Alberto Di Donato, Lelio Mazzarella, and Renata Piccoli I. II. Ill. IV.
13
Introduction ......................................................................... Isolation and P r o d u c t i o n of S e m i n a l R N a s e . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Structures . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Functions ............................................................................ References ...........................................................................
383 385 388 408 417
Eosinophil-Associated Ribonucleases
Melissa R. Snyder and Gerald J. Gleich I. II. III. IV. V.
14
Introduction ......................................................................... The Ribonuclease Superfamily ................................................... Eosinophil-Derived Neurotoxin .................................................. Eosinophil Cationic Protein . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Conclusion ........................................................................... References ...........................................................................
426 429 431 436 439 440
Structure and Function of Angiogenin
James F. Riordan I. II. III. IV.
Introduction ......................................................................... Angiogenesis ........................................................................ Isolation of A n g i o g e n i n . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . C h a r a c t e r i z a t i o n as M e m b e r of the R i b o n u c l e a s e F a m i l y . . . . . . . . . . . . . . . . . . . . .
446 447 450 453
~Contents V. VI. VII. VIII. IX. X. XI.
15
R e l a t i o n s h i p of R N a s e A c t i v i t y a n d A n g i o g e n i c A c t i v i t y . . . . . . . . . . . . . . . . . . . . . I n t e r a c t i o n w i t h E n d o t h e l i a l Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A n g i o g e n i n E n h a n c e m e n t of A c t i n A c c e l e r a t i o n of P l a s m i n o g e n Activation ........................................................................... A n g i o g e n i n S u p p o r t of E n d o t h e l i a l Cell A d h e s i o n . . . . . . . . . . . . . . . . . . . . . . . . . . . . M e c h a n i s m of A c t i o n . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B i o l o g y of A n g i o g e n i n . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Epilogue ............................................................................. References ...........................................................................
ix 467 470 476 477 478 480 483 483
Antitumor RNases
Richard J. Youle and Giuseppe D'Alessio I. II. III. IV.
16
Introduction ......................................................................... Bovine Seminal RNase ............................................................ Amphibian RNases ................................................................. Concluding Remarks ............................................................... References ...........................................................................
491 492 499 508 509
2-5A-Dependent RNase L: A Regulated Endoribonuclease in the Interferon System
Robert H. Silverman I. II. III. IV. V.
17
P e r s p e c t i v e s on the 2 - 5 A S y s t e m . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . S t r u c t u r e a n d F u n c t i o n of R N a s e L . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B i o c h e m i c a l P r o p e r t i e s of R N a s e L . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D i s t r i b u t i o n , L o c a l i z a t i o n , a n d R e g u l a t i o n of R N a s e L a n d Its G e n e . . . . . . . Biological A c t i v i t i e s of R N a s e L . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References ...........................................................................
516 518 524 532 536 541
RNA-Processing RNases in Mammalian Cells
Jeff Ross I. II. III. IV.
18
Introduction ......................................................................... rRNA-Processing Ribonucleases ................................................. mRNA-Degrading Ribonucleases ................................................ Summary ............................................................................. References ...........................................................................
553 554 563 576 577
Messenger RNA Ribonucleases and mRNA Turnover in
Saccharomyces cerevisiae Christine E. Brown and Alan B. Sachs I. II.
Introduction ......................................................................... P a t h w a y s of m R N A D e c a y . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
589 590
Contents III. IV.
19
mRNA Ribonucleases ............................................................. Discussion and Future Directions ................................................ References ........................................................................... Ribonuclease
596 611 613
Inhibitor
Jan Hofsteenge I. II. III. IV.
Index
Introduction ......................................................................... Biological Properties ............................................................... Molecular Properties ............................................................... Applications ......................................................................... References ...........................................................................
621 622 632 646 649
.........................................................................................
659
Contributors Numbers in parentheses indicate the pages on which the authors' contributions begin.
Pauline A. Bariola (163), Department of Energy Plant Research Laboratory and Department of Biochemistry, Michigan State University, East Lansing, Michigan 48824. Jaap J. Beintema (245), Department of Biochemistry, University of Groningen, 9747 AG Groningen, The Netherlands. Steven A. Benner (213), Department of Chemistry, Laboratory for Organic Chemistry, E. T. H. Zentrum, CH-8092 Zurich, Switzerland, and Department of Chemistry, University of Florida, Gainesville, Florida 32611. Heleen J. Breukelman (245), Department of Biochemistry, University of Groningen, 9747 AG Groningen, The Netherlands. Christine E. Brown (589), Department of Molecular and Cell Biology, University of California, Berkeley, Berkeley, California 94720. Antonella Carsana (245), Department of Organic and Biological Chemistry, University Federico II of Naples, 80134 Naples, Italy. Mauro I. Ciglie (213), Department of Chemistry, Laboratory for Organic Chemistry, E. T. H. Zentrum, CH-8092 Zurich, Switzerland. Adrienne E. Clarke (191), Plant Cell Biology Research Centre, School of Botany, University of Melbourne, Parkville, Victoria 3052 Australia. Claudi M. Cuchillo (271), Departament de Bioqufmica i Biologfa Molecular, Facultat de Ci~ncies and Institut de Biologia Fonamental V. Villar-Palasi, Universitat Autbnoma de Barcelona, 08193 Bellaterra, Spain. Giuseppe D'Alessio (383,491), Dipartimento di Chimica Organica e Biologica, Universit~ Degli Studi di Napoli Federico II, 80134 Naples, Italy. Alberto Di Donato (383), Dipartimento di Chimica Organica e Biologica, Universit~ Degli Studi di Napoli Federico II, 80134 Naples, Italy. Adriana Furia (245), Department of Organic and Biological Chemistry, University Federico II of Naples, 80134 Naples, Italy. xi
xii
Contributors
Gary L. Gilliland (305), The Center for Advanced Research in Biotechnology, Biotechnology Institute and National Institute of Standards and Technology, University of Maryland, Rockville, Maryland 20850. Gerald J. Gleieh (425), Allergic Disease Research Laboratory, Departments of Immunology and Medicine, Mayo Clinic and Foundation, Rochester, Minnesota 55905. Carlos Gonztilez (343), Instituto de Estructura de la Materia, CSIC, 28006 Madrid, Spain. Pamela J. Green (163), Department of Energy Plant Research Laboratory and Department of Biochemistry, Michigan State University, East Lansing, Michigan 48824. Robert W. Hartley (51), Laboratory of Cellular and Developmental Biology, National Institute of Diabetes and Digestive and Kidney Diseases, National Institutes of Health, Bethesda, Maryland 20892. Monika Haugg (213), Department of Chemistry, Laboratory for Organic Chemistry, E. T. H. Zentrum, CH-8092 Zurich, Switzerland. Jan Hofsteenge (621), Friedrich Miescher Institut, CH-4002 Basel, Switzerland. Masachika Irie (101), Department of Microbiology, Hoshi College of Pharmacy, Tokyo 142, Japan. Thomas M. Jermann (213), Department of Chemistry, Laboratory for Organic Chemistry, E. T. H. Zentrum, CH-8092 Zurich, Switzerland. Ying-hong Liu (191), Department of Plant Science, Waite Institute, University of Adelaide, Adelaide SA 5005, Australia. Lelio Mazzarella (383), Dipartimento di Chimica, Universith Degli Studi di Napoli Federico II, 80134 Naples, Italy. Ed Newbigin (191), Plant Cell Biology Research Centre, School of Botany, University of Melbourne, Parkville, Victoria 3052, Australia. Allen W. Nicholson (1), Department of Biological Sciences, Wayne State University, Detroit, Michigan 48202. Victoria Nogu~s (271), Departament de Bioqufmica i Biologfa Molecular, Facultat de Ci~ncies Universitat Aut6noma de Barcelona, 08193 Bellaterra, Spain. Jochen G. Optiz (213), Department of Chemistry, Laboratory for Organic Chemistry, E. T. H. Zentrum, CH-8092 Zurich, Switzerland. Simon K. Parry (191), Plant Cell Biology Research Centre, School of Botany, University of Melbourne, Parkville, Victoria 3052 Australia. Renata Piccoli (383), Dipartimento di Chimica Organica e Biologica, Universita Degli Studi di Napoli Federico II, 80134 Naples, Italy.
Contributors
xiii
Sun-Ai Raillard-Yoon (213), Department of Chemistry, Laboratory for Organic Chemistry, E. T. H. Zentrum, CH-8092 Zurich, Switzerland. Manuel Rico (343), Instituto de Estructura de la Materia, CSIC, 28006 Madrid, Spain. James F. Riordan (445), Center for Biochemical and Biophysical Sciences and Medicine, Harvard Medical School, Boston, Massachusetts 02115. Jeff Ross (553), McArdle Laboratory for Cancer Research, Department of Pathology, University of Wisconsin-Madison, Madison, Wisconsin 53706. Alan B. Sachs (589), Department of Molecular and Cell Biology, University of California, Berkeley, Berkeley, California 94720. Jorge Santoro (343), Instituto de Estructura de la Materia, CSIC, 28006 Madrid, Spain. Robert H. Silverman (515), Department of Cancer Biology, Research Institute, The Cleveland Clinic Foundation, Cleveland, Ohio 44195. Melissa R. Snyder (425), Departments of Biochemistry and Molecular Medicine, Mayo Clinic and Foundation, Rochester, Minnesota 55905. Josef Soucek (213), Institute of Hematology and Blood Transfusion, 128 20 Prague 2, Czech Republic. Joseph Stackhouse (213), Department of Chemistry, Laboratory for Organic Chemistry, E. T. H. Zentrum, CH-8092 Zurich, Switzerland. Nathalie Trabesinger-Riif (213), Department of Chemistry, Laboratory for Organic Chemistry, E. T. H. Zentrum, CH-8092 Zurich, Switzerland. Katrin Trautwein (213), Department of Chemistry, Laboratory for Organic Chemistry, E. T. H. Zentrum, CH-8092 Zurich, Switzerland. Maria Vilanova (271), Unitat de Bioqufmica i Biologia Molecular, Departament de Biologia, Facultat de Ci6ncies, Universitat de Girona, 17071 Girona, Spain. Ira G. Wool (131), Department of Biochemistry and Molecular Biology, The University of Chicago, Chicago, Illinois 60637. Richard J. Youle (491), Biochemistry Section, Surgical Neurology Branch, National Institute of Neurological Disorders and Stroke, National Institutes of Health, Bethesda, Maryland 20892. Todd C. Zankel (213), PGEC-Plant Gene Expression Center, Albany, California 94710.
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Preface
Few enzymes have contributed so much to our knowledge of protein structure and function as ribonuclease A. The reasons for its remarkable impact are manifold, chief among them being its availability to the early giants of protein chemistry who chose it as the principal object of their attention. One of the first enzymes to be isolated (by Rene Dubos in 1938) and purified in crystalline form (by Moses Kunitz in 1939), it could be obtained in adequate amounts from bovine pancreas, a major byproduct of the beef industry (sweetbreads were never a favorite in the American diet even during the Great Depression era of the late 1930s). World War II interrupted basic research on ribonuclease, yet nevertheless provided unexpected benefits to protein science. As noted by Richards and Wyckoff in their 25-year-old review in Advances in Protein Chemistry, one of the original industrial-academic interactions--a warinspired cooperation between E. J. Cohn at Harvard Medical School and Armour, Inc.mled to the preparation of 1 kilogram of crystalline ribonuclease which Armour generously portioned out to researchers well into the 1950s. This not only determined the glorious fate of the enzyme, but played a key role in the development of the study of proteins. With the end of the war, scientists could once again return to the Mecca of protein chemistry, The Carlsberg Laboratory which was directed by one of the truly great giants, Kai LinderstrCm-Lang (whose centennial is being celebrated this year). Many who later went on to become leaders in their own right received their introduction to ribonuclease in his unique environment. It was also in this laboratory that the first experiments on limited proteolysis of pancreatic ribonuclease were performed and where Richards began his distinguished series of studies on ribonuclease S, an exceptionally rich source of important protein structurefunction information. The post-war resurgence of research witnessed a XV
xvi
Preface
period of many ribonuclease-related monumental firsts: the first purification of a macromolecule by ion-exchange chromatography, the first quantitative amino acid analysis of a protein (and, as a corollary, the first automated amino acid analyzer), and the first amino acid sequence of an enzyme. This rich legacy, much of it emanating from the laboratory of Moore and Stein at the Rockefeller Institute, was later supplemented by one of the first X-ray structures of an enzyme, the first investigation of a protein by NMR, the first chemical synthesis of an enzyme, and important firsts in protein engineering. In 1969, Barnard pointed out in his remarkable Annual Review of Biochemistry chapter that the announcement of the crystal structure of ribonuclease 1 was an epochal event that signaled an end to groping in the dark. The crystallographers had switched on the lights, and no longer would protein chemists be limited to indirect methods. In fact, Barnard proposed that henceforth all studies of enzymes should start from the crystal structure. (It is unlikely that he intended that enzymes that could not be crystallized should not be studied.) Thus, the structure of ribonuclease marked a defining moment in protein chemistry and it is only fitting that we recognize this achievement by portraying an up-to-date version of that structure on the cover of this volume. We are indebted to Gary Gilliland for providing such a striking rendition. The golden age of ribonuclease provided one of if not the most seminal of concepts in protein chemist r y - t h e thermodynamic hypothesis of Anfinsen and his colleagues. Based on the now classic observation that fully unfolded ribonuclease could be completely renatured, it asserts that the three-dimensional structure of a native protein is the one with the lowest Gibbs free energy and is determined by the amino acid sequence. It is difficult to estimate the amount of effort that has been expended in pursuit of this hypothesis, but it is safe to say that the folding problem has been the central theme and holy grail of the field for at least two decades. Ribonuclease A as well as barnase have featured prominently in this endeavor and still do, as described in Chapters 9 and 2 of this book, respectively. Another stellar achievement to emerge from this era, and one which culminated a long history of ribonuclease research at Rockefeller Instii We note that actually three ribonuclease structures were reported in 1967: one from Carlisle's group in London, a second from Harker's in Buffalo, and a t h i r d - - f o r ribonuclease S - - f r o m Richards and Wyckoff at Yale. Moreover, in the Journal of General Physiology of 1941, sandwiched between descriptions of the effects of hexylresorcinol on Valonia and the flicker response contour of the horned lizard, is the very first report of a crystallographic analysis of ribonuclease by Fankuchen at MIT.
Preface
xvii
tute/University, was the total synthesis of the protein by Merrifield. Not only did this substantiate the Anfinsen hypothesismthe synthetic enzyme folded properly and was catalytically active--but it opened entirely new avenues to the investigation of protein structure/function relationships. Barnard emphasized this by stating that the synthesis made it possible to test any residue for essentiality by replacement or deletion, thus revolutionizing active center studies. This, of course, antedated the advent of cloning. These signal accomplishments ultimately led to the awarding of four Nobel prizes for studies on ribonuclease. No other enzyme commands such distinction. Moore and Stein were recognized in 1972 "for their contribution to the understanding of the connection between chemical structure and catalytic activity of the active centre of the ribonuclease molecule." They shared the prize with Anfinsen who was honored "for his work on ribonuclease, especially concerning the connection between amino acid sequence and the biologically active conformation." Twelve years later Merrifield received the prize "for his development of methodology for chemical synthesis on a solid matrix" spectacularly exploited in the total synthesis of ribonuclease. Many others have expressed appreciation for the contributions of these outstanding scientists, including the Protein Society which features both the Stein and Moore Award and the Anfinsen Lecture at its annual meetings. We wish to express our own appreciation by dedicating this volume to these four titans who together established the basic tenets underlying the structures and functions of the ribonucleases. By the end of the 1960s bovine pancreatic ribonuclease had "yielded many of its secrets," and Barnard suggested that it was inevitable that the main interest in ribonucleases would shift to other proteins having this function. Such unusual prescience is amply corroborated by the broad range of ribonucleases described in this volume. Still, pancreatic ribonuclease continues to be both a mother lode of protein science and the matriarch of a superfamily. Moreover, it seems likely that as a group these proteins will remain the subject of intense investigation for many years to come, since, surprisingly, despite mountains of material published about them, little is actually known about their biological functions. It is, in fact, amazing that so many ribonucleases with special biological actions have been discovered. Why they should have antifertility, antitumor, antihelminthic, neurotoxic immunosuppressive, angiogenic, and ribosome-inactivating activities is not immediately intuitive. Yet enzymes with all of these properties and more are described in this
xviii
Preface
book in some detail. In addition, outstanding research still continues on the molecular properties of ribonuclease A, a clear reminder that it has not yet left the scientific mainstream and that the approach to the truth progresses asymptotically. The next phase of the ribonuclease saga promises to be another age of enlightenment in which the powerful tools of structural analysis combine with the seemingly endless capabilities of molecular and cellular biology to elucidate the intricacies of this fascinating group of enzymes. We have designed this volume to present a summary of the current status of ribonuclease-related research and to convey a sense of its dimensions and scope. The core chapters focus on the latest structures and functions of pancreatic ribonuclease. These are augmented by chapters on the newer, nondigestive members of the family, again with emphasis on structures and functions. Together these illustrate how nature has exploited a given molecular scaffold and adapted it to execute a range of biological functions. Yet other chapters are intended to demonstrate how ribonucleases continue to serve as excellent systems with which to uncover the secrets of protein chemistry. While ribonuclease A has been a pillar of the biochemical community, it and its superfamily of enzymes are just the tip of the ribonucleolytic iceberg. Nature seems to have been rather extravagant when it came to nucleases. After all, how many enzymes should it take to hydrolyze a molecule of R N A ? Of course we could also ask how many physicians it should take to care for a patient. Different problems require different specialists, sometimes even teams of specialists. So too with RNA. A perspective on the multiplicity of nonpancreatic-type enzymes involved in cellular R N A metabolism is provided by the chapters on E. coli ribonucleases, on mammalian R N A processing enzymes, and on m R N A turnover in yeast. A different ribonuclease, RNase L, is a prominent component of a host defense mechanism, and a chapter updating its unique properties adds important coverage. Other interesting ribonucleases that participate in a host of processes in plants, such as selfincompatibility, and having promising practical applications have been included as well. We could not encompass every enzyme that acts on RNA, but no book with a title like this would be complete without a chapter on the ribonuclease inhibitor, if only to show what has been called one of the most beautiful structures yet revealed by X-ray crystallography. We hope that this volume will serve to stimulate interest, provide an overview, and delineate a context for future ribonuclease research. We
Preface
xix
are, of course, grateful to all of our contributors who share our conviction that the ribonucleases are important biological modulators that deserve the attention of the scientific community. We regret that because of space limitations we were unable to invite the participation of the many other ribonucleologists who have significant stories to relate, but we thank them nonetheless for all that they have done for the field. We also thank Shirley Light of Academic Press for her tolerance and understanding as well as for being a constant source of encouragement and advice. Giuseppe D'Alessio James F. Riordan
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1 Escherichia coli Ribonucleases"
Paradigms for Understanding Cellular RNA Metabolism and Regulation ALLEN
W. NICHOLSON
Department of Biological Sciences Wayne State University Detroit, Michigan 48202
I. Introduction II. Gallery of Escherichia coli Ribonucleases III. Endoribonucleases A. Ribonuclease P B. Ribonuclease III C. Ribonuclease E D. Ribonuclease I E. Ribonucleases HI and HII F. Other Endoribonuclease Activities IV. Exoribonucleases A. Ribonuclease II B. Polynucleotide Phosphorylase C. Ribonuclease D D. Ribonuclease BN E. Ribonuclease T F. Ribonuclease PH G. Oligoribonuclease H. Ribonuclease R V. Ribonuclease Functional Roles A. Ribonucleases Involved in Ribosomal R N A Maturation B. Ribonucleases Involved in Transfer RNA Maturation C. Ribonucleases Involved in Messenger R N A Maturation and Turnover
1 RIBONUCLEASES: STRUCTURES AND FUNCTIONS
Copyright 9 1997 by Academic Press, Inc. All rights of reproduction in any form reserved.
Allen W. Nicholson
VI. Other Ribonucleases A. Ribonuclease Activity Associated with RNA Polymerase B. Prr Anticodon Nuclease C. Coliphage T4 RegB Nuclease VII. Ribonuclease Regulation and RNA Metabolic Control VIII. Conclusions and Prospects References
I. I N T R O D U C T I O N
Studies on the gram-negative bacterium Escherichia coli have provided a wealth of information on the myriad biological processes that constitute cellular metabolism (Neidhardt et al., 1987). This organism has been especially informative of the mechanisms of synthesis, maturation, function, and decay of RNA. Early molecular genetic studies of E. coli and the coliphages showed that RNA is the information link between DNA and protein (Brock, 1990), and that RNA processing is carried out by specific enzymes (e.g., see Robertson et al., 1972). The value of this organism as a paradigm for understanding cellular RNA function and metabolism has not diminished: the sequencing and analysis of the E. coli chromosome soon will be completed, which, along with the extensive genetic and biochemical data, should provide the first full description of the enzymes of RNA metabolism, and their functional roles. This prospect assumes added importance, because studies are now indicating the universality of the enzymes and mechanisms of RNA maturation and decay. Escherichia coli, like all other cells, makes a major commitment to RNA synthesis. Approximately 20% of the cell dry mass is RNA, most of which is ribosomal RNA (---81%) and tRNA (---14%). The most metabolically labile fraction is mRNA, which comprises only 4% of the RNA (Neidhardt et al., 1990). The instability of mRNA is underscored by the fact that many species begin to decay before their synthesis is complete. The biochemical properties and functional roles of the E. coli ribonucleases that participate in RNA maturation and decay are examined in this chapter. Acknowledging the likelihood that many readers are studying eukaryotic ribonucleases, the similarities of the E. coli ribonucleases with specific eukaryotic activities will be emphasized where
1 Escherichia coli Ribonucleases
possible. The biochemical properties of the ribonucleases and their involvement in the variety of R N A metabolic pathways will be summarized. Several recently characterized, atypical RNase activities will be examined. Currently unanswered questions concerning RNase structure and function, and the pathways of R N A processing and decay, will be pointed out as guides for future studies. The reader also is referred to several excellent reviews of R N A processing and decay in E. coli and other eubacteria (Gegenheimer and Apirion, 1981; King et al., 1986; Ehretsmann et aL, 1992a; Petersen, 1992; Belasco and Brawerman, 1993), and the associated ribonuclease activities (Deutscher, 1993a-c).
II. G A L L E R Y OF Escherichia coli R I B O N U C L E A S E S
Tables I and II list the currently known endoribonucleases and exoribonucleases, respectively, of E. coli. The most relevant biochemical and genetic properties are also provided [see also Linn and Deutscher (1993)]. The gallery is incomplete; there will be new entries, and more information obtained on existing entries. Several of the poorly characterized activities may be alternate manifestations of the same ribonuclease. This situation in part reflects the fact that different laboratories have used different protein purification schemes and substrates in characterizing the ribonuclease activities (Deutscher, 1993b).
III. E N D O R I B O N U C L E A S E S
The endoribonucleases (Table I) are biochemically diverse, and include phosphodiesterases as well as phosphotransferases. Some require a divalent metal ion for activity, whereas others are active in the presence of EDTA. One enzyme contains a catalytic RNA subunit.
A. Ribonuclease P
RNase P was the first ribonuclease shown to be an RNA-processing enzyme (Robertson et al., 1972). Studies on this enzyme have provided
TABLE I
Escherichia coil Endoribonucleases" Gene a rnpA, 83; rnpB, 70 rnc, 55
Mg 2+
pre-tRNAs
t R N A maturation
Phosphodiesterase, 5'
C5 protein, 14 M1RNA. 377 nt 25.6, c~2
Mg 2+
dsRNA
--
Phosphodiesterase, 5'
120
me, 24
Mg 2+
mRNA, 9S r R N A
EC 3.1.27.6 EC 3.1.26.4
Phosphotransferase, 3' Phosphodiesterase, 5'
27 17.6
rna, 14 rnh, 5.1
Mg 2+
ssRNA R N A / D N A hybrids
mRNA, r R N A maturation 5S r R N A maturation, m R N A degradation R N A degradation Replication, D N A repair
Phosphodiesterase, 5' Phosphotransferase, 3' (?) Phosphotransferase(?), 3'
23.2 27 24 31
rnhB, 4.5 NR NR NR
(?)
(?)
Phosphodiesterase, 5'
120, a/3 (?)
EC number
RNase P
EC 3.1.26.5
Phosphodiesterase, 5'
RNase III
EC 3.1.24
RNase E RNase I, I* RNase HI RNase RNase RNase RNase
HII M R IV, F
RNase P2, O, PIV, PC RNase N
m
Mechanism b
Cofactor(s)
Mg2 ~-
Substrates
(?) (?) (?)
R N A / D N A hybrids ssRNA ssRNA ssRNA
t R N A maturation (?)
NR
ssRNA
t R N A maturation
NR
ssRNA, dsRNA
(.9)
"See also Deutscher (1993b,c). b The 5', 3' notation indicates whether cleavage leaves a 5'- or 3'-phosphate-containing RNA terminus. c Numbers refer to subunit mass (kDa); subunit structure is indicated. a Numbers refer to map position (minutes) on the E. coli chromosome): NR, not reported. See text for further discussion. e
Function(s) e
Structure'-
Name
TABLE
II
Escherichia coil E x o r i b o n u c l e a s e s ~ Name
EC number
RNase II
EC 3.1.27.1
Phosphodiesterase, 5'
70
rnb, 28
Mg 2+, Mn 2+
ssRNA, poly(A)
PNPase
EC 2.7.7.8
Phosphotransferase, 5'
85, a 3
pnp, 69
Phosphate, Mg 2+
ssRNA
Phosphodiesterase, 5' Phosphodiesterase, 5' Phosphodiesterase, 5' Phosphotransferase, 5' Phosphodiesterase, 5'
42.7 60 23.5, a2 25.5, a2 (?) 38
rnd, 40 rbn, 88 rnt, 36 rph, 81.7 NR
Mg 2+ Co 2§ acidic pH Mg 2+ Mg 2§ Mf +
(?)
80
NR
pre-tRNAs pre-tRNAs pre-tRNAs pre-tRNAs Short oligoribonucleotides Ribosomal RNA, homopolymers
RNase D RNase BN RNase T RNase PH OligoRNase RNase R
m m
m
Mechanism b
Structure C
Gene d
Cofactor(s)
Substrates
Function(s) e m R N A degradation, t R N A maturation m R N A degradation, t R N A maturation t R N A maturation t R N A maturation t R N A maturation t R N A maturation R N A degradation (?) R N A degradation (?)
a See also Deutscher (1993a-c). RNase PIII, Q, and Y were not included, because they are probably manifestations of RNase II and/or RNase D (Deutscher, 1985). b The 5' notation indicates that cleavage leaves a 5'-phosphate on the R N A terminus. c Numbers refer to subunit mass (kDa); subunit structure is indicated. d Numbers refer to map position (minutes) on the E. coli chromosome; NR, not reported. e See text for further discussion.
6
Allen W. Nichoison
subsequent paradigm-shattering surprises, including the discovery that it has an RNA subunit (Stark et al., 1978), and that the R N A catalyzes the processing reaction (Guerrier-Takada et al., 1983). RNase P is predominantly involved in tRNA maturation, catalyzing the endonucleolytic cleavage of tRNA precursors to remove the 5' leader segment, creating the mature tRNA 5' ends. Because all tRNAs undergo this processing event, it is not surprising that RNase P is found throughout the Eukarya and Archaea, as well as the Bacteria (Pace and Brown, 1995). The involvement of RNase P in tRNA maturation is further discussed in Sections V,A and V,B. There is also evidence that RNase P is involved in the processing and stabilization of a cellular m R N A (Alifano et al., 1994). The complexities of RNase P structure, mechanism, and function have been excellently reviewed from several perspectives (Altman, 1990; Pace and Smith, 1990; Cech, 1993; Pace and Brown, 1995). RNase P occupies a unique position in the E. coli ribonuclease gallery, because it is a ribonucleoprotein enzyme. The single protein subunit, termed C5 protein, is encoded by the rnpA gene, which maps at 83 minutes on the E. coli chromosome. C5 protein is basic and has a molecular mass of approximately 14 kDa. The single RNA subunit, termed M1 RNA, is 377 nucleotides in length, with a molecular mass of approximately 130 kDa. The RNase P holoenzyme is a 1 : 1 complex of C5 protein and M1 RNA (Talbot and Altman, 1994). From these considerations it is clear that the bulk of RNase P is RNA. M1 RNA is encoded by the rnpB gene, mapping at 68 minutes. M1 RNA is transcribed as a precursor, which is matured in part by RNase E (Lundberg and Altman, 1995). Both the protein and RNA are essential for RNase P activity in vivo (Altman, 1990; Pace and Smith, 1990). A remarkable finding was that the RNA subunit of RNase P is a biochemical catalyst capable on its own of the accurate processing of tRNA precursors in vitro (Guerrier-Takada et al., 1983). Although C5 protein is required for RNase P function in vivo, it can be replaced by high salt concentrations in vitro. However, the catalytic activity of the isolated RNA subunit may not be a universal feature: studies on the RNA subunits from eukaryotic and archaeal RNase P activities have not demonstrated measurable catalytic activity (Pace and Brown, 1995). It is possible in these instances that the RNA subunits are indeed catalytic, but that the protein subunit is required to promote the active conformation. Alternatively, the protein may provide one or more side chains that participate in catalysis (Cech, 1993; Pace and Brown, 1995). RNase P is a phosphodiesterase; it activates a water molecule to hydrolyze the scissile phosphodiester bond, leaving 5'-phosphate, 3'-hydroxyl
1
Escherichia coli Ribonucleases
R N A termini. Divalent metal ion (Mg 2+) is required for the cleavage reaction. Biochemical studies indicate that RNase P requires at least three Mg 2+ ions to catalyze the reaction (Smith and Pace, 1993). Ca 2+ supports substrate binding as well as Mg 2+, but only weakly supports cleavage activity (Smith et al., 1992; Smith and Pace, 1993). Because M1 R N A is sufficient to provide efficient and accurate processing in vitro, it carries the substrate binding and the catalytic sites, which include divalent metal ion sites. The mechanistic basis for RNase P catalysismas may be the case with many other phosphodiesterases (Gerlt, 1993)mmay be provided by Mg 2+ ions that are precisely placed in the M1 R N A structure. The biochemical activity of M1 RNA, as with any biomolecule, derives from its three-dimensional shape. The sequence and proposed secondary structure of M1 R N A are shown in Fig. 1A. The secondary structure was solved by enzymatic and chemical structure-probing studies, mutational analysis, and computer-assisted folding (Pace and Brown, 1995). Phylogenetic analysis was also crucial in verifying and refining the secondary structure model, as well as in identifying long-range base pair interactions and pseudoknots, and in deriving a minimum consensus RNase P RNA structure, shown in Fig. 1B (Pace and Brown, 1995). Specific base-paired stem structures and conserved nucleotides in formally single-stranded regions are the hallmarks of the consensus secondary structure. In the absence of a crystal structure of RNase P or RNase P RNA, molecular modeling studies have provided a preliminary three-dimensional structural model of the RNA subunit (Harris et al., 1994; Westhof and Altman, 1994). The models will be further refined as additional biochemical and genetic data are obtained. How are the pre-tRNA substrates recognized by RNase P? Studies show that RNase P recognizes the tertiary structure in the mature domain of the pre-tRNAs. M1 RNA establishes contacts with the helix consisting of the coaxially stacked tRNA acceptor stem and common arm (containing the T ~ C loop) (Altman, 1990; Pace and Smith, 1990; Pace and Brown, 1995). This portion of the tRNA structure is sufficient for recognition, because RNase P can accurately process small substrates containing just the common arm/acceptor stem, the 5' leader segment, and the 3' CCA end (Altman, 1990; Pace and Smith, 1990). It is interesting to note that there is no strong evidence that Watson-Crick base pairing between enzyme and substrate is required for processing. C5 protein plays an essential in vivo role in substrate recognition. The basic nature of the C5 protein reduces the charge repulsion between substrate and RNA
Allen W. Nicholson CA
A
160
G A" c-G G-.-C c--G c-G c-G G.U
A
Escherichia coli
AGue'CA 1)12 Gc-G c-G GeUA 140 C -GA C
A'AAC-~"U~AA 180 A
o UA
P13
a_Qu oe Q,
Q
A G
A
Ul~l ,.. G I c A C G~" G A
A
C
G A C
"~
G 200 CGc.. P14 le. ur;. ,.,
A AC
Gu ~ Pll CA C AG~"A AA U G G 1~) CA C A "- " - A n " U . ~ P 1 0
u C
_Qua! r ~-UQQ
~Cn. / U 220/ U - A C A A
AG~;~oCCAc~P7 1)5 G " / / vC GGAGCAA 1)8 U~..~P C..- A
"Au -C n
. c - ,~
u ,-IO0"CA A U
.,,,
CCUC A GI
UCGG G-30 o A U
~.,,,,u,e -G
~1
c
" ~
1~1
U
P18
..
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G A
G
/
, C~.
/
~1
" G I ~
P3
U
GAGCCAGUGA
\uccaauc~uuA~
_6~1~
O|
"
uI
cGGGGGA GACG GCGGAGUGG ~ I 9I I I I I I I e l I 9I I I I ~ 40 uCUCCU CUGC UGCUUCGCCGc_ G G 20 U -A
A UA
. %A . p ] 7
CCCAAGur//~'A_
" AU" ,"~ Q(;~
o ,.,
\ ~ 8O.u I
0
PI5 AAAUAG Pl' GGCC ^ GGGU
,,,,
o'_e"uQ'-'
f
O,.
2a0"
n~ rq
C - G-~.to A-U I GAAGCUGACC A G-C C A C G AC A I I I el I I I I I 9 CUUUGACUGGC G A A U A ,y u c G Q c c C, A c
PI
C U
36o
Fig. 1. (A) Secondary structure of M 1 R N A of E. coliRNaseP. Double-helicalelements are indicated by Pl-P12 notation. (B) Phylogenetic minimum consensus secondary structure for bacterial RNase P RNA subunits (Pace and Brown, 1995). Conserved doublehelical elements are indicated by P1-Pl2. The conservation of specific nucleotides or nucleotide positions is indicated. Reprinted with permission from American Society for Microbiology.
1
Escherichia coli R i b o n u c l e a s e s
o 9
B 9
P12o
G
a
oA
9
9 0
"o
9 ~
O
A
U G
9A g ~ " e a 9
GAUC - 100% conserved nts gauc - 80% conserved nts 9- nt present in all RNAs o - nt present in 80% of RNAs
a9
CG 9 ~ o
9
Cx\oO g g
c a9
o o o ' P~__ (
99
~x ~o
9
A
P1 ] a
c
9
g ~149
C
i ~149149
o o O/o//
P8o
~
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"o
o~"o
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9
o~
oO !1
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Go
uC
9 9 9
9
o
g
~
o
a
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o
G
P3
9
9 oooooooG
9 9
A
I I I I I o I I
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A
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G O _ _ Y- O--O O--O
p2o-o
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oooo ooooooo I I I I, I,,,,.
OOOO
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o-o 9
o oooAC
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3" Fig. 1.
A
g
9 9 9
(Continued)
subunit, and promotes the productive binding of a wider range of substrate types, including the precursor to 4.5S RNA (Reich et al., 1988; Altman, 1990).
B. Ribonuclease III
Ribonuclease III is a double-stranded (ds) RNA-specific endoribonuclease. Originally detected as an activity that hydrolyzes dsRNA
10
Allen W. Nichoison
in vitro (Robertson et aL, 1968), the enzyme was later shown to
participate in ribosomal RNA maturation, cleaving the T7 phage polycistronic early mRNA precursor [reviewed by Court (1993) and Nicholson (1996)]. RNase III processes other phage RNAs, as well as cellular mRNAs, and plasmid transcripts, including sense-antisense RNA duplexes. The functions encoded by the processed cellular mRNAs participate in the synthesis, maturation, translation, and turnover of mRNA (Nicholson, 1996). RNase III can regulate gene expression because it controls the functional activities and physical half-lives of several of the mRNAs. The RNase III gene (rnc) maps at 55 minutes and is cotranscribed with the downstream era and recO genes. Cells that lack RNase III are viable, indicating that alternate processing pathways can provide functional rRNA and that RNase III cleavage of other cellular transcripts is not essential, at least under typical laboratory growth conditions. RNase Ill-related activities are apparently ubiquitous in the bacteria, and a homologous enzyme has been identified in yeast (Iino et al., 1991; Rotondo et aL, 1995). The yeast enzyme, termed Pacl, participates in sporulation and small nuclear RNA maturation. However, the substrates for Pacl have yet to be determined, dsRNA-cleaving activities exist in mammalian cells [reviewed by Nicholson (1996)], but it is not clear whether any of these activities are mechanistically or functionally similar to RNase III. The RNase III structural gene, rnc, has been cloned and sequenced (Nashimoto and Uchida, 1985), and RNase III can be overexpressed from plasmid vectors without toxic effects (Chen et aL, 1990; March and Gonzalez, 1990; Li et al., 1993). Purification of active enzyme is easily accomplished by affinity chromatography on dsRNA-agarose (Dunn, 1976) or by Ni 2+ affinity column purification of the N-terminal histidine-tagged protein (A. W. Nicholson and H. Li, 1995, unpublished). RNase III is active as a homodimer (Dunn, 1976), with a subunit size of 226 amino acids (dimer molecular mass, approximately 50 kDa) (Nashimoto and Uchida, 1985). RNase III is a phosphodiesterase and cleaves substrate to provide 5'-phosphate, 3'-hydroxyl product termini. The only required cofactor is Mg 2+, with Mn 2+, Co 2+, and Ni 2+ as functional substitutes (Court, 1993; Nicholson, 1996). A possible involvement of at least one carboxyl group in the RNase III catalytic mechanism is suggested by the behavior of the rnc[Ell7K] (rnc70) mutant, in which glutamic acid-ll7 is changed to lysine. This mutant can bind but cannot cleave substrate (Court, 1993; Li and Nicholson, 1996).
1
Escherichia coli Ribonucleases
11
The RNase III polypeptide contains a dsRNA-binding motif (dsRBM) (St. Johnstone et aL, 1992) near the carboxyl terminus. The dsRBM is a sequence of 65-70 amino acids, also present in other proteins that bind dsRNA (St. Johnstone et al., 1992). The dsRBM is probably the primary substrate-binding determinant, in that the isolated dsRBM of RNase III can bind dsRNA in vitro but is incapable of catalyzing cleavage (A. Amarasinghe and A. W. Nicholson, 1995; unpublished). The solution structure of the RNase III dsRBM has been recently solved by nuclear magnetic resonance (NMR) techniques, which show that it possesses an c~/3/3/3c~ structure (Kharrat et aL, 1995). The domain is a tightly folded ellipsoid, with the two c~ helices packed on one side of the antiparallel/3 sheet. Extensive hydrophobic contacts between the /3 sheet and the c~ helices apparently stabilize the structure. Site-directed mutagenesis of a dsRBM from another protein indicate that helix c~2 contains residues that contact dsRNA (Bycroft et aL, 1995). All RNase III processing signals feature a more or less regular dsRNA element, within which specific cleavage occurs. Exhaustive digestion of dsRNA yields short duplexes, averaging 10-15 bp in length, or little over one turn of the A helix (Robertson, 1982; Dunn, 1982). The presence of specific sequence and structural elements, in addition to dsRNA, establishes the specific reactivity pattern. It has been determined for one processing substrate that the length of the dsRNA element influences processing reactivity but not specificity. Thus, shortening the dsRNA element reduces the cleavage rate, but does not alter cleavage site choice (Chelladurai et al., 1993). A formal similarity between RNase III and the D N A restriction endonucleases has been suggested, because there may be an involvement of base pair sequence in processing reactivity (Court, 1993; Li et al., 1993; Nicholson, 1996). A proposed consensus RNase III processing substrate contains a conserved base pair sequence element which exhibits hyphenated dyadic symmetry about the cleavage sites (Krinke and Wulff, 1990). However, the base pair conservation is weak, suggesting that if the processing signal identity elements involve specific base pairs, they may be somewhat degenerate (Chelladurai et al., 1991). Whereas dsRNA undergoes coordinate double cleavage, RNase III processing substrates with internal loops undergo single cleavage. Perhaps single cleavage occurs because only one of the two strands in the internal loop can be placed in a catalytic site (Li and Nicholson, 1996).
12
Allen W. Nicholson
C. Ribonuclease E
Ribonuclease E is a major participant in mRNA decay, but also plays a role in rRNA maturation. RNase E was originally identified as an activity that cleaved 9S rRNA in vitro, yielding the immediate precursor to 5S rRNA (Ghora and Apirion, 1978). It was subsequently shown to be an essential activity, involved in the decay of cell, phage, and plasmid transcripts (reviewed by Melefors et aL, 1993). Immunological analysis indicates that RNase E-related activities are ubiquitous in the prokaryotes (Taraseviciene et al., 1994), and eukaryotic RNase E-related activities are likely (see later). RNase E is a phosphodiesterase, requiring a divalent metal ion (Mg 2+, Mn 2+) to cleave RNA, leaving 5'-phosphate, 3'-hydroxyl termini (Misra and Apirion, 1979). The single gene encoding RNase E ( m e ) maps at 23.5 minutes (Misra and Apirion, 1978; Casaregola et al., 1992). The rne gene is coallelic with the a m s (altered mRNA stability) locus, which was previously shown to control the rate of bulk mRNA decay (Mudd et al., 1990; Babitzke and Kushner, 1991; Melefors and von Gabain, 1991; Taraseviciene et al., 1991" Mackie, 1991), and with the h m p l gene, encoding a high molecular mass polypeptide, which cross-reacts with yeast heavy chain myosin antibodies (Casaregola et al., 1992). RNase E autoregulates its expression by cleavage of the rne transcript (Mudd and Higgins, 1993; Jain and Belasco, 1995). The rne gene has been cloned, and the encoded polypeptide overexpressed and purified (Cormack et al., 1993). The purified rne polypeptide can cleave substrate with the same specificity as native RNase E (Cormack et al., 1993). There is no requirement for any cofactor other than Mg 2§ However, there is evidence for the association of RNase E with other polypeptides in vivo, forming a macromolecular, multienzymic complex involved in RNA degradation (see Section V,D). RNase K, an activity related to RNase E, was shown to participate in site-specific cleavage of specific mRNAs and to control mRNA half-lives (Lundberg et al., 1990). However, further analysis revealed that RNase K is a proteolytic fragment of RNase E (Carpousis et al., 1994; Lundberg et aL, 1995). The rne gene sequence has been determined and encodes a polypeptide of 1061 amino acids with a predicted molecular mass of 116 kDa [(Casaregola et aL, 1992); G. Mackie, 1995, personal communication].
1
Escherichia coli Ribonucleases
13
The mobility of the rne polypeptide in SDS-polyacrylamide gels is anomalously slow (approximately 180 kDa), which may derive from unusual sequence and structural features near the carboxyl terminus (Casaregola et aL, 1992). The position of the rne polypeptide in twodimensional gels indicates a species with a pI value of 5.0 (Taraseviciene et al., 1994). Two temperature-sensitive mutations (ams-1 and rne3071) map within the N-terminal domain of the rne polypeptide, near a consensus nucleotide-binding site (McDowall et aL, 1993). These mutations block RNase E action in vivo at the nonpermissive temperature, perhaps by disrupting specific interactions with other factors, rather than directly inactivating the catalytic mechanism (McDowall et al., 1993). The antigenic cross-reactivity of the rne polypeptide to myosin-specific antibodies suggests that RNase E may be involved in RNA movement, in addition to RNA decay (Casaregola et aL, 1992). The cleavage reactivity and specificity determinants in RNase E substrates are only partially defined. The target site must be single-stranded for cleavage reactivity (Cormack and Mackie, 1992). Comparison of RNase E cleavage site sequences yielded the consensus 5'-RAUUW-3' (where R is A or G, and W is A or U), within which occurs the scissile phosphodiester bond (Ehretsmann et al., 1992b). However, considerable sequence variation is allowable, thus the "consensus" sequence may exhibit considerable degeneracy, as long as it is A,U-rich (Lin-Chao et al., 1994; McDowall et al., 1994). Third, the target sites are often flanked by secondary structure, such as RNA hairpins (Cormack and Mackie, 1992). These elements may directly participate in enzyme binding, or enforce the single-stranded nature of the cleavage sites (Cormack and Mackie, 1992). Because RNase E can accurately process short, singlestranded RNA oligonucleotides that lack flanking stem-loops, such secondary structures are not strictly required, at least for in vitro reactivity (McDowall et al., 1995). The accurate prediction of RNase E cleavage sites will depend on knowledge of the RNase E-substrate interaction at the atomic level. RNase E-related activities exist in mammalian cells. The human ard1 gene encodes a basic, proline-rich 13.3-kDa molecular mass protein, which is similar to a sequence in E. coli RNase E (Wang and Cohen, 1994). Expression of the ard-1 gene in E. coli complements rne mutants: bulk mRNA decay rates are restored to wild type, and the site-specific
14
Allen W. Nicholson
cleavages produced in vivo and in vitro are essentially the same as that of RNase E (Wang and Cohen, 1994). Another study described an activity in human cell extracts, which cleaves 9S RNA and o m p A m R N A in vitro with the same specificity as RNase E (Wennborg et al., 1995). Antibodies to RNase E recognize the human protein, which exhibits a molecular mass of approximately 65 kDa (Wennborg et al., 1995). This enzyme cleaves within the 5 ' - A U U U A - 3 ' sequence, which is a cis-acting determinant of mammalian m R N A stability and is reminiscent of the A,U-rich RNase E cleavage consensus sequence (Wennborg et aL, 1995).
D. Ribonuclease I
RNase I is a periplasmic enzyme that nonspecifically cleaves RNA, yielding mononucleotides and short oligonucleotides (reviewed by Shen and Schlessinger, 1982). The enzyme is a phosphotransferase, activating the ribose 2'-hydroxyl group to attack the adjacent phosphodiester, cleaving the RNA chain and creating 2',3'-cyclic phosphodiester groups. The products are subsequently hydrolyzed to the 3'-monophosphates (Shen and Schlessinger, 1982). RNase I does not require divalent metal ion for activity (Shen and Schlessinger, 1982). The gene encoding RNase I (rna) maps at 14.3 minutes and the predicted amino acid sequence is 245 amino acids, with a predicted molecular mass of 27.1 kDa (Meador and Kennell, 1990). Escherichia coli strains carrying a deletion of the rna gene are viable, and overexpression of RNase I from recombinant plasmids is not toxic to the cell (Zhu et al., 1990). RNase I- strains were used to show that there are one or more additional EDTA-resistant RNase activities in addition to RNase I and RNase I* (Zhu et al., 1990) (see also below). Purified RNase I is active as a monomer and exhibits an apparent molecular mass of 27 kDa in SDS-containing polyacrylamide gels (Meador et al., 1990). The enzyme is synthesized with an N-terminal leader peptide, which is cleaved on secretion into the periplasm (Meador and Kennell, 1990; Meador et aL, 1990). The function of RNase I in RNA metabolism is unclear. However, RNase I may allow the cell to utilize the nucleotides from R N A present in the external environment. Insight on the mechanism of RNase I may be provided by analysis of the crystallized protein (Lim et al., 1993).
1
Escherichia coli Ribonucleases
15
RNase I*, a modified form of RNase I, can be isolated from the soluble cytosolic fraction of spheroplasts (Cannistraro and Kennell, 1991). RNase I* exhibits a gel electrophoretic mobility (27 kDa) identical to that of RNase I, and deletion of the rna gene removes RNase I* activity (Cannistraro and Kennell, 1991). However, RNase I* and RNase I exhibit different physicochemical properties, including differential reactivities toward sulfhydryl reagents. The data indicate that RNase I* is expressed from the rna gene and has the signal peptide removed. However, RNase I* is not secreted through the cytoplasmic membrane, but rather is maintained in the cytoplasm in a form exhibiting free sulfhydryl groups (Cannistraro and Kennell, 1991). RNase I* activity may also be masked by a specific inhibitor (Beppu and Arima, 1969). The role of RNase I* may be in intracellular R N A turnover and nucleotide salvage (see Section VII). However, because RNase I* is not essential, other degradative enzymes are sufficient.
E. Ribonucleases HI and HII
The ribonucleases H refer to a ubiquitous class of phosphodiesterases that endonucleolytically cleave the R N A strand of R N A - D N A hybrids. Escherichia coli contains two RNase H activities, RNases HI and HII (for a recent review, see Hostomsky et al., 1993). Although the functional roles of these enzymes have not been fully defined, RNase HI is the best understood of the E. coli RNases from the standpoint of physical structure and enzymatic mechanism. RNase HI has been implicated in removing R N A primers within R N A - D N A (R-loop) structures formed during initiation of chromosome replication at oriC. RNase HI may also remove R N A primers at the 5' ends of lagging strand D N A synthesis fragments. However, because RNase HI- E. coli strains are viable, other activities, including recombination nucleases, DNA polymeraseassociated exonuclease activities, and RNase HII, may provide alternate routes of R-loop and R N A primer removal. One role for RNase HI has, however, been clearly defined: it is responsible for R N A primer 3' end formation for ColE1 plasmid replication (Hostomsky et al., 1993). RNase HI is active as a monomer and requires Mg 2+ for catalysis. RNase HI cleavage creates R N A products with 5'-phosphate, 3'hydroxyl termini (Hostomsky et al., 1993). RNase HI is encoded by the
16
Allen W. Nicholson
r n h A gene, at 5 minutes on the genetic map. The r n h A gene has been cloned and sequenced (Kanaya and Crouch, 1983) and the protein purified (Itoh and Tomizawa, 1982). The polypeptide consists of 155 amino acids, with a predicted molecular mass of 17.6 kDa. RNase HI has been crystallized and its structure determined (Yang et al., 1990; Katayanagi et aL, 1990). The protein consists of a five-stranded, mixed /3 sheet, with five c~ helices, four of which are positioned on one face of the/3 sheet. The amino acid side chains implicated in the RNase HI catalytic mechanism have been identified by in vitro mutagenesis and physical structure analysis. Specifically, a triad of carboxylate side chains, Asp-10, Glu-48, and Asp-70, clustered on the concave surface of the/3-sheet, are essential for catalysis (Katayanagi et al., 1993a,b). Two cleavage mechanisms have been proposed. One mechanism involves a single Mg 2+ ion, which coordinates the scissile phosphodiester, and Asp-70, which activates the water nucleophile (Fig. 2) (Nakamura et al., 1991; Oda et al., 1993). The other proposed mechanism involves two divalent metal ions (Yang et al., 1990; Davies et al., 1991), and is utilized by other phosphotransferases (Kim and Wyckoff, 1991; Beese and Steitz, 1991). The latter mechanism is based in part on the structural similarity of RNase HI with the RNase H domain of HIV-1 reverse transcriptase and the exonuclease domain of DNA polymerase I (Yang et al., 1990; Davies et al., 1991). Because current evidence indicates that only one Mg 2§ ion binds in the catalytic site (Huang and Cowan, 1994; Uchiyama et al., 1994; Katayanagi et al., 1993a; Oda et al., 1991), the first mechanism is favored. However, additional studies are required to define more precisely the catalytic mechanism and divalent metal ion participation. RNase HII is encoded by the r n h B gene, at 4.5 minutes on the genetic map (Itaya, 1990). The predicted amino acid sequence of RNase HII (23.2 kDa molecular mass) exhibits only a modest similarity (approximately 17%) to RNase HI. The protein exhibits a gel electrophoretic mobility of 24 kDa, in agreement with the predicted molecular mass (Itaya, 1990). The role of RNase HII remains to be determined, although it may provide a backup function for RNase HI or may perform specialized functions. The occurrence of more than one RNase H activity in E. coli suggests a gene duplication event, followed by sequence (and possible functional) divergence. The occurrence of two RNase H activities in E. coli is consistent with the widespread occurrence of multiple RNase H activities in other prokaryotic and eukaryotic cells,
1
17
Escherichia coli Ribonucleases
0
B
/ O, I
Ixo-~ 0
\ I
9
"H
pro-Rp H--O
\
H
O
, @ I I
o-"" H
~
B
H I
I Il
O"
O
O.
H
Fig. 2. Proposed catalytic mechanism of RNA cleavage by E. coli RNase HI (Uchiyama et aL, 1994). The water-activating amino acid residue (Asp-70) and single Mg 2+ ion are indicated. Reprinted with permission.
and is reminiscent of the similarity of RNase I and RNase M (see Section Ill,F).
F. Other Endoribonuclease Activities
The following activities are only partially characterized and their functional roles are obscure. Some of the activities may be manifestations of the same enzyme, partially purified using different protocols, and using different substrates. RNase M cleaves YpA bonds (Y = C or U) in unstructured RNAs, yielding 3'-phosphate, 5'-hydroxyl RNA termini (Cannistraro and Kennell, 1989). RNase M has a molecular mass of approximately 26 kDa
18
Allen W. Nichoison
and does not require divalent metal ion for activity (Cannistraro and Kennell, 1989). RNase M is distinct from RNase I, in that it occurs in RNase I- strains, and exhibits thermal stability and substrate specificity different from that of RNase I (Cannistraro and Kennell, 1989). However, RNase M is structurally similar to RNase I: both exhibit essentially the same molecular mass and have similar tryptic peptide maps (Meador et al., 1990). It was speculated that RNase M and RNase I are related by a gene duplication event, followed by sequence and functional divergence (Meador et al., 1990). RNase M and RNase I share some functional similarity to pancreatic RNase A, which is a cyclizing-decyclizing phosphotransferase that recognizes YpN bonds (Cannistraro and Kennell, 1989). RNase R (R for residual) represents a low-level activity present in RNase I- cell extracts (Srivastava et al., 1992). The activity degrades a variety of substrates in the absence of added divalent metal ion and exhibits an apparent molecular mass of approximately 24 kDa. The levels of RNase R activity are approximately 100-fold lower than that for RNase I (Srivastava et al., 1992). The functional role of RNase R has not yet been established, and a relation to RNase M is possible (Deutscher, 1993b). RNase IV is an activity that cleaves phage R17 RNA in a site-specific manner (Shen and Schlessinger, 1982). RNase IV does not require monoor divalent metal ion for activity and may be related to RNase F, which catalyzes specific cleavage of a phage T4 RNA, creating 3'-phosphate, 5'hydroxyl product termini. RNase F is approximately 31 kDa in molecular mass (Deutscher, 1995). The roles of RNase IV and RNase F in cellular RNA metabolism remain to be determined. RNase P2 catalyzes sitespecific cleavages in 5' leader sequences of tRNA precursors and may be related to RNase PIV (Deutscher, 1995). RNase PC was detected by its ability to cleave phage T4 tRNA transcripts in vitro (Schmidt and Apirion, 1984), and may be another manifestation of RNase P2 (Deutscher, 1993b). RNase N digests single- or double-stranded synthetic and natural RNAs, providing 5'-mononucleotides (Misra and Apirion, 1978). The enzyme is active in the absence of added divalent metal ion. However, an involvement of divalent metal ion is suggested because RNase N activity is inhibited by E D T A (Misra and Apirion, 1978). The enzyme is approximately 120 kDa in size and may consist of two subunits of approximately 60 kDa each (Misra and Apirion, 1978). The functional role of RNase N is unknown.
1
Escherichia coli Ribonucleases
19
IV. E X O R I B O N U C L E A S E S
Table II lists six well-characterized 3' --> 5' exoribonucleases and two less well-described activities. All are hydrolytic phsphodiesterases, except two that are phosphorolytic [polynucleotide phosphorylase (PNPase) and RNase PH]. No 5' ~ 3' exoribonucleases have been characterized.
A . R i b o n u c l e a s e II
Ribonuclease II, the major exoribonucleolytic activity in cell-free extracts, hydrolytically degrades RNA chains from the 3' end, releasing 5'-rNMPs (Shen and Schlessinger, 1982). The enzyme is active in monomeric form, and requires divalent metal ions (Mg 2§ Mn 2§ and monovalent cations (K § NH4 § for maximal activity. The enzyme is processive, with the homoribopolymer poly(A) as the most reactive substrate (Shen and Schlessinger, 1982; Cannistraro and Kennell, 1994; Coburn and Mackie, 1996). RNase II is encoded by the rnb gene, which maps at 29 minutes (Donovan and Kushner, 1983; Zilh~o et al., 1995). The rnb gene has been cloned and sequenced and the protein overexpressed and purified (Zilh~o et al., 1993; Coburn and Mackie, 1996). The DNA sequence predicts a protein of 644 amino acids of 67.6 kDa molecular mass, and the gel electrophoretic mobility of purified RNase II corresponds to a polypeptide of approximately 70 kDa molecular mass (Zilh~o et aL, 1993, 1996; Coburn and Mackie, 1996). Transcription of the rnb gene is initiated at either of two promoters, and the 3' end of the mRNA is defined by a rho-independent terminator immediately downstream of the coding sequence (Zilh~o et al., 1996). RNase II has been implicated along with polynucleotide phosphorylase in mRNA decay (Section V,D), and perhaps also in tRNA 3' end maturation (Section V,B). RNase II- cells are viable (Piedade et al., 1995), but cells that lack both RNase II and polynucleotide phosphorylase activity are inviable (Donovan and Kushner, 1986). Discrete mRNA decay intermediates accumulate in the absence of these enzymes and RNase E (Arraiano et al., 1988). RNase II action is inhibited by RNA secondary structure, such as hairpins, and dissociates from its substrates in vitro approximately 10 nucleotides from such structures (Coburn and Mackie,
20
Allen W. Nichoison
1996). These products are resistant to further digestion by RNase II or by polynucleotide phosphorylase. Thus, in addition to a degradative role, under certain circumstances RNase II may protect RNA sequences from further degradation (Hajnsdort et al., 1994b; Pepe et al., 1994; Coburn and Mackie, 1996) (see also below). It remains to be shown whether RNase II associates with other proteins in vivo.
B. Polynucleotide Phosphorylase Polynucleotide phosphorylase (PNPase) is distinct from the other exoribonucleases in that it utilizes inorganic phosphate to carry out phosphorolytic cleavage of RNA, creating 5'-ribonucleoside diphosphates (Littauer and Soreq, 1982). It is noteworthy that, in contrast to a hydrolytic degradation reaction, the PNPase-catalyzed reaction conserves free energy in the 5'-rNDP products, which may be important to the cell under energy-poor conditions (Deutscher, 1993b). PNPase efficiently and processively degrades unstructured RNAs, including ribohomopolymers, but can be impeded by RNA secondary structure (Guarneros and Portier, 1991; Causton et al., 1994). PNPase also catalyzes the polymerization of 5'-rNDPs, forming RNA chains with release of phosphate, and can catalyze inorganic phosphate exchange with 5'-rNDPs (Littauer and Soreq, 1982). PNPase possesses an c~3 subunit structure (Littauer and Soreq, 1982). The a subunit is encoded by the p n p gene, mapping at 69 minutes, and is cotranscribed with the gene ( r p s O ) encoding ribosomal protein S15 (Regnier et al., 1987). At the carboxyl terminus, the c~ subunit contains a 69-amino-acid sequence that is similar to a repetitive sequence element in ribosomal protein S1, which also binds RNA (Regnier et al., 1987). Perhaps the two proteins share the same RNA recognition mechanism. Biochemical and genetic studies indicate that the RNA binding and catalytic functions of the a subunit are separable (Littauer and Soreq, 1982). An c~3/32 form of PNPase can be isolated from cells (Littauer and Soreq, 1982). The/3 subunit does not have an assigned function, but may confer regulation or substrate specificity. It has not yet been determined whether the two forms of PNPase are functionally distinct. PNPase is involved in mRNA decay and is a component of a macromolecular complex including RNase E (Section V,D). PNPase autoregulates trans-
1
Escherichia coli Ribonucleases
21
lation of its message, in conjunction with the action of RNase III and RNase E (Hajnsdorf et al., 1994a; Robert-Le Meur and Portier, 1994). PNPase-like activities are widely distributed in prokaryotic and eukaryotic organisms (Littauer and Soreq, 1982), and may therefore have a conserved biological role.
C. Ribonuclease D
RNase D was originally identified as an activity that degrades denatured RNA in vitro, and was subsequently shown to participate in 3' end maturation of tRNA (Deutscher, 1995). RNase D is specific for tRNA substrates, because it lacks detectable activity toward homopolymers. The enzyme is encoded by the rnd gene, mapping at 40 minutes (Zhang and Deutscher, 1988a,b). RNase D is a protein of 375 amino acids with a predicted molecular mass of 42.7 kDa and an isoelectric point of 6.2. The protein is active as a monomer and hydrolytically removes 5'-rNMPs from tRNA precursor 3' termini in a random fashion. RNase D requires a divalent metal ion (Mg 2§ Mn 2§ Co 2§ for activity (Cudny et al., 1981a). Because RNase D is not essential for cell growth, other enzymatic activities can effectively substitute (Blouin et al., 1983). However, elevated expression of RNase D can inhibit cell growth, which may suggest a deleterious action of RNase D on otherwise unreactive substrates (Zhang and Deutscher, 1988b).
D. Ribonuclease B N
RNase BN was uncovered in studies of the maturation of phage T4encoded tRNAs (reviewed in Deutscher, 1995). RNase BN is encoded by the rbn gene, at approximately 88 minutes. The enzyme has been partially purified and exhibits a molecular mass of 60 kDa. The enzyme is most active in vitro at acidic pH, and in the presence of Co 2§ (Deutscher, 1995) RNase BN participates in the maturation tRNA precursor 3' ends. However, it appears to be less efficient than the other exoribonucleases in this process (Deutscher, 1995).
22
Allen W. Nicholson
E. Ribonuclease T
RNase T participates in the 3' end maturation of tRNA (Deutscher et al., 1985) and has been shown to mature the 3' end of 5S rRNA (Li and Deutscher, 1995). RNase T catalyzes 3' end turnover of tRNA, which is defined by the removal and readdition of the 3'-terminal nucleotide. Thus, RNase T removes the 3'-terminal adenosine of an uncharged tRNA, whereas the 3' end is regenerated by tRNA nucleotidyltransferase (Deutscher et al., 1985). The significance of this event is unknown; however, RNase T- strains grow more slowly, suggesting an importance of RNase T-dependent reactions (Li and Deutscher, 1995). RNase T is encoded by the rnt gene, at 36 minutes, and encodes a polypeptide of approximately 23.5 kDa molecular mass (Huang and Deutscher, 1992). The enzyme is active as an c~2 dimer, and a cysteine residue (C168) is important for enzyme dimerization and activity (Li et al., 1996a,b).
F. Ribonuclease PH
RNase PH is a phosphorolytic enzyme that participates in tRNA 3' end maturation (reviewed by Deutscher, 1995). RNase PH utilizes inorganic phosphate to cleave substrate, yielding ribonucleoside 5'-diphosphate products. RNase PH is encoded by the rph gene, at 81.7 minutes, which encodes a 238-amino-acid protein with a predicted molecular mass of 25.5 kDa (Deutscher, 1995). Because the rph gene is dispensible, RNase PH is nonessential. RNase PH aggregates in solution, but the smallest active form is a 50-kDa or2 dimer (Deutscher, 1995). Like PNPase, RNase PH can act as an RNA synthetic enzyme as well as a nuclease. However, the low intracellular phosphate concentration would favor the degradative activity. Cells containing only RNase PH as the single representative of the five exoribonucleases are viable. However, mutants that lack both RNase PH and PNPase grow poorly, suggesting that the inorganic phosphate-utilization activity of these enzymes is important (Deutscher, 1995).
G. Oligoribonuclease
Oligoribonuclease (oligoRNase) was first characterized in 1975 (Niyogi and Datta, 1975; Datta and Niyogi, 1975). However, it was not known
1
Escherichia coli Ribonucleases
23
whether this activity is distinct, or a manifestation of another exoribonuclease. A new study confirmed oligoRNase as a distinct enzyme (Yu and Deutscher, 1995). OligoRNase has an apparent molecular mass of 38 kDa (Niyogi and Datta, 1975), and acts in a hydrolytic fashion. OligoRNase prefers short oligonucleotides as substrates, with the cleavage rate inversely proportional to RNA chain length (Datta and Niyogi, 1975). It was originally suggested that oligoRNase degrades the products of other ribonucleases, including RNase II and PNPase (Datta and Niyogi, 1975). Interestingly, oligoRNase was recently shown to copurify with PNPase (Yu and Deutscher, 1995). If the association with PNPase is specific, then oligoRNase may be an integral component of the mRNA degradation complex (Section V,D).
H. Ribonuclease R
RNase R was first detected as a residual exoribonuclease activity in extracts of cells deficient in the other exoribonucleases (Deutscher, 1993b). It should be noted that the designated name is identical to that given to an endoribonucleolytic activity (Table I) (Deutscher, 1993b). The apparent size of the enzyme is approximately 80 kDa, but the putative gene remains to be mapped (Deutscher, 1993a). In vitro substrates for RNase R include ribosomal RNA, which is degraded more efficiently than homopolymers, and mRNA (Kasai et al., 1977; Deutscher, 1993a). A degradative function for this activity is a possibility, but confirmation of this role awaits further study.
V. RIBONUCLEASE FUNCTIONAL ROLES A. Ribonucleases Involved in Ribosomal R N A Maturation
The E. coli ribosome contains three RNA species" 16S rRNA, 23S rRNA, and 5S rRNA. The rRNAs are cotranscribed in the listed order from seven rrn operons. An additional feature of the rrn transcription units is the presence of tRNA sequences between the 16S and 23S rRNAs, and occasionally following the 23S sequence, rRNA processing (and ribosomal subunit assembly) occurs during transcription, thus the
24
Allen W. Nichoison
primary transcript (30S R N A , approximately 5500 nucleotides) is not normally observed (King et al., 1986; Srivastava and Schlessinger, 1990). A n u m b e r of exo- and endoribonucleases participate in r R N A m a t u r a tion, several of which are well characterized. T h e enzymes recognize specific features in the transcript and catalyze specific cleavage. The R N a s e s do not act on naked r R N A , but on nascent ribosomal particles containing growing amounts of the ribosomal proteins. In fact, the final r R N A m a t u r a t i o n steps occur on actively translating ribosomes (Srivastava and Schlessinger, 1990). Figure 3 provides a diagram that indicates the sites of enzymatic cleavage in the primary transcript. Ribonuclease III carries out specific cleavage of sequences, which flank the 16S and 23S r R N A s , releasing the immediate precursors to 16S and 23S r R N A . The R N a s e III processing signals for 16S r R N A are f o r m e d
( 16S/RNA~ 10~ 7, 8"~ 3 ...~:
',,-4
~11
9
~9
tit
Spacer tRNA (~
5S RNA Distal tRNA
Fig. 3. Overall secondary structure scheme of the primary transcription product RNA elements that flank the 16S, 23S, and 5S ribosomal RNA sequences (Srivastava and Schlessinger, 1990). Numbers indicate the cleavage events that produce the mature 5' and 3' ends of 16S (1 and 2), 23S (10 and 11), and 5S (12) rRNAs. RNase III cleavage sites are indicated by 3, 4, 7, 8, and 9. RNase P cleavage sites are indicated by 5, and the cleavage(s) creating the mature 3' end of tRNA is indicated by 6 (see text for further details). Asterisks (single, double, triple) represent sequences required for 16S and 23S rRNA formation. Reprinted with permission from Annual Review of Microbiology, volume 44. 1990, by Annual Reviews, Inc.
1
Escherichia coli Ribonucleases
25
by complementary base pairing between the 5' and 3' flanking sequences, creating a dsRNA element (King et al., 1986; Srivastava and Schlessinger, 1990; Court, 1993). Coordinate double cleavage releases a 17S rRNA species in which the mature 5' and 3' ends of 16S rRNA are created by separate endonucleolytic events. The corresponding enzymatic activities ("maturases") have been detected, but not characterized (Hayes and Vasseur, 1976; Dahlberg et al., 1978; Srivastava and Schlessinger, 1989). A similar dsRNA structure can be formed between sequences flanking the 23S rRNA. Moreover, this structure in purified form can be accurately processed by RNase III in vitro (Robertson, 1982; Court, 1993; King et al., 1986). However, the proposed dsRNA element does not appear to participate in the in vivo pathway. Specifically, RNase III cleavage of the 5' flanking sequence occurs prior to the synthesis of the 3' flanking sequence (King and Schlessinger, 1983). It was suggested that the 5' flanking segment participates in an alternate intramolecular (or perhaps intermolecular) structure that is cleaved by RNase III (King and Schlessinger, 1983). It would follow that the 3' flanking sequence also forms a separate processing signal for RNase III. RNase III cleavage provides the immediate precursor to 23S rRNA, from which the mature 5' end is fashioned by an endonucleolytic activity (King et al., 1984; Srivastava and Schlessinger, 1988) and the 3' end is created by an exonucleolytic activity (Sirdeshmukh and Schlessinger, 1985a,b). Both of these activities remain to be characterized, but appear to use polysomal ribosomes as substrates (Srivastava and Schlessinger, 1990). The precursor of 5S rRNA is 9S RNA, the 5' end of which is created by RNase III cleavage downstream of the 23S rRNA sequence; the 3' end is either the transcription terminator or is created by RNase P cleavage of the trailer tRNA sequence, when present (Fig. 3). 9S RNA is a substrate for RNase E, which creates the immediate precursor to 5S rRNA, with three extra nucleotides at the 5' and 3' ends (King et al., 1986; Srivastava and Schlessinger, 1990). The 5S rRNA 3' end is created through the action of RNase T (Li and Deutscher, 1995). Because RNase T- strains are viable, 5S rRNA 3' end maturation is not required for ribosome function (Li and Deutscher, 1995). The 5' end maturation activity has not been characterized. However, it is distinct from RNase T, and may be a (long sought after!) 5' ~ 3'-exoribonuclease (Li and Deutscher, 1995). Are the various steps involved in rRNA maturation precisely coordinated, or otherwise occur in an obligatory fashion? Because RNase IIIstrains are viable, RNase III cleavage is not required to create functional
26
Allen W. Nicholson
ribosomes (King et al., 1986; Srivastava and Schlessinger, 1990). However, in the absence of RNase III, r R N A processing slows down, causing accumulation of the 30S rRNA precursor, which can be seen as a distinct species by gel electrophoresis (King et al., 1986; Srivastava and Schlessinger, 1990). Correct maturation of 16S rRNA still occurs; thus the endonucleolytic cleavages that create the 16S 5' and 3' ends can act independently of RNase III. However, the mature 5' and 3' ends of 23S rRNA are not formed, and 50S subunits that are assembled contain a 23S rRNA carrying 20-100 extra nucleotides at its 5' and 3' ends (Sirdeshmukh and Schlessinger, 1985a,b). One explanation for the slower growth of RNase III- cells is that the 50S subunits containing altered 23S rRNA exhibit suboptimal protein synthetic function (King et al., 1984).
B. Ribonucleases Involved in Transfer R N A Maturation Escherichia coli contains about 60 different tRNA species, which collectively constitute about 14% of the cellular RNA (Neidhardt et al., 1990; Komire et al., 1990). The tRNAs are generated from a large number of transcription units of differing genetic structures. The mature tRNAs are fashioned from the precursor transcripts by the action of specific endo- and exonucleases. Several phages encode tRNA genes, whose expression requires specific processing. The coliphage T4 encodes a tRNA gene cluster, where eight tRNA species are cotranscribed and processed (Mazzara et al., 1981). Studies on the expression of the T4 tRNA gene cluster have provided perhaps the best understood model for tRNA maturation (reviewed by Schmidt and Apirion, 1984). A recent comprehensive review of tRNA processing ribonucleases is provided by Deutscher (1995). Some E. coli tRNAs are synthesized in monocistronic form; other species are cotranscribed with other tRNAs, rRNAs, or protein coding sequences (Deutscher, 1995). For the multicistronic transcripts, endonucleolytic cleavage is absolutely required before commencement of further reactions that are specific to the maturation of the excised pretRNA (Deutscher, 1995). A consensus scheme for tRNA maturation is provided in Fig. 4. In all cases, the mature 5' end is created by the action of RNase P, and the 3' end is created by a series of exonucleolytic trimming reactions, involving several enzymes. The clipped off 5' leader sequence is presumably degraded, unless it itself contains a tRNA,
1
27
Escherichia coli Ribonucleases
0
3
1
0
Fig. 4. Summary diagram representing the processing pathway for E. coli tRNA precursors (Deutscher, 1995). Endonuclease cleavage reactions are indicated by vertical arrows. Numbers represent the overall order of the reactions. Exonucleolytic reactions are indicated by horizontal arrows. Points 0 and 1 represent cleavages that excise the tRNA precursor from a larger transcript. Point 3 represents RNase P processing; 2 and 4 represent steps most efficiently performed by RNase PH and RNase T, respectively (see text for further details). Reprinted with permission from the American Society for Microbiology.
rRNA, or coding sequence. RNase P recognizes specific aspects of the tRNA structure embedded within the variety of precursors, and the protein subunit of RNase P increases the versatility of RNase P in handling the many different tRNA precursors (Reich et al., 1988; Altman, 1990). The processing events that occur at the tRNA 5' end are not necessarily independent of events at the 3' end. Thus, efficient RNase P action may require a substrate that has been subjected to prior 3'end-specific processing (Deutscher, 1995). The tRNA 3' end is created by exonucleolytic removal of 3' trailer sequences. This presents a formally simple process from the standpoint of reaction type, but is actually much more elaborate due to the number of enzymatic activities involved. At least five exonucleases have been implicated: RNases II, D, BN, T, and PH. Mutant cells lacking four of the RNases (II, D, BN, and T) still grow well, and specific tRNAs can be matured. However, the additional loss of RNase PH is lethal. The presence of any one of these RNases alone can support cell growth, with the order of ribonuclease effectiveness being T > PH > D > II > BN (Kelly and Deutscher, 1992). In vivo and in vitro studies on model tRNA substrates have provided a preliminary choreography for the exonuclease activities (Fig. 4) (Deutscher, 1995). RNase II and/or PNPase provide rapid initial shortening of long 3' trailer sequences, yielding pre-tRNAs with two to four additional 3' nucleotides. Additional shortening to a + 1 species is most effectively accomplished by RNase PH, with RNase T catalyzing removal of the final nucleotide. The final two steps probably require prior RNase P cleavage. Why is there a need for multiple exoribonucleases for tRNA 3' end maturation? Although the RNases exhibit distinct preferences in vitro, they can substitute for each other in vivo.
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Given the tremendous energy investment in and importance of protein synthesis, tRNAs need to mature rapidly. The most efficient maturation of tRNA 3' ends may only be provided by this route.
C. Ribonucleases Involved in Messenger R N A Maturation and Turnover
The enzymes and RNA elements involved in mRNA decay are only now beginning to yield to experimental analysis. A brief discussion of the general features of bacterial mRNA decay and the associated enzymes is provided herein. The reader is also referred to several reviews (Ehretsmann et al., 1992a; Petersen, 1992; Nierlich, 1995) and a comprehensive monograph (Belasco and Brawerman, 1993).
1. Messenger RNA Maturation Pathways A number of E. coli and coliphage mRNAs undergo maturation prior to function. RNase III is involved in mRNA maturation [reviewed by Court (1993) and Nicholson (1996)]. The phage T7 polycistronic early mRNA precursor (approximately 7000 nucleotides) is cleaved at five sites by RNase III, generating the mature mono- and dicistronic mRNAs (Dunn and Studier, 1983). Several of the T7 late transcripts also contain RNase III cleavage sites (Dunn and Studier, 1983). RNase III processing influences the translational activities of several of the T7 messages, probably by influencing the accessibility of the Shine-Dalgarno sequence to the ribosome. However, because T7 can grow on RNase III- strains, T7 mRNA activity does not require cleavage. RNase III carries out maturation cleavage of several cellular mRNAs, but the functional importance of these cleavages is not known (Nicholson, 1996). It was recently shown that RNase III cleavage within the coding sequence of a plasmid transcript can down-regulate expression of the encoded protein (Koraimann et al., 1993). RNase III processing can provide the rate-limiting step in mRNA decay. RNase III action can remove or otherwise alter secondary structures (e.g., hairpins) from the 3 ' or 5' end of mRNAs, thereby allowing further rapid degradation by other enzymes [reviewed by Court (1993) and Nicholson (1996)]. Inhibition of phage lambda int (integrase) gene expression is provided by RNase III cleavage downstream of the int gene
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in the PL transcript. Cleavage removes a hairpin, allowing subsequent exonucleolytic digestion into the int gene by RNase II and/or PNPase. RNase III cleavage in the 5' untranslated region (UTR) of its own message accelerates subsequent decay of the message, providing negative autoregulation (Bardwell et aL, 1989). RNase III cleavage within the 5' UTR of the p n p - r p s O mRNA (encoding PNPase and ribosomal protein $15) also allows subsequent RNase E cleavage and message decay (Portier et al., 1987).
2. Messenger RNA Decay Pathways Mature E. coli mRNAs are naturally resistant to exonucleolytic action. Because there are apparently no 5' ~ 3'-exoribonucleases, mRNAs are not normally exposed to such attack at their 5' termini, mRNAs usually contain a hairpin or related secondary structure at their 3' ends, which provides an impediment to the 3' exonucleolytic action of RNase II and PNPase. As a consequence, initiation of bacterial mRNA decay is usually established by endonucleolytic cleavage, which may be the rate-limiting step. RNase E and RNase III have been shown to provide this event, but other endonucleases may also participate (e.g., see Arraiano et al., 1993). Thus, cleavage upstream from a 3'-terminal hairpin can allow access of exonucleases to the body of the mRNA (Higgins et al., 1993). Other primary endonucleolytic cleavage sites are near the mRNA 5' end (Bechhofer, 1993). In this case, primary cleavage is followed by a 5' ~ 3' propagated "wave" of endonucleolytic cleavages, providing short fragments with 3' ends accessible to exonucleolytic action. Current evidence suggests that the pathway of decay of the majority of E. coli mRNAs exhibits 5' ~ 3' directionality (Hansen et al., 1994). There is no strong evidence that random endonucleolytic cleavage within the coding sequence provides the rate-limiting step. If this were so, then longer mRNAs would have shorter half-lives, and this is not uniformly observed (Belasco, 1993). Any scheme for mRNA degradation must take into account the fact that bacterial transcription and translation are coupled events. Endonucleolytic cleavage sites can be occluded by translating ribosomes, and mRNA decay rates can be influenced by the frequency of translation initiation (reviewed by Petersen, 1993). In the situation where transcription and translation are uncoupled, mRNA synthesis may be largely completed before appreciable translation occurs. In this situation the mRNA may be substantially less stable, because it would be fully exposed
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to degradative endonucleases (e.g., see lost and Dreyfus, 1995). Cleavage within the 5' UTR, as well as the coding sequence, can functionally inactivate the mRNA and accelerate degradation. Cleavage can inhibit new rounds of translation initiation and expose downstream sequences to nuclease attack following transit of the last ribosome (Petersen, 1993). Several E. coli mRNAs with prolonged physical and functional lifetimes exhibit at their 5' ends specific RNA structures that confer resistance to degradation. The o m p A mRNA has a 5' hairpin structure that impedes RNase E cleavage near the 5' end (Emory and Belasco, 1990; Emory et al., 1992; Hansen et aL, 1994). Appending a 5' single-strand extension to the 5' hairpin abrogates the ability of the hairpin to block RNase E cleavage (Emory et al., 1992). Statistical analysis indicates that the initiation of RNase E-dependent degradation of a majority of E. coli mRNAs occurs near the 5' end, and that this pathway appears to require a 5' end in single-stranded form (Hansen et al., 1994). 3. R o l e o f 3 ' - E n d Polyadenylylation in R N A Degradation Escherichia coli contains a poly(A) polymerase activity (PAP I), which catalyzes the template-independent addition of adenylate residues onto RNA 3' ends (Deutscher, 1978; Cao and Sarkar, 1992a,b; He et al., 1993; Xu et al., 1993). PAP I is active in vivo, in that mRNAs can be isolated with 3' poly(A) tails (Cao and Sarkar, 1992a). PAP I is a monomeric enzyme with a molecular mass of approximately 55 kDa (Cao and Sarkar, 1992b) and is encoded by the pcnB gene, mapping at 3 minutes (Lopilato et al., 1986). PAP I- mutants are viable, although they grow more slowly (Liu and Parkinson, 1989). A second poly(A) polymerase activity (PAP II) has been characterized and exhibits a molecular mass of 35 kDa (Kalapos et al., 1994). It preferentially utilizes poly(A) as a substrate and is more thermolabile than PAP I (Kalapos et al., 1994). The viability of a PAP I-, PAP II- double mutant has not been determined. Studies on the degradation of the ColE1 plasmid-encoded transcript have shown that 3' polyadenylylation is a determinant of RNA stability (Xu et al., 1993; He et al., 1993). RNA I, an antisense repressor of ColE1 plasmid replication, has a short-life, comparable to that of mRNA. RNA I undergoes site-specific cleavage by RNase E, which provides the ratelimiting step for RNA I decay. The subsequent rapid breakdown of the RNase E cleavage products requires the action of PAP I and PNPase (Xu and Cohen, 1995). Specifically, polyadenylylation accelerates PNPasedependent decay. The binding of PNPase to a poly(A) tail over a certain
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length allows efficient exonucleolytic digestion into the body of the RNA, which possesses a secondary structure that, in the absence of a poly(A) tail, would impede PNPase digestion (Xu and Cohen, 1995). RNase II may also remove 3' poly(A) tails, but is also sensitive to RNA secondary structure (Coburn and Mackie, 1996), and cannot compensate for the loss of PNPase (Xu and Cohen, 1995). Interestingly, the absence of PNPase enhances RNA I stability by inhibiting initial cleavage by RNase E (Xu and Cohen, 1995). This and other evidence (see section V,C,4) indicate that RNase E and PNPase functionally interact. Because E. coli mRNAs can be shown to possess 3' poly(A) tails, it was anticipated that the rapid decay of mRNA may reflect the relative efficiency of the 3'polyadenylylation/PNPase digestion pathway. Recent evidence implicates this pathway in mRNA decay (O'Hara et al., 1995; Hajnsdorf et al., 1995).
4. Macromolecular Complex Mediating Messenger RNA Degradation Gentle fractionation of E. coli extracts in the presence of protease inhibitors affords the purification of a complex that includes RNase E and PNPase, as well as several other polypeptides (Carpousis et al., 1994; Py et al., 1994). Because this complex can degrade RNA both by endoand exonucleolytic cleavage, it contains the minimal activities necessary for mRNA decay (Carpousis et al., 1994; Py et al., 1994). Moreover, because RNase E can cleave near the 5' end of mRNA, and PNPase recognizes RNA 3' ends, the association of these two enzymes in a single complex implies a functional association of the 5' and 3' ends of bacterial RNA sequences undergoing degradation (Py et al., 1994). PNPase also copurifies with oligoRNase (Yu and Deutscher, 1995). The latter activity may complete the breakdown of the small oligonucleotide products of PNPase action. The identities of the other polypeptides remain to be determined, and may include other activities that alter RNA structure, or localize the complex and regulate its activity. An energy (ATP) requirement for this complex would derive, at the minimum, from the formation of poly(A) tails. Part of the free-energy expenditure would be recovered from the action of PNPase. An additional energy requirement for RNA degradation remains a possibility. The discovery of 3' poly(A) tails as an important determinant of bacterial RNA stability and the identification of an RNA degradative
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Allen W. Nicholson
complex provide an initial model for m R N A decay (Fig. 5) ( O ' H a r a et aL, 1995). In one pathway, which involves net 3' ~ 5' decay, the formation of the 3' poly(A) tail by PAP I allows "loading" of PNPase in the R N A degradation complex. PNPase shortening of the poly(A) tail is accompanied by upstream endonucleolytic cleavage by RNase E, generating a new 3' end, which can be polyadenylated, and the cycle repeated. Alternatively, because it is known that R N A degradation can still proceed in the absence of RNase E and PNPase, a separate pathway can be proposed (involving an unidentified endonuclease, "RNase X " ) (Fig. 5). The 5' 3' decay of R N A would require a modification of the 3' --~ 5' decay
Fig. 5. Schemefor mRNA decay in E. coli (O'Hara et al., 1995). Enzymes and enzyme cleavage sites are indicated; PBP, poly(A) binding protein. Shaded boxes indicate regions in mRNA that influence RNA stability. Initial cleavage products are further degraded by RNase II, PNPase, and perhaps oligoRNase (see text). Reprinted with permission from the National Academy oI Science.
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model. In this pathway, the complex would have to remain associated with the upstream cleavage product, which is fated for PAP I/PNPasedependent degradation, while simultaneously binding to the downstream RNA, prior to recognition of the next accessible RNase E cleavage site.
VI. O T H E R R I B O N U C L E A S E S
A. Ribonuclease Activity Associated with RNA Polymerase Premature arrest of elongation by E. coli RNA polymerase is countered by the action of the GreA and GreB transcription factors (Borukhov et aL, 1992, 1993). Suppression of early arrest is also associated with specific endonucleolytic cleavage near the 3' end of the nascent transcript. The mechanism of RNA cleavage in the arrested transcription complex is apparently conserved, because it is also seen with eukaryotic and viral RNA polymerases. Specifically, the mammalian transcription factor SII suppresses early arrest of transcription by cleavage upstream from the stalled RNA 3' end (Reines, 1992). Transcript cleavage is promoted by the interaction of GreA or GreB protein with the RNA polymerase. GreA and GreB induce cleavage either 2-3 or approximately 9 nucleotides, respectively, from the stalled mRNA 3' end (Borukhov et al., 1992, 1993). The newly formed RNA 3' end provides a site for renewed transcription elongation, which proceeds through the arrest site. GreA and GreB proteins are the same size (158 amino acids) and are closely similar in sequence (Borukhov et al., 1993). GreA protein has been crystallized and its structure has been determined (Stebbins et al., 19995). GreA and GreB are not ribonucleases. Instead, the two factors may cause a conformational change in the transcription complex, inducing RNA cleavage. The action of GreA and GreB can be mimicked by raising the pH, and the pH dependence has an inflection at approximately 8.6, presumably due to ionizable protein side chain(s) (Orlova et al., 1995). One possible mechanism is that the GreA and GreB proteins induce "strain" at specific sites in the nascent transcript, resulting in cleavage. Alternatively, a latent ribonuclease in RNA polymerase may be activated by GreA or GreB binding, which can be mimicked by elevated pH (Orlova et al., 1995). It is not known exactly how cleavage occurs, but for the mammalian cleavage/antiarrest factor SII,
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Allen W. Nicholson
pyrophosphorylysis (i.e., the reverse of polymerization) occurs ( R u d d e t al., 1994). Whatever the precise mechanism, cleavage must provide a free 3'-hydroxyl group to allow chain elongation.
B. Prr Anticodon Nuclease A n E. coli clinical isolate, CT196, contains a genetic element, prr, at 29 minutes. The prr element includes the prrC gene, which encodes a ribonuclease (termed the anticodon nuclease, or ACNase) specific for the anticodon loop of the cell-encoded tRNA Lys3.T4 infection specifically activates the ACNase, causing cleavage of tRNA Lys3.The latent ACNase is unmasked by expression of the T4 Stp polypeptide, which is 26 amino acids in length. However, a translational block to T4 phage development occurs only when the T4 fails to express its polynucleotide kinase and/ or RNA ligase activities, which can repair the cleaved tRNA Lys3.Successful repair can sustain protein synthesis and permit T4 development (for a recent review, see Snyder, 1995). The ACNase is a component of the Type Ic D N A restrictionmodification system, EcoprrI (Levitz et al., 1990; Tyndall et al., 1994). The ACNase protein interacts with the restriction/modification proteins, which masks its activity. The T4 Stp polypeptide primarily inhibits the action of the restriction-modification system. However, in doing so, it releases the active form of the ACNase (Amitsur et al., 1992; Morad et al., 1993). The ACNase therefore may be regarded as a cellular selfdestruction agent, which along the restriction system provides an effective antiphage System (Snyder, 1995). The phage polynucleotide kinase and RNA ligase activities represent the T4 solution to inactivating the otherwise lethal translational block.
C. Coliphage T4 RegB Nuclease Phage T4 infection of E. coli induces an activity that introduces cuts within the ribosome binding sites of specific T4 mRNAs (Uzan et al., 1988). Cleavage occurs 3' to the second G residue in the sequence G G A G , which represents the core of the Shine-Dalgarno (S-D) sequence that directs m R N A binding to the 30S ribosomal subunit. The
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nucleolytic activity copurifies with the product of the T4 regB gene, which encodes a protein of 18 kDa, and is termed the RegB nuclease (Ruckman et al., 1994). The RegB nuclease is important for the temporal down-regulation of specific T4 early protein expression. RegBdependent cleavage of the S-D sequence within the T4 mRNAs would directly inactivate their translation. However, additional factors are involved, because other mRNAs with GGAG-containing S-D sequences are not cleaved. Because the regB mRNA is a target for the RegB nuclease, the enzyme autoregulates its own synthesis (Ruckman et al., 1989). There is evidence that the RegB nuclease associates with the 30S ribosomal subunit. First, the RegB nuclease preferentially cleaves G G A G sequences specifically associated with translation initiation regions (Sanson and Uzan, 1993). Second, there is also a strong correlation between the strength of a translation initiation region and the susceptibility of the associated S-D sequence to RegB nuclease cleavage (Sanson and Uzan, 1993; Ruckman et al., 1994). Third, the activity of RegB nuclease is significantly enhanced by S1, a 30S ribosomal protein involved in translation initiation (Ruckman et al., 1994). The association of a phage-encoded RNase with the small ribosomal subunit would provide an efficient means to cleave selectively actively translated mRNAs. However, RegB cleavage can also occur at other sites in mRNA, which may play a role in the degradation of the mRNAs (Sanson and Uzan, 1993).
VII. R I B O N U C L E A S E R E G U L A T I O N A N D R N A
METABOLIC CONTROL The bacterial cell adapts to a changing environment by altering its pattern of gene expression. The regulation of bacterial gene expression at the transcriptional and translational levels is well established (Neid hardt et al., 1987). However, gene expression is also controlled by mRNA decay rates. Bacterial mRNAs exhibit upward of a 50-fold difference in decay rates, ranging from ---0.5 to >20 minutes, with an average halflife of 2-4 minutes (Belasco, 1993). The rate of decay establishes the steady-state mRNA concentration and therefore the rate of synthesis of the encoded protein. The instability of mRNA permits rapid change in gene expression patterns, allowing the cell to adjust quickly to new growth conditions (Belasco, 1993). On the other hand, there are situa-
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tions where RNA stability is required (e.g., see Thisted et al., 1994). Perhaps the extreme case of RNA stability is exhibited by RNA phage chromosomes, which not only must be translated, but also packaged in undamaged form in the progeny viral particles (Zinder, 1975). It has been shown that a protein complex involving heat-shock protein GroEL binds to and confers physical stability to RNA (Georgellis et al., 1995). The interaction is reversible, as the presence of ATP and Mg 2+ releases the protection (Georgellis et al., 1995). The RNA-binding/protection activity is enhanced in cell growth conditions known to stabilize specific mRNAs, and under these conditions the GroEL protein is in modified form. The covalent modification of GroEL [possibly phosphorylation; see Sherman and Goldberg (1992)] may provide a rapid response to altered growth conditions and provide a mechanism for the observed o m p A mRNA stability and its control by growth rate shifts (Georgelis et al., 1995). However, a global control mechanism involving chaperonin function has not been uncovered. Thus, the heat-shock response, which includes increased expression of GroEL, does not cause noticeably different change in overall mRNA stability (Henry et aL, 1992). An earlier report described an interaction between GroEL and RNase E, detected by immunoprecipitation (Sohlberg et al., 1993). It is not clear whether this interaction is related to the RNA protection function mediated by GroEL, or whether RNase E requires the chaperonin function of GroEL for activity. Phage infection can alter RNA processing and degradative activities. T7 phage expresses a protein kinase, which induces the phosphorylation of a large number of proteins (Robertson et al., 1994), including RNase III (Mayer and Schweiger, 1983). Phosphorylation stimulates RNase III activity in vitro (Mayer and Schweiger, 1983), which may allow more efficient maturation of the abundant T7 mRNAs. T4 infection creates a modified RNase D, wherein a T4 infection-specific polypeptide associates with RNase D (Cudny et al., 1981b). The functional consequences of this modification are not understood. Finally, as discussed above, T4 also expresses the RegB nuclease, which cleaves and inactivates specific T4 early transcripts (see section VI,C). Glucose starvation increases bulk mRNA stability (Albertson and Nystrom, 1994). A primary source of the mRNA stabilization may be the slower translation rate, where ribosomes provide enhanced protection of mRNA (Albertson and Nystrom, 1994). On the other hand, starvation of E. coli leads to enhanced degradation of stable RNA (Apirion, 1975; Kaplan and Apirion, 1975a,b). Under these conditions, the cell may
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utilize stable RNA (approximately 95% percent of the total RNA) as a source of nucleotides. Little is known of this mechanism of r R N A turnover. However, RNase I (or perhaps RNase I*) is implicated in this process (Kaplan and Apirion, 1975a,b; Deutscher, 1993b). In another study, exposure of E. coli cells to mercuric (Hg 2+) ion induced degradation of rRNA (Beppu and Arima, 1969). Mercuric ion may inactivate an intracellular inhibitor of RNase I* (Beppu and Arima, 1969). RNase I* is also apparently involved in degradation of rRNA in cells expressing the F plasmid s r n B + gene (Ito and Ohnishi, 1983).
VIII. C O N C L U S I O N S A N D P R O S P E C T S
Contemporary studies suggest a broad phylogenetic conservation of RNA-processing enzymes and their functions. RNase P is a ubiquitous enzyme with a conserved role, and there is evidence for RNase III and RNase E-related activities in eukaryotic cells, perhaps also with functional roles similar to the E. coli enzymes. RNA splicing reactions, once thought to be confined to eukaryotic cells, are now being uncovered in bacterial systems with increasing frequency (Belfort et al., 1995). RNA 3'-end polyadenylylation, originally identified in eukaryotes, is now an established reaction in E. coli and other bacteria, and may be a universal determinant of RNA stability. Because polyadenylylation affects translational activity of eukaryotic mRNAs (Sachs, 1990), it would be interesting to determine whether it can influence the translation of bacterial mRNAs. The pathway of bacterial rRNA maturation has been considered to be quite different from that of eukaryotic rRNA. However, investigations on eukaryotic (Morrissey and Tollervey, 1995) and archaeal (Durovic and Dennis, 1994; Potter et al., 1995) rRNA processing are beginning to reveal an underlying formal conservation of the maturation pathway, and have prompted fresh speculation on the evolution of rRNA processing. Thus, although there is no evidence for small (nucleolar-like) RNA involvement in E. coli rRNA processing, there may be RNase Ill-related enzymes participating in eukaryotic rRNA maturation, which may recognize and cleave dsRNA elements, formed between snoRNAs (small nucleolar RNAs) and the processing sites in the primary rRNA transcript.
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New experimental directions now include structural and mechanistic studies of the E. coli ribonucleases. Determining how substrate is recognized and phosphodiester bonds are cleaved will broaden our understanding of RNA-processing mechanisms. In this regard, RNase HI provides an important paradigm in how a R N A phosphodiesterase works at the atomic level. The role of R N A structure in determining processing reactivity is now being experimentally addressed. Multidimensional NMR analysis of processing substrates (e.g., see Schweisguth et al., 1994) will shed light on how R N A structure determines processing reactivity. Another primary focus is the structure of the m R N A degradation complex, what the resident activities are, and how they interact to carry out this essential process. RNase III and RNase HI have served as valuable reagents for probing RNA structure and function. It is anticipated that bacterial RNases and specific enzymatic derivatives may provide therapeutic advantage in fighting disease. Thus, RNase P is being exploited as an endogenous activity for the targeted cleavage of RNA and inactivation of gene expression (Li et al., 1992; Frank et al., 1994; Liu and Altman, 1995). A potentially exciting application of the phage T4 ACNase is indicated by the observation that the primer for human immunodeficiency virus reverse transcriptase, tRNA Lys3, can be cleaved by the ACNase (Snyder, 1995). The ACNase nuclease may provide therapeutic promise by selectively inactivating HIV-1 replication. Clearly, the full biotechnological potential of the E. coli ribonucleases remain to be fully explored and exploited.
ACKNOWLEDGMENTS
The author greatly appreciates the reprints and information provided by colleagues. Research on RNase III in the author's laboratory is supported by the National Institutes of Health (GM41283).
REFERENCES
Albertson, N., and Nystrom, T. (1994). Effects of starvation for exogenous carbon on functional mRNA stability and rate of peptide chain elongation in Escherichia coli. FEMS MicrobioL Lett. 117, 181-188.
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Alifano, Y., Kivellini, F., Piscitelli, C., Arraiano, C., Bruni, C. B., and Carlomagno, M. S. (1994). Ribonuclease E provides substrates for ribonuclease P-dependent processing of a polycistronic mRNA. Genes Dev. fl, 3021-3031. Altman, S. (1990). Ribonuclease P. J. Biol. Chem. 265, 20053"20056. Amitsur, M., Morad, I., Chapman-Shimshoni, D., and Kaufmann, G. (1992). HSD restriction-modification proteins partake in latent anticodon nuclease. E M B O J. 11, 31293134. Apirion, D. (1975). The fate of mRNA and rRNA in Escherichia coli. In "Processing of RNA" (J. J. Dunn, ed.), Brookhaven Symposium in Biology, No. 26, pp. 286-306. U.S. Department of Commerce, Springfield, VA. Arraiano, C. M., Yancey, S. D., and Kushner, S. R. (1988). Stabilization of discrete mRNA breakdown products in ams pnp rnb multiple mutants of Escherichia coli K-12. J. Bacteriol. 170, 4625-4633. Arraiano, C. M., Yancey, S. D., and Kushner, S. R. (1993). Identification of endonucleolytic cleavage sites involved in decay of Escherichia coli trxA mRNA. J. Bacteriol. 175,10431052. Babitzke, P., and Kushner, S. R. (1991). The Ams (altered mRNA stability) protein and ribonuclease E are encoded by the same structural gene of Escherichia coli. Proc. Natl. Acad. Sci. U.S.A. 88, 1-5. Bardwell, J. C. A., Regnier, P., Chen, S.-M., Nakamura, Y., Grunberg-Manago, M., and Court, D. L. (1989). Autoregulation of RNase III operon by mRNA processing. E M B O J. 88, 3401-3407. Bechhofer, D. (1993). 5' mRNA stabilizers. In "Control of Messenger RNA Stability" (J. G. Belasco and G. Brawerman, eds.), pp. 31-52. Academic Press, New York. Beese, L. S., and Steitz, T. A. (1991). Structural basis for the 3'-5' exonuclease activity of Escherichia coli DNA polymerase I: A two metal ion mechanism. E M B O J. 10, 25-33. Belasco, J. G. (1993). mRNA degradation in prokaryotic cells: An overview. In "Control of Messenger RNA Stability" (J. G. Belasco and G. Brawerman, eds.), pp. 3-12. Academic Press, New York. Belasco, J. G., and Brawerman, G. (1993). "Control of Messenger RNA Stability." Academic Press, New York. Belfort, M., Reaban, M. E. Coetzee, T., and Dalgaard, J. Z. (1995). Prokaryotic introns and inteins: A panoply of form and function. J. Bacteriol. 177, 3897-3903. Beppu, T., and Arima, K. (1969). Induction by mercuric ion of extensive degradation of cellular ribonucleic acid in Escherichia coli. J. Bacteriol. 98, 888-897. Blouin, R. T., Zaniewski, R., and Deutscher, M. P. (1983). Ribonuclease D is not essential for the normal growth of Escherichia coli or bacteriophage T4 or for the biosynthesis of a T4 suppressor tRNA. J. Biol. Chem. 258, 1423-1426. Borukhov, S., Polyakov, A., Nikiforov, V., and Goldfarb, A. (1992). GreA protein: A transcription elongation factor from Escherichia coli. Proc. Natl. Acad. Sci. U.S.A. 89, 8899-8902. Borukhov, S., Sagitov, V., and Goldfarb, A. (1993). Transcript cleavage factors from E. coll. Cell (Cambridge, Mass.) 72, 459-466. Brock, T. D. (1990). "The Emergence of Bacterial Genetics." Cold Spring Harbor Laboratory, Cold Spring Harbor, New York. Bycroft, M., Grunert, S., Murzin, A. G., Proctor, M., and St. Johnston, D. (1995). NMR solution structure of a dsRNA binding domain from Drosophila staufen protein reveals homology to the N-terminal domain of ribosomal protein $5. E M B O J. 14, 3563-3571.
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Piedade, J., Zilh~o, R., and Arraiano, C. M. (1995). Construction and characterization of an absolute deletion mutant of Escherichia coli ribonuclease II. FEMS Microbiol. Lett. 127, 187-194. Portier, C., Dondon, L., Grunberg-Manago, M., and Regnier, P. (1987). The first step in the functional inactivation of the Escherichia coli polynucleotide phosphorylase messenger is ribonuclease III processing at the 5' end. E M B O J. 6, 2165-2170. Potter, S., Durovic, P., and Dennis, P. P. (1995). Ribosomal RNA precursor processing by a eukaryotic U3 small nucleolar RNA-like molecule in an Archaeon. Science 268, 1056-1060. Py, B., Causton, H., Mudd, E. A., and Higgins,. C. F. (1994). A protein complex mediating mRNA degradation in Escherichia coli. Mol. Microbiol. 14, 717-729. Regnier, P., Grunberg-Manago, M., and Portier, C. (1987). Nucleotide sequence of the pnp gene of Escherichia coli encoding polynucleotide phosphorylase. J. Biol. Chem. 262, 63-68. Reich, C., Olsen, G. J., Pace, B., and Pace, N. R. (1988). Role of the protein moeity of ribonuclease P, a ribonucleoprotein enzyme. Science 239, 178-181. Reines, D. (1992). Elongation factor-dependent transcript shortening by template-engaged RNA polymerase II. J. Biol. Chem. 267, 3795-3800. Robertson, E. S., Aggison, L. A., and Nicholson, A. W. (1994). Phosphorylation of elongation factor G and ribosomal protein $6 in bacteriophage TT-infected Escherichia coll. Mol. Microbiol. 11, 1045-1057. Robert-Le Meur, M., and Portier, C. (1994). Polynucleotide phosphorylase of Escherichia coli induces the degradation of its RNase III processed messenger by preventing its translation. Nucleic Acids Res. 22, 397-403. Robertson, H. D. (1982). Escherichia coil ribonuclease III cleavage sites. Cell (Cambridge, Mass.) 30, 669-672. Robertson, H. D., Webster, R. E., and Zinder, N. D. (1968). Purification and properties of ribonuclease III from Escherichia coll. J. Biol. Chem. 243, 82-91. Robertson, H. D., Altman, S., and Smith, J. D. (1972). Purification and properties of a specific Escherichia coli ribonuclease which cleaves a tyrosine transfer ribonucleic acid precursor. J. Biol. Chem. 247, 5243-5251. Rotondo, G., Gillespie, M., and Frendewey, D. (1995). Rescue of the fission yeast snRNA synthesis mutant snml by overexpression of the double-strand-specific Pacl ribonuclease. Mol. Gen. Genet. 247, 698-708. Ruckman, J., Parma, D., Tuerck, C., Hall, D. H., and Gold, L. (1989). Identification of a T4 gene required for bacteriophage mRNA processing. New Biol. 1, 54-65. Ruckman, J., Ringquist, S., Brody, E., and Gold, L. (1994). The bacteriophage T4 regB ribonuclease. Stimulation of the purified enzyme by ribosomal protein S1. J. Biol. Chem. 269, 26655-26662. Rudd, M. D., Izban, M. G., and Luse, D. S. (1994). The active site of RNA polymerase II participates in transcript cleavage within arrested ternary complexes. Proc. Natl. Acad. Sci. U.S.A. 91, 8057-8061. Sachs, A. (1990). The role of poly(A) in the translation and stability of mRNA. Curr. Opin. Cell Biol. 2, 1092-1098. Sanson, B., and Uzan, M. (1993). Dual role of the sequence-specific bacteriophage T4 endoribonuclease RegB. J. Mol. Biol. 233, 429-446. Schmidt, F. J., and Apirion, D. (1984). T4 transfer RNAs: Paradigmatic system for the study of RNA processing. In "Bacteriophage T4: (C. K. Matthews, E. M. Kutter, G. Mosig, and P. B. Berget, eds.), pp. 208-217. ASM Press, Washington, D.C.
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2 Barnase and Barstar R O B E R T W. H A R T L E Y Laboratory of Cellular and Developmental Biology National Institute of Diabetes and Digestive and Kidney Diseases National Institutes of Health Bethesda, Maryland 20892
I. Introduction II. Gene Structure A. Barnase B. Barstar III. Activity and Structure A. Assay B. Barnase Structure C. Barnase Activity D. Barstar Structure E. Barnase-Barstar Complex IV. Barnase, Barstar, and the Folding Problem A. Barnase B. Barstar C. Barnase-Barstar Reaction V. Barnase and Barstar Homologs A. Bacillus B. Streptomyces VI. Other Practical Applications of Barnase VII. Concluding Remarks References
I. I N T R O D U C T I O N
Barnase and barstar are two small bacterial proteins, one a ribon u c l e a s e a n d t h e o t h e r its specific i n h i b i t o r . I n a d d i t i o n t o t h e s e p r o t e i n s , 51 RIBONUCLEASES: STRUCTURESAND FUNCTIONS
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several known homologs of barnase and possible relatives of barstar will also be discussed in this chapter. The study of all these proteins is driven primarily by their usefulness as models of protein folding, but an attempt will be made to cover all aspects of their study. Barnase, an extracellular enzyme secreted by Bacillus amyloliquefaciens strain H (IAM1521), is one of a family of small microbial ribonucleases (Hartley, 1980; Hill et aL, 1983). Known homologs, in addition to those from other species of Bacillus, include products of several Streptomyces strains and the group of fungal enzymes related to ribonuclease T1 (the last group will be covered in Chapter 3, this volume). Barnase was first reported and purified as an extracellular ribonuclease of Bacillus subtilis strain H by Nishimura and Nomura (1959). Once considered a strain of B. subtilis, strain H (IAM1521) and several other highly amylolytic strains were shown to be genetically distinct by Welker and Campbell (1967), who proposed the designation B. amyloliquefaciens. The study of barnase in this author's laboratory was initiated with the specific idea that it might be developed as an ideal protein for investigation of protein folding. This was based on its small size (110 amino acids, 12,382 Da) and early findings that it contains no disulfide bonds (Nishimura and Nomura, 1959) and requires no nonpeptide adjuvants such as divalent cations for function (Hartley, 1968). Also, it unfolds reversibly in a highly cooperative, two-state fashion under the influence of high temperature or destabilizing reagents such as urea or guanidine hydrochloride (Hartley, 1968). It is this two-state behavior, with an equilibrium between molecules that are either native (N) or unfolded (U) and negligible populations of intermediate states, which allows complete thermodynamic analysis of the molecular folding, including determination of the Gibbs free energy (AG) separating the end states. As we shall see, the small size and simplicity of barnase have also allowed considerable progress in understanding its folding pathways and its folding intermediates by a combination of kinetic analysis and directed mutagenesis (protein engineering). Barstar is an even smaller protein, with 89 amino acids (10,213 Da). Produced intracellularly by the same organism, it is a specific inhibitor of barnase. Although barstar has two cysteine residues, it is now clear that these do not form a disulfide bond and probably carry free sulfhydryls in vivo. Barstar(CCAA), a mutant with alanines in place of its two cysteines, is functional in vitro and in vivo (Hartley, 1993). First reported by Smeaton et al. (1965), barstar inhibits barnase by forming with it a tight, one-to-one, noncovalent complex (Hartley and Smeaton, 1973), thereby
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prohibiting access of substrates to the active site of the enzyme (Guillet et al., 1993a). The dissociation constant for the wild-type barnase-barstar complex is on the order of 10 -14 (Hartley, 1993; Schreiber and Fersht, 1993a). Like barnase, barstar can undergo reversible two-state unfolding, albeit in a more limited range of solvent conditions and with perhaps more residual structure in the unfolded state. It provides, therefore, still another small protein composed of nothing more than a linear peptide, ideal for thermodynamic and kinetic studies of protein folding, and its use for such purposes is also being pursued vigorously in several laboratories. It is generally agreed that a very early phase of protein folding involves a compaction from a more or less random conformation to one that is much more compact and contains elements of the secondary structures seen in the final folded form. The extent to which this collapse to a "molten globule" (Ptitsyn, 1995; Christensen and Pain, 1994; Kuwajima, 1989) begins with secondary structure (the framework model) or with a coalescence of hydrophobic groups (the hydrophobic collapse model) remains to be seen. Barnase and barstar are on high the short list of proteins being used to relate these models to the real world. At low pH some proteins denature to a stable "A state" (Kuwajima, 1989), which bears some resemblance to the compact intermediates seen in folding and unfolding at neutral pH. As we shall see, these include barstar and some barnase mutants. In addition to the two individual proteins, barnase and barstar form a bimolecular complex (or two-subunit protein) of near minimum size that can be approached by both kinetic and equilibrium methods and manipulated by directed mutagenesis of either or both proteins. Because the genes for both barnase and barstar have been cloned and expressed in Escherichia coli, they and many of their mutants can be produced in multimilligram quantities. It should be noted at this point that expression of the barnase gene without concomitant barstar production is lethal, in E. coli or elsewhere. This applies as well to any barnase mutant genes whose products retain activity. Genes for those active mutants which are poorly inhibited by barstar are pathogenic or lethal in our normal barstar-carrying vectors and require special handling. Before the genes for barnase and barstar were cloned, it was, of course, necessary to obtain these proteins from their original source. Little was or is known about the control of the genes for synthesis of barnase or barstar in B. amyloliquefaciens. Investigation of a paradoxical stimulation of ribonuclease synthesis by actinomycin D (May et al., 1968) has suggested this effect involves the lifting of repression by inorganic phos-
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Robert W. Hartley
phate. Direct control by phosphate in the medium, however, is not seen. The gene for the related binase from Bacillus intermedius, on the other hand, is tightly controlled (repressed) by inorganic phosphate (Znamenskaya et al., 1995). In a very rich complex medium barnase is produced during exponential growth and for several hours thereafter. Crude yields are on the order of 3 or 4 mg per liter. Final yields of reasonably pure protein from complex media are seldom more than 5% of the crude initial yield. Defined media have been devised, simplifying the isolation of a cleaner product in comparable final yield (---0.2 mg/liter culture) in spite of much lower initial production (Hartley and Rogerson, 1972). Barstar yields were unpredictably variable in the range of 0.01-0.15 mg/ liter. Today, the recombinant barnase gene in E. coli can provide yields of 80-100 mg/liter, most of which survives the purification procedure. Recombinant barstar yields range up to 200-300 mg/liter. Whereas this author was largely responsible for the early development of the barnase-barstar system, its present popularity is based on the extensive study of the folding of barnase, and more recently of barstar and the complex, by the large group under Alan Fersht at the Institute for Protein Engineering in Cambridge. Their application of a broad spectrum of modern physical techniques, combined with protein engineering, has made the folding kinetics and pathways of barnase among the best understood for any protein. Several papers that review and pull together much of their work are highly recommended to anyone with an interest in how proteins fold (Serrano et al., 1992c; Fersht, 1993; Matouschek et al., 1994). A smaller but also excellent group under Jayant Udgaonkar in Bangalore have also mounted a similar attack specifically on the barstar fold. A short historical review of the barnase-barstar system was published in 1989 (Hartley, 1989).
II. G E N E S T R U C T U R E
A. Barnase
It is now clear that early difficulties in the cloning of the barnase gene were due the toxicity of even very low levels of expression in either E. coli or B. subtilis. Eventual success involved cloning the transposon-
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Barnase and Barstar
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inactivated gene, mutating the codon for the essential active site residue His-102 to aspartic acid and recombining with a separately cloned fragment that overlapped the transposon insertion site (Paddon and Hartley, 1985, 1987; Hartley and Paddon, 1986). With the complete gene for an inactive barnase on a plasmid in either B. subtilis or most strains of E. coli, barnase antigen secretion can be detected with the help of a rabbit antibarnase serum. As originally reported (Paddon and Hartley, 1987), E. coli strain BMH71-18 does not correctly process and secrete the protein, but this is not true of most strains. Western analysis after S D S P A G E revealed that the B. subtilis product contained, in addition to mature barnase, two probarnases, one with 4 extra amino acids as seen with B. amyloliquefaciens, and another with 14. The latter would appear to be the primary probarnase released by the signal peptidase. A study of the role of its signal peptide in the posttranslational export of barnase from B. subtilis (Chen and Nagarajan, 1993) found that barnase on a heterologous signal peptide is secreted faster than on its own, and yet the barnase signal sequence exports another protein faster than it does barnase. With transfer of the mature barnase sequence to the signal sequence of the E. coli phoA gene, on any of several promoter sequences, barnase is secreted by E. coli in much larger quantities. Under some conditions, however, the barnase is released with a portion carrying an extra seven N-terminal amino acids derived from the C terminus of the phoA signal peptide (R. W. Hartley and L. V. Znamenskaya, 1995, unpublished). When the gene for barnase(H102D), preceded by the phoA promoter and signal sequence, was placed on a plasmid containing a functional barstar gene and Asp-102 then mutated back to histidine, active barnase was secreted by E. coli into the medium (Hartley, 1988). Substitution of the synthetic tac promoter, a hybrid of the trpA and lac promoters (DeBoer et al., 1983), for the phoA promoter allows control by the lac inducer isopropyl-/3-D-thiogalactopyranoside (IPTG) in a rich medium without the need for low phosphate. A host carrying the lac[ Q gene for overproduction of the lac repressor is required and overinduction of the barnase gene by IPTG is lethal, but, with limited induction (4 to 10/xg/ ml IPTG), yields of 20 mg/ml are comparable to those obtained with the p h o A promoter. Okorokov et al. (1994) replaced the tac promoter by the PR of A phage on a cassette that included the gene for the temperature-sensitive A repressor CI857. With this construction, yields of 100 mg/liter or so can be obtained simply by growing the E. coli host in a rich medium at 37~
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Robert W.
Hartley
In E. coli or B. amyloliquefaciens barnase is secreted first into the periplasm and then, more slowly, into the growth medium. In E. coli its partition between periplasm and medium at harvest appears to depend both on culture conditions and on the host strain. It can be released from the periplasm by osmotic shock or by acidification. A common practical procedure for recovering the enzyme at harvest is to add acetic acid to 5% to the entire culture and then to adsorb the barnase from the clarified medium onto a strong cation exchanger such as phosphocellulose (Hartley and Rogerson, 1972). After step elution from this ion exchanger, a single salt-gradient chromatographic run on a cation exchanger such as CM-cellulose or SP-Trisacryl provides barnase pure enough for most purposes. The same procedure works well with most barnase mutants.
B. Barstar
The barstar gene as cloned on its own promoter provides a crude yield of about 50 mg per liter of culture. Substitution of the tac promoter increases this to 300-500 mg/liter. Unlike barnase, barstar continues to increase for 12 to 18 hours after cell growth ceases in a rich medium at 37~ In E. coli HB101, which lacks the lacl Q gene for overproduction of lac repressor, the IPTG inducer has little effect and is unnecessary. The harvested cells may be extracted with acetone and dried, after which the barstar is readily released into an aqueous buffer such as 0.2 M ammonium acetate, pH 8. Alternatively, the cells may be lysed by added lysozyme or by using the self-lysing strains of E. coli, BL21 (pLysE) or BL21 (pLysS) (Schreiber and Fersht, 1993b). Purification involves gelfiltration chromatography on Sephadex G-75 and salt-gradient chromatography on a D E A E ion exchanger. Purification can also be carried out on an affinity column of immobilized barnase (Hartley et al., 1972), but this procedure suffers from the difficulties of scaling up and the necessity of allowing the protein to renature slowly as the denaturing detergent used for its elution is removed by dialysis. As far as is known, no one has tried the reverse procedure of purifying barnase on a barstar column, which might actually be more practical. The requirement for barstar in a bacterium carrying the barnase gene is very stringent. In E. coli, even to carry a barnase gene without any promoter in the absence of barstar requires extraordinary measures,
2 Barnase and Barstar
57
such as concomitant synthesis of antisense RNA and/or the appropriate insertion of transcription terminators. It is no wonder, then, that evolutionary selection has produced and maintained such an excellent inhibitor.
III. ACTIVITY AND STRUCTURE A. Assay
The ribonuclease activity of barnase can be determined by the classical procedure of measuring acid-soluble products released from RNA (Rushizky et al., 1964), or, with more precision but less sensitivity, by measuring spectral changes in dinucleotide substrates (Osterman and Walz, 1978). A fluorogenic substrate, polyethenoadenosine, now provides an assay which is both sensitive and precise (Fitzgerald and Hartley, 1993). It also sometimes has the advantage of being relatively specific, being unaffected by most other ribonucleases, including those of E. coli, the barnase homologs from S t r e p t o m y c e s or the fungi, or ribonuclease A. This polymer is, however, a substrate for staphylococcal nuclease. Barstar may be accurately measured by assaying a standard barnase solution as it is titrated by barstar. The three-dimensional structures of barnase, barstar, and that of their complex in crystals have been determined by X-ray diffraction crystallography and the structure of each of the two proteins alone in solution by NMR. The crystallographic structure of barnase was first solved by Mauguen et al. (1982). That of the complex of barnase with barstar (CCAA) was solved independently by Guillet et al. (1993a) and by Buckle et al. (1994), and that of the wild-type barstar, by A. Lapthorn and Y. Mauguen (1994, personal communication). The Cambridge group is also responsible for the NMR solution structures of barnase (Bycroft et al., 1991) and barstar (Lubienski et aL, 1994).
B. Barnase Structure
The structure of barnase (Mauguen et al., 1982) is shown in Fig. 1. It consists of a five-stranded antiparallel/~ sheet, three a helices, and an
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Robert W. Hartley
Fig. 1. Orthogonal views showing the structural elements of barnase. Rotation about the y axis. Produced using Molscript (Kraulis, 1991).
assortment of loops and turns. The elements of secondary structure, in order from the N terminus, are (6-18) helix 1; (19-25) loop 1, with (21-24) forming a type 1 turn; (26-34) helix 2; (35-40) loop 2; (41-46) helix 3; (46-49) a type 2 turn; (50-55)/31 strand; (56-69) loop 3; (70-76) /32 strand; (77-84) loop 4; (85-91) /33 strand; (91-94) a type 1 turn;
2
59
Barnase and Barstar
Fig. 1. (Continued)
(94-99)/34 strand; (100-105) loop 5; (106-108)/35 strand. Except for a /3 bulge in the/31 strand at residues 53 and 54, the hydrogen bonding pattern of the/3 sheet is regular (Bycroft et al., 1991). The longest helix, helix 1, is packed against a side of the/3 sheet to form the largest of three internal hydrophobic regions, core 1. Core 2 is made up of side chains from helices 2 and 3,/31, and loops 1 and 2. Core 3, on the face
60
Robert W. Hartley
of the/3 sheet opposite helix 1, is centered on Leu-63 of loop 3 and Leu89 of the/3 sheet (strand/33).
C. Barnase Activity Barnase is, of course, a ribonuclease, and its principle activity is a transesterification, converting 3 ' ~ 5' internucleotide phosphate groups to 2' ~ 3' intranucleotide cyclic phosphates, with a slower secondary function of hydrolyzing the 2'-phosphate bond. The structurally very different bovine ribonuclease A (see Chapter 9, this volume) catalyzes these same reactions, although with different nucleotide base specificities. The early assumption that the general mechanism of transesterification by the microbial ribonuclease family would be similar to the classic ribonuclease A mechanism (Richards and Wyckoff, 1971) appears to be true for barnase (Mossakowska et al., 1989). Where ribonuclease A uses two histidines as proton donor and receiver, barnase uses a histidine (His-102) as donor and a glutamic acid (Glu-73) as receiver. These two residues are conserved in all the prokaryotic homologs of barnase and the histidine is conserved in its eukaryotic relatives as well. Mutation of either of these residues to alanine virtually eliminates activity. Other conserved residues that appear to be involved in binding or catalysis are Lys-27, Glu-60, Arg-83, and Arg-87. Mutation of Lys-27 to alanine (Mossakowska et al., 1989) reduces activity against both GpA and RNA by two orders of magnitude. The binding (Kin) of GpA is hardly affected, almost the whole change being due to a decrease in kcat, implying that Lys-27 is involved in lowering the energy of the transition state. The corresponding lysine residue in binase, the close homolog of barnase from Bacillus intermedius, appears to be much less critical (Yakovlev et al., 1994), suggesting that there must be subtle differences between the two structures which affect the manner by which the transition state is stabilized. The active site of barnase lies on the/3 sheet opposite the large helix 1 in a groove between loop 2 and helix 2 on one side and loop 3 on the other. Against dinucleotides of the form NpN, barnase is specific for GpN and favors A for the second site, with GpA > GpG > GpC --~ GpU (Mossakowska et al., 1989, Day et al., 1992). The pH optimum for hydrolysis of these substrates is pH 5. Against an RNA substrate, where the optimum pH is 8.5, there still is a preference for
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Barnase and Barstar
61
cutting on the 3' side of G, but barnase cuts other sites as well, so that a complete digest contains only mono- and dinucleotides (Rushizky et al., 1963). Also, although the specific activity of barnase against R N A is about double that of its more consistent guanine specific homolog, ribonuclease T1, its activity against the dinucleotide substrates is about three orders of magnitude lower. Clearly, there are secondary binding sites for R N A in addition to the principle G-favoring site. Two papers from the Fersht group have addressed this problem. Day et al. (1992) carried out kinetic studies with oligoribonucleotide substrates to determine the specificities of such subsites and the sequence positions of the nucleotides involved relative to the primary guanidine. Buckle and Fersht (1994) achieved a high-resolution crystallographic structure of barnase complexed with a deoxytetranucleotide inhibitor d(CGAC). As the substrate is lengthened from GpN, by far the greatest increase in the rate of transesterification, by about three orders of magnitude, comes with the addition of a 3' phosphate to the second nucleotide. The increase is due to a 10-fold reduction in Km and a 100-fold increase in kcat. Addition of a C to make GpUpC or GpApC provides another substantial increase. Because the Km actually increases with this step, the authors suggest that the binding of the third base forces a change in the position of the second phosphate in such a way as to reduce the energy of the transition state. There is some specificity to the binding site for the third nucleotide also, in that A is preferred over C, most of the difference, which is not great, being in Km. Addition of Cp 5' to the primary ~uanine has little effect, nor is that nucleotide seen in the crystal structure, implying that there is no significant binding site for a nucleotide or its phosphate at this position. Several groups have approached questions of barnase specificity by protein engineering. Vuilleumier and Fersht (1994) note that RNase T1 has extra residues, relative to barnase, in the loop after the active site histidine (102 in barnase), but that deciding which residues are extra depends on whether one uses the best sequence alignment or an alignment based on a structural comparison. Attempting to see which might be the better choice, they inserted sequences based on each alignment into barnase and compared the resulting enzymes, finding that both had very low activity. The one based on a structural alignment was affected only in kcat but its overall activity was much lower than the other, leaving no clear choice between the two alignments. On a suggestion that the lower specificity of barnase relative to RNase Sa might be due to an extra hydrogen bond between barnase Ser-37 and the substrate (Sevcik
62
Robert W. Hartley
et al., 1990), this residue has been mutated to glutamic acid (Yakovlev et al., 1993) and to alanine (Kolbanovskaya et al., 1994). The first ap-
peared to increase the guanine specificity for polynucleotide substrates but the second did not, nor does the barnase-d(CGAC) structure support the hypothesis. Bastyns et al. (1994) attempted to affect the guanine specificity of barnase by mutating Glu-60 to glutamine with ambiguous results, finding some reduction of activity against dinucleotides but no effect against polymeric substrates. Figure 2 shows the arrangement of the inhibitor d(CGAC) with respect to the barnase active site, with the target phosphate group centered among the polar ends of Lys-37, Arg-83, Arg-87, and His-102. The guanosine plays a major role in the binding of the tetranucleotide, its guanine base being stacked against the aromatic rings of Phe-56 and Tyr-103 and forming several hydrogen bonds: two to the carboxyl group of Glu-60, two to the peptide backbone, and two to bridging water molecules. The adenine base also contributes to the binding by stacking with the imidazole of His-102 and making several hydrogen bonds. The better part of the interaction energy, however, is provided by the negative charge on the 3'-phosphate of the guanosine, and to a lesser extent on that of the adenosine, interacting with basic groups on barnase. Both NMR (Meiering et al., 1993) and crystallographic (Guillet et al., 1993b) structures are available for complexes of the hydrolysis product 3'-GMP with barnase, as is the crystallographic structure of 3'-GMP with ribonuclease Sa (Sevcik et al., 1991). The nucleotide in each case lies in essentially the same position as the G of the d(CGAC) in the catalytically productive site. The exact conformations of the nucleotide in the crystallographic structures do not all conform to what is expected for the accepted mechanism to operate, but the flexibility seen in the NMR (Meiering et al., 1993) suggests that this is because unproductive conformations are being locked in by intermolecular contacts in the crystals. The same explanation would seem to apply where, in crystals of the complex of barnase with d(GpC) (Baudet and Janin, 1991), the guanidine lies almost precisely in the unproductive adenine site of the d(CGAC). Although it seems clear that the barnase-tetradeoxynucleotide structure accurately represents the barnase-RNA conformation, minor corrections may be necessary because of the missing 2'-hydroxyls. Also, in this structure the two adenine bases in the two complexes of the crystalline asymmetric unit are stacked together, sandwiched between the two His102 imidazole groups. The energetics of this arrangement could perturb the structure.
Barnase and Barstar
63
Fig. 2. The inhibitor d(CGAC) bound to barnase. "P" marks the target 3'-phosphate group of deoxyguanosine. Structure of Buckle and Fersht (1994). Produced using Molscript (Kraulis, 1991).
A n in vivo system has b e e n devised (Jucovic a n d H a r t l e y , 1995) to detect r i b o n u c l e a s e activity in u n s t a b l e b a r n a s e m u t a n t s of low activity for which no in vitro activity can be seen. This involves the use of a plasmid with a cassette carrying b a r n a s e and b a r s t a r g e n e s in such a way
64
Robert W. Hartley
that the barnase gene, from which the signal sequence has been deleted, is strongly repressed and the barstar gene is being expressed. Reversal of the cassette by the lambda int function, triggered by a heat shock, turns on the barnase gene and suppresses that of barstar. Any ribonuclease activity in the barnase, the production of which can be increased by induction of its lac promoter, is lethal. Survival of the bacterium after this reversal establishes a very low limit on the activity of the barnase mutant being tested. The mutants H102K, E73D, R87K, and R83K all test positive for activity by this test, but H102G, H102A, and H102L do not.
D. Barstar Structure
Because it contains no cysteines and none of its residues exhibit any abnormal behavior, the covalent structure of barnase is clear. Barstar, on the other hand, has two cysteines, which immediately suggest the possibility of a disulfide bond, although only a very stable disulfide bond should be expected to form in the reducing milieu of the bacterial cytoplasm. As noted earlier, it seems clear now that the side chains of the two cysteines of barstar exist as free sufhydryls. Confusion on this point may be traced to a publication by this author (Hartley, 1993). Although it was made clear that its activity could not depend on a disulfide bond, because substitution of both cysteines by alanines reduced its binding to barnase only slightly, it was reported that wild-type barstar was destabilized by 2-mercaptoethanol with respect to thermal unfolding in 3 M urea. Indeed, its melting temperature was reduced by 8~ bringing it close to that of the double alanine mutant. This behavior clearly suggested a disulfide bond, although earlier we had found (Hartley and Fitzgerald, 1989) that other barstar preparations had various numbers of free sulfhydryls. I have not been able to prepare another sample of barstar that repeats the anomalous melting behavior, nor have other laboratories seen anything but free sulfhydryls in freshly prepared barstaro(Shastry and Udgaonkar, 1995; Frisch et al., 1995). The more than l l - A distance between the o~ carbons of Ala-40 and Ala-82 (cysteines in the wild type) in the structure (Guillet et al., 1993a) makes the formation of a disulfide bond between them unlikely. Barstar(CCAA) has been much studied, in part because of its less ambiguous primary structure and
2
Barnase and Barstar
65
in part because its crystals, in complex with barnase, provided our first solution of its structure (Guillet et al., 1993a; Buckle et al., 1994). Frisch et al. (1995) have reported that their recombinant barstar preparations uniformly carry the N-terminal methionine from the start codon but without the formyl group. A precise molecular weight obtained by mass spectroscopy on our most recent barstar preparation is in perfect agreement with their structure, although N-terminal sequence analyses of early preparations showed that the methionine had been removed from a fraction of the molecules. The discrepancy may be due to differences in culture conditions and it is probably best to assume that in most preparations most or all of the molecules have 90 residues rather than 89. Authentic barstar from B. a m y l o l i q u e f a c i e n s has no methionine (Hartley et al., 1972). To avoid confusion in barstar numbering, when reference to this residue is necessary, it might be called Met-0. The structure of barstar(CCAA), as it is found in complex with barnase (Guillet et al., 1993a; Buckle et al., 1994), is shown in Fig. 3. Like barnase, barstar is an c~/B protein. A three-stranded parallel/3-sheet forms one side, with one face exposed to solvent. Three helices, very roughly parallel, are packed against the/3 sheet and a fourth closes the open end of this arrangement perpendicular to it. The order and nomenclature of the structural elements are (1-7)/31 strand; (8-11) loop 1; (12-25) helix 1; (26-32) loop 2; (33-44) helix 2; (45-48) loop 3; (49-54) strand/32; (55-63) helix 3; (66-81) helix 4; (83-89) strand/33. The/3 sheet and all four helices surround and contribute to a well-developed hydrophobic core.
E. Barnase-Barstar Complex The structure of the barnase-barstar interface is shown in Fig. 4. Most of the contacts of barstar with barnase in the complex are made by side chains from helix 2 and the adjacent loop 2. Buckle et al. (1994), comparing this structure with the NMR structure of free barstar (Lubienski et al., 1994), report that the entire helical region as a unit moves away slightly from the/3 sheet on binding to barnase. This produces a volume increase calculated at 502 ~3 and may be driven in part by the uncovering of 43 ~2 of the surface of helix c~2, which is then free to contact barnase. Such movement implies considerable plasticity in the hydrophobic core. In this connection, Lubienski et al. (1994) note that, deep within the
66
Robert W. Hartley
Fig. 3. Orthogonal views showing the structural elements of barstar. Rotation about the y axis. Produced using Molscript (Kraulis, 1991).
core, the benzene ring of Phe-74 is able to flip on a time scale fast relative to that of the NMR. Over 800 /Sk2 of surface from each protein is buried in the interface, as barstar helix 2 fits into the active site groove of barnase, parallel to its/3 sheet but perpendicular to t h e / 3 strands. As might be expected,
2
Bamase and Barstar
67
Fig. 3. (Continued)
barstar mimics an RNA substrate to some extent. The carboxyl group of Asp-39 of barstar stands in well for the target 3'-phosphate of the productive binding site, forming hydrogen bonds with barnase residues Arg-83, Arg-87, and His-102, and with a water molecule bound in turn to Lys-27. A main-chain carbonyl and a fixed water partly occupy the other phosphate site. The phenolic group of barstar Tyr-29 takes the
68
Robert W.
Hartley
Fig. 4. Orthogonal views of the b a r n a s e - b a r s t a r interface. The 90 ~ rotation is about the x axis. Barnase is to the right. Elements of barstar are marked by asterisks. Produced using Molscript (Kraulis, 1991)
place of the next base, fitting edgewise against the face of the His-102 imidazole. The barstar Trp-28 interaction with the guanidinium group of Arg-59 corresponds very roughly with the stacking of the latter on guanine in the productive binding site. That guanine site itself, however,
2
69
Barnase and Barstar
Fig. 4. (Continued)
is largely filled with water molecules, including five of nine that mediate hydrogen bonds between the two proteins. These and a few other scattered water molecules fill much of the relatively small amount of free space between the well-fitted proteins. The Ala-40 of the mutant barstar lies at the bottom of the barstar pocket, into which fits the side chain
70
Robert W. Hartley
of barnase His-102. Buckle et al. (1994) suggest that the loss of an - - S H - - i m i d a z o l e hydrogen bond might account for the small decrease in binding of the double Cys-to-Ala mutant. As expected, because barstar binds to barnase much more tightly than does its substrate, as indeed it must in view of its function, there are other links between the two not available to RNA. Barstar covers about 800 A 2 Of barnase, compared to 305 A 2 for d(CGAC) (the first C of which does not contribute). There are a number of polar interactions in the extra surface, the most energetic being provided by barnase Arg-59 in contact with Asp-35 and Glu-76 of barstar (Schreiber and Fersht, 1995). In addition, 45% of the residues in the interface are nonpolar and must contribute substantial hydrophobic energy. Except for the rigid movement of the barstar helices away from the/3-sheet, changes in either structure on binding are minor. Several barstar side chains, notably Asp39 and Tyr-29 of barstar and His-102 of barnase, rotate to better fit their opposing surfaces. While there is little change in the mobility of residues central to the barnase active site, the guanine binding loop (loop 3) is substantially immobilized, especially Arg-59, as well as residues 35-45. Glu-76 of barstar is similarly immobilized. Essentially the same pattern of stabilization in the barnase structure takes place on the binding of d(CGAC). The good preformed fit between the two minimizes entropic energy loss on complex formation. Nor should there be significant energy loss due to the freezing in of water molecules. Half of those seen in the interface are also seen on free barnase. Very likely as many will be found on barstar.
IV. BARNASE, BARSTAR, A N D THE FOLDING PROBLEM
A. Barnase
As mentioned in the introductory paragraphs of this chapter, barnase is ideal for investigations of many aspects of protein folding. In addition to being no more than a simple linear peptide that can unfold reversibly in vitro, two further advantageous properties became evident once the structure was known. (1) Many proteins that unfold reversibly have, while folded, proline residues in the cis conformation. On unfolding, these residues will mostly revert with time to the favored (where uncon-
2
Barnase and Barstar
71
strained) trans conformation. On returning to conditions that favor the folded form, the relatively slow isomerization of such residues back to cis can be the rate-limiting steps on the path or paths back to the native fold (Brandts et al., 1975). In some cases this may allow transient observation of trapped intermediate states, although it masks the more general, and especially the final, folding steps, which are much faster than the proline isomerization. All three prolines in barnase, however, are in trans. The cis-prolines that appear at equilibrium in the unfolded form of barnase are too few to interfere seriously with kinetic studies of the complete folding process (Matouschek et al., 1990). (2) The major elements of secondary structure, the three helices and the/3 sheet, are separated along the sequence so that these structures can be isolated and studied alone or as complementing fragments. It is important to stress the two-state nature of the barnase unfolding transition. A highly cooperative transition, with a negligible fraction of the molecules in states other than F or U (folded or unfolded) at any time, is required if real thermodynamic parameters are to be determined simply from transition measurements. For a true two-state transition, all measurable local or global parameters that change through the transition must yield the same curve when plotted as a fraction (fu) of the total change versus temperature or denaturant concentration. Another strong requirement for thermal unfolding is that the van't Hoff enthalpy (AHvH = - R T d In K v / d T , K u is equilibrium constant for unfolding) be equal to the calorimetrically measured heat Ancal of unfolding. It was shown early (Hartley, 1968, 1969) that, in the neutral pH range, barnase met the first criterion for several disparate parameters and the very high value of AHvH (1.4 kcal per residue) suggested it might meet the other as well. Calorimetric measurements (Makarov et al., 1993; Griko et al., 1994; Martinez et al., 1994) have recently indicated that it does indeed and several groups (Pace et aL, 1992; Martinez et al., 1994) have presented evidence that the unfolding of barnase is more complete (at pH 6.3) than for other proteins studied. It has also been demonstrated that barnase undergoes a strictly two-state transition on titration with the anionic detergent sodium dodecyl sulfate (SDS) (Hartley, 1975), but in this case a less than random conformation in the nonnative state is to be expected. Since recombinant barnase became available in 1988, it has been the subject of intensive study by the Fersht Cambridge group. Considerable progress has been made toward measuring and interpreting the energetic contributions to stability of various residues and structures and in under-
72
Robert W. Hartley
standing the pathways through which barnase unfolds and folds. An attempt here will be made to outline the experimental procedures used and to summarize their major results and conclusions and related reports by other groups as well. Equilibrium and kinetic observation of wildtype and mutant barnases have been the primary approaches, but other methods, in concert or independently, have not been neglected. For observing folding or unfolding in the two-state transition of barnase, the main parameter of choice has been tryptophan fluorescence (Hartley, 1975), which has the dual advantages of being very sensitive and undergoing a very large decrease ( - 7 0 % ) as tryptophans are exposed to solvent. The fraction unfolded, fu, is a simple function of the fluorescence and from it can be derived the equilibrium constant Ku and, hence, the Gibbs free energy, A G, within the transition range. A commonly accepted procedure in urea or guanidine denaturation (Pace, 1986) is to make a linear extrapolation of AG from the transition range to zero denaturant to derive AGu,w, the free energy of unfolding in the absence of denaturant. For barnase, at pH 5 or pH 6, where it is most stable, there is a consensus value of AGu,w by this method of about 9 kcal mol -~ at 25~ (Pace et al., 1992, Kellis et aL, 1989, Serrano et al., 1992a), falling slightly with pH up to pH 9 and dropping off more rapidly below pH 5 (Pace et al., 1992). Note that barnase is a basic protein, with an isoelectric point of about pH 9, so that its stability is not a strong function of net charge. Johnson and Fersht (1995), applying differential scanning calorimetry (DSC) to barnase in urea concentrations over a range from 0 to 4.5 M, have concluded that the linear extrapolation yields an underestimate of AGu.w, with 10.5 kcal mol -~ giving the best fit to their data at pH 6.3. The AGu,w for barnase can be determined by examining its thermal transition as well. This requires a rather more complicated argument (see, for example, Kellis et al., 1989), but the values obtained are in reasonable agreement with those derived from unfolding in urea. AGu is a measure of stability and the accurate distribution of its value between AHu and - T u A S u can tell us a good deal about the factors contributing to that stability. The absolute value of each is therefore of considerable interest. In most current work on the role of particular residues and their contexts in determining the fold of a protein and its stability, however, it is the differences in these values in mutants and in the wild type, or among different mutants, that are of special interest. These differences can be determined with much greater accuracy than the
2
Barnase and Barstar
73
absolute values, especially where measurements are made in parallel at the same time. This point has been made often by the Fersht group. [See especially Matouschek et al. (1994), where the accuracy of both the absolute and the differential measurements of kinetic as well as the thermodynamic parameters is assessed in depth.] The change AAGu,w in the free energy of folding produced by a mutation in barnase can, then, be determined with some confidence. The problem remains of rationalizing the change found in AGu in terms the gain or loss of specific bonds or degrees of freedom. Any element of a protein normally contacts more than one other element, and changing it or deleting it will reflect the energy gain or loss due to changes in each contact, and further changes due to consequent motions of the neighboring elements, and so on. In some cases this is reasonably straightforward, where the energy can be attributed to changes in van der Waals contacts alone or the loss of hydrogen bonds, with no new bonds, steric interference, or local disorganization introduced by the mutation. In other cases, double mutant cycles (Serrano et aL, 1991, Fersht et aL, 1992) can be carried out. This involves measuring AAGu,w for mutations at each of two neighboring residues and for the double mutant, allowing partial subtraction of the influence of other neighbors. The procedure can be used successively on a series of connected residues (Horovitz et al., 1991). A large number of mutants have been studied in this manner, chosen, for the most part, so as to produce minimum disruption of surrounding structures. A paper by Serrano et al. (1992a) collects and extends previous data on mutations affecting hydrophobic, hydrogen bonding, and charge-charge interactions and analyzes their respective contributions toward stability. Deletion of a methylene group buried in a hydrophobic region produces an average loss of free energy of about 1.5 kcal mo1-1, in fair agreement with a similar estimate by Pace et al. (1992). The deletion of partially buried hydrogen bonds produces more variable effects than for those either deeply buried or on the surface. For five instances where holes are produced in a hydrophobic core by - - C H z - deletions, it was found by X-ray crystallography (Buckle et al., 1993) that the holes contain no solvent molecules and that as little as 10% and as much as 90% of each hole is filled by the moving in of neighboring groups. The variation in AAGu.w is high even for mutations that are descriptively very similar, and the detailed context for each case is clearly important. The importance of context is even more evident for mutations affecting hydrogen bonds and charged groups, which are also more difficult to
74
Robert W.
Hartley
categorize. The AAGu,w for mutations disrupting H bonds range from negligible, where fully solvent-accessible H-bonding groups are deleted, to near 3 kcal mo1-1 for some affecting fully buried groups. Two deletions of hydrogen-bonding hydroxyl groups that contribute 1.4 and 1.9 kcal mo1-1 to stability (Serrano et al., 1992a) have been studied by X-ray crystallography (Chen et al., 1993). These mutations do not introduce new interactions or disrupt other aspects of structure so that their effect on stability can be ascribed primarily to the loss of hydrogen bonds (two each). Mutations removing one charge from a fully buried pair is even more destabilizing (Fersht, 1972, 1988), but no such mutants are available for barnase. The much more common charge-charge interactions on the surface are of minor importance, with the closest pairs contributing no more than about 1 kcal mo1-1 to folding energy. A pair of tyrosines on the barnase surface interact edge to face and contribute 1.3 kcal mo1-1 (Serrano et al., 1991). A series of papers from Cambridge dealing with the both the C and N caps and internal sequences of the helices were summarized and extended in two papers by Serrano et al. (1992a) and Horovitz et al. (1992). For helix 1 and helix 2, substitution of each residue in turn by Ala, and multiple substitutions at selected positions, allow the authors to compare and contrast the preferences of these particular protein helices with others and with synthetic helical peptides. Using stopped-flow kinetics, the rates of unfolding and folding can be directly measured over much wider ranges of denaturant concentration than can equilibrium constants, which are limited to a narrow melting range. The rate constant for unfolding, ku, can be measured from zero denaturant to the upper part of the transition range, whereas kF, the rate constant for folding, is available from the lower end of that range to the limit of solubility of the denaturant. Sanz and Fersht (1994) have even extended the measurement of kF above the transition range by trapping refolding barnase with barstar. Matouschek et al. (1990) found that while refolding at low urea concentration barnase passes through a relatively long-lived intermediate state. As barnase unfolds with what appears to be a single rate constant, they identify this intermediate state with the rate-limiting state for unfolding. Using an analysis analogous to that used to get AAGu,w from equilibrium melting curves [see Fersht et al. (1992) for theoretical details], they find values of AAG~,w and AAG$,w for various barnase mutants, where AAGI,w is the increment to the difference in energy between the unfolded state and the intermediate state produced by the mutation, and AAG:~,w is the
2
Barnase and Barstar
75
same for the transition state that separates the folded and intermediate states. The unfolded molecule is taken as the standard state, where Gu = O, on the assumption that there the differences in free energy between wild type and mutants are minimal. Parameters of the form Cz = A A G z / A A G F are defined, where z identifies some state on the unfolding pathway. Thus we have CF = 1 for the folded state, Cv = 0 for the unfolded state, and r and r refer to the most stable intermediate and the transition state for the ratelimiting step. A sample of how these r values are plotted against the transition parameter is illustrated in Fig. 5. The r parameters may be safely interpreted when their values are near to either 0 or 1, which is frequently the case. A value of r of near zero, for example, means that the mutated residue is in a region that is unfolded in the intermediate state, whereas a value near one puts it in a folded region. The fact that the r values are near either 0 or 1 for many mutants gives us confidence that each of these states represents a limited number of similar conformations (see also Hakoshima et al., 1991). The Cambridge group has applied this analysis to kinetic measurements of a large number of mutants (Serrano et al., 1992b; Matouschek et al., 1992a). By examining mutants in the various structural motifs of barnase, they have been able to build up a remarkably detailed picture of the order of folding and, perhaps even more important, have established that such a compulsory order of folding exists. Folding appears to begin with a rapid collapse of a central hydrophobic core (core 1) with early formation of the central portion of the/3 sheet (/32,/33, and 134) and an N-terminal portion of helix 1, which provides most of the side chains of the core. On the other side of the 13 sheet, in conjunction with parts of loops 3 and 5, the precursor of core 3 is also formed early, as is the one-turn helix 3 and part of helix 2. The latter appear to form separately and only pack, along with loop 4 and the edge of the 13 sheet, to form core 2, after the rate-limiting transition. Between the intermediate and transition states, cores 1 and 3 are reorganized to something close to their folded forms, perhaps with the removal of some solvent. Independent corroboration of this sequence of events has been provided (Bycroft et al., 1990; Matouschek et al., 1992b) by application of the deuterium exchange-NMR method of Udgaonkar and Baldwin (1990). This procedure observes folding by following the time course by which particularly H-bonding amide hydrogens become protected from rapid exchange with solvent. The results provide clear evidence that
76
Robert W. Hartley 1.20 1.00 0.80
c 14V o N5A -.--e--- N23A
/ ,/I
N77A
0.60
II
///
0.40 0.20 0.00 -0.20
I
I
I
I
Unfolded IntermediateTransition Folded State
1.20 1.00 0.80 0.60 0.40 0.20 0.00 -0.20
/ d/
••o
D12/R110 Y13/Y17 r T16S ---=-- N58A = K62R
~' /'
o
I
l
I
l
Unfolded Intermediate Transition Folded State
1.20 1.00 0.80 0.60 0.40 0.20 0.00 -0.20
i• I
l
1
r
V10A L14A 188V
=
196V 1
Unfolded Intermediate Transition Folded State
Fig. 5. Examples of $ plots. From (a) Residues seen to be close to the unfolded condition in the intermediate and transition states. (b) Residues are already fixed in the intermediate and transition states. (c) Intermediate cases. Fersht (1993). Matouschek, et al. (1994), in Mechanisms of Protein Folding. (R. H. Pain, ed.). Reprinted by permission of Oxford University Press.
2
Barnase and Barstar
77
barnase folds through at least one relatively stable intermediate. Where there is overlap with the protein engineering data, the agreement is excellent. The study of complementing peptides, where independent fragments can combine to produce a complex with all or part of the activity of the whole protein, has been of considerable importance in the history of the folding problem beginning with the discovery of ribonuclease S by Richards (1958) (see Chapter 9, this volume). In early studies of barnase (Hartley, 1970, 1977) it was shown that barnase (1-102), which had no activity but could adopt a less stable but apparently native fold, could be complemented by barnase (95-110) and longer C-terminal peptides but not by barnase (99-110) or shorter peptides. The last eight residues, although they stabilize barnase and indeed are in position early in its folding (Fersht, 1993), are, therefore, not crucial to the barnase fold. His-102 is essential for activity, and the complementation results imply that the complementing peptide must be able to displace the f14 strand in order to fix loop 5 and His-102 in place. The Cambridge group has produced and studied two complementing barnase systems, fragments (1-36) plus (37-110) (Sancho and Fersht, 1992; Sancho et al., 1992) and (1-21) plus (23-110) (Kippen et al., 1994; Kippen and Fersht, 1995). Barnase was cut by substituting a methionine at the target site and cleaving with cyanogen bromide and the sites were chosen as lying between units of secondary structure along the sequence. In the first case, both peptides are separately inactive and mostly random but with some traces of structure, including probably some of helices 1 and 2 in (1-36). In combination, a largely native structure is formed with substantial activity. Cut at Met-22, both fragments again show traces of native structure, and fragment (23-110), which contains all the residues involved in the active site, alone has about 1% of the activity of uncut barnase(I36M). Furthermore, fragment (23-110) binds to and is stabilized by both the deoxytetranucleotide inhibitor d(CGAC) and by barstar, which inhibits it at a 1:1 molar ratio. Note that whole barnase is stabilized by the binding of 3'-GMP (Martinez et al., 1994) and (at least at pH 4.5) by phosphate alone (Meiering et al., 1991). The equivalent experiment with d(CGAC) has not been reported. The activity of (23110) increases sevenfold in the presence of (1-22) or synthetic (5-22) and regains much of the native circular dichroism (CD) spectrum. Among other things, these experiments show that, as with ribonuclease S and a number of other proteins, a continuous peptide chain is not a necessary condition for folding, which must then involve multiple nuclei or incipient
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Robert W. Hartley
or transient secondary structures that come together and are finally reorganized by tertiary interactions. The 6 to 12 portion of helix c~l appears to be one such nucleus and the hairpin formed by/33 and/34 has been suggested (Serrano et al., 1992c) for another, and these results combined with the kinetic pathways (Fersht, 1993) suggest that the helix 2-helix 3 region is a third. Another group, pursuing the theory that all globular domains are made up of modules derived from exon-shuffled ancestors, have divided barnase into six such modules ("compact contiguous segments") and synthesized each (Ikura et al., 1993; Yanagawa et al., 1993). Two of these, modules (24-52) and (52-73), were found to have some residual structure by NMR. The first, which includes helices 2 and 3, showed signs of the helix, and some/3 turn was indicated in both. Module (1-24) (Yoshida et al., 1993, including the large helix 1 in barnase, polymerizes slowly (in 50 mM Tris-HCl, pH 7.5, at 5~ into helical filaments 10 to 20 nm in diameter and containing a substantial amount of c~ helix as seen by CD. The unfolding kinetics of barnases with two different disulfide bonds introduced by protein engineering (Clarke and Fersht, 1993) lend strong support to the Cambridge picture of ordered folding. These bonds were introduced by substituting cysteines for Ser-85 and His-102 or for Ala43 and Ser-80. The first joins the 133 strand with loop 5 in a portion of the structure found to form early, whereas the second joins helix c~3 with the late-forming loop 4, which, in the wild type, come together only in the final steps of folding. In both cases the disulfide form is more stable than the wild type, although in reduced form the substitutions are slightly destabilizing. When the equilibria and unfolding kinetics of the oxidized and reduced forms are analyzed, however, it was found that the 43-80 bond stabilizes only the final fold and has a large effect on the rate of unfolding, whereas the 85-102 bond stabilizes both the intermediate and transition states as well but has much less effect on the rate of unfolding. These are exactly the results expected for links that stabilize structures formed early (85-102) and late (43-80). Perrett et al. (1995) have used amide proton exchange to identify those amides that exchange, at least in part, by local breathing of the structure, and those that only exchange on global unfolding. They conclude that exchange by breathing correlates with nearness to the surface of the structure but not necessarily with lateness in folding. Matouschek and Fersht (1993), using an extension of their theory, have added information from the perturbation of folding rates and equi-
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Barnase and Barstar
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libria by the denaturant (urea), finding that mutations can move the transition state along the reaction coordinate and that the closer the energy of the transition state is to that of the folded state, the closer also is its structure. This has been applied to a study of Ala and Gly mutations of all solvent-exposed residues of the two major helices. Their most surprising conclusion is that mutants in the helix 1 that bring the transition state, and hence the overall structure, closer to the folded state are those mutants that destabilize the helix. They suggest that in such mutants the helix 1 forms later and hence the order of folding can be changed without affecting the final fold. In the course of their studies on many mutants, the Cambridge group found a number that stabilize the intermediate state with respect to both the folded and unfolded states. Although the intermediate state did not accumulate appreciably in any of these during equilibrium unfolding, Sanz and Fersht (1993) were able to combine such mutations to produce double and multiple mutants that allow the accumulation of 40-50% of the molecules as intermediate forms. It is suggested that, as the denaturant concentration is increased, the first intermediate that appears after the rate-limiting step subsequently unravels its secondary structure in a less cooperative manner. They predict that the structure of these intermediates should be accessible by NMR. Meiering et al. (1992) compared the effects of active site mutations on stability and activity. For three of the positively charged residues clustered in the active site, mutation to remove a position charge increases overall stability, implying that this clustering represents a compromise between stability and activity. Thus suggest further that the local instability engendered by these charges may be important for full activity. At low pH the thermal behavior of barnase unfolding becomes more complex (Makarov et al., 1993; Sanz et al., 1994; Martinez et al., 1994). In DSC of barnase, below pH 4, at low ionic strength the excess heat capacity peak shows signs of asymmetry, accomplished by a decrease in the ratio of van't Hoff to calorimetric enthalpy, which reaches a minimum at about pH 2.7. [The asymmetry is much more pronounced in its close homolog binase (Protasevich et al., 1987), but without the effect on enthalpy.] Adding 0.2 M KC1 eliminates both the asymmetry and the effect on enthalpy. Barnase(S85C, H102C), with a disulfide bond stabilizing the 13 sheet and major core, maintains two-state behavior at low pH, even at low salt. All of these reactions, two-state or not, are fully reversible. Those mutants designed to stabilize the folding intermediate (Sanz and Fersht, 1993), however, show an asymmetry that is concentration
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and rate dependent, indicating aggregation, and are not reversible. The long- and short-wave UV CD spectra of these aggregates suggest nativelike secondary structure but no burial of aromatic side chains. There is a strong implication that the longer lifetimes of the folding intermediates of these mutants allow them to be trapped irreversibly into aggregates. In two papers, Oliveberg et al. (1994, 1995) have begun an in-depth study of the effects of pH on unfolding and on titratable groups, using a thermodynamic analysis of denaturation at low pH and a titration of each barnase acidic group by NMR and protein engineering. They conclude that at low pH the denatured barnase is not so expanded as not to be affected by intramolecular charge repulsion, and that in the thermally denatured state the pK values of the carboxyl groups average 0.4 units lower than expected from model compounds. Arcus et al. (1994) have reported complete NMR assignments for the acid-denatured state. In examining their peaks, they find evidence of relatively stable structures in two regions, Gly-9 to Tyr-17 and Asp-93 to Tyr-97. These are just those regions found by protein engineering to be among the first to fold. [For more detailed summaries and analyses of these works on barnase folding pathways by the Cambridge group, see Serrano et al., 1992c; Fersht, 1993; Matouschek et al., 1994.] The computational approach to protein folding has also been applied to barnase. Prevost et al. (1991) have applied molecular dynamics to the effects of a buried hydrophobic hole, and Braxenthaler et al. (1995) have looked at the unfolding of a helix c~l/loop 1 fragment and Puglicse et al. (1995) have examined the hairpin formed by /33 and /34. A more ambitious study by Caflisch and Karplus (1994, 1995) considered the initial stages of unfolding of the whole molecule. Their computed traces agree in most respects with the experimental deductions of the Cambridge group and are not changed drastically by the addition of 15 protons to simulate strongly acid conditions. They emphasize the role of water, suggesting that solvation of the major hydrophobic core represents the rate-limiting step in unfolding. Loewenthal et al. (1991) have applied directed mutagenesis to uncover the role of each tryptophan residue in fluorescence. Not surprisingly, the buried Trp-35 dominates, but they also report that the fluorescence of Trp-94 is strongly affected by titration of the nearby His-18 and suggest further that there may be energy transfer to Trp-94 from Trp-71. In a similar work, Vuilleumier et al. (1993) evaluated and catalogued the contributions of every aromatic residue of barnase to the long- and shortwave CD spectra.
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The Fersht group has used barnase, along with slower and faster folding mutants, to investigate the function of the E. coli chaperonin, GroEL (Gray et al., 1983; Gray and Fersht, 1993; Corrales and Fersht, 1995). The GroEL 14-mer binds up to four molecules of denatured barnase, preferentially binding those in an early stage of folding. The barnase continues to fold while bound to GroEL but much more slowly than when free. Binding of ATP, not its hydrolysis, speeds the release and folding of barnase, the reaction ending with one molecule of folded barnase bound. Addition of GroES in substoichiometric amounts to the A T P - G r o E L - b a r n a s e complex further accelerates the release. Hojo and Aimoto (1993) have achieved a complete chemical synthesis of barnase at 11% yield and full ribonuclease activity. Their synthetic barnase was site specifically labeled with two atoms of 13C. This accomplishment clearly opens the possibility of observing in unprecedented detail, by NMR, the local effects of any change in barnase.
B. Barstar
With the recent solutions of the barstar structure, alone and in complex with barnase, there has been a marked increase in interest in its folding behavior and in the details of complex formation. That barstar can unfold reversibly has been clear since early preparative procedures included elution from a barnase affinity column by guanidine hydrochloride (Hartley et al., 1972). A number of papers have made it clear that barstar will indeed be another useful folding model and one that differs considerably from barnase. Agashe et al. (1995) have reported that the earliest event of the major folding pathway of barstar is a nonspecific hydrophobic collapse that does not involve concomitant secondary structure formation. Their conclusion is based on the binding of 8-anilino-l-naphthalene-sulfonic acid (ANS) before spectroscopic evidence of secondary structure appears. This fluorescent dye binds and reports on relatively large hydrophobic clusters (Stryer, 1965). This initial collapsed state might resemble the condensed Dphys state in the nucleation-condensation mechanism of Fersht (1995). In equilibrium unfolding experiments between pH 7 and pH 9, both barstar(CCAA) and, in the presence of a reducing agent such as dithioo threitol (DTT), wild-type barstar unfold in a two-state manner with
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a negligible population of intermediate forms in the transition zone (Schreiber and Fersht, 1993b; Khurana and Udgaonkar, 1994). The free energy of unfolding at 25~ about 5 kcal mol-1 for barstar and somewhat less for barstar(CCAA), is only about half that of barnase and indeed lower than most small globular proteins. The enthalpy of unfolding per residue, at any temperature, determined either by differential scanning calorimetry (DSC) (Martinez et aL, 1994; Wintrode et al., 1995) or by isothermal unfolding over a range of temperatures by guanidine hydrochloride (Agashe and Udgaonkar, 1995), is also unusually low. This suggests a lower than expected contribution by hydrophobic interactions to the enthalpy and has been interpreted as a looser than usual packing in the hydrophobic core. Both Wintrode et al. (1995) and Agashe and Udgaonkar (1995) found that the relative temperature dependencies, at pH 8, of folding enthalpy and entropy were such that the free energy passes through a maximum. The latter authors went further and showed that, with a maximum stability at 27~ between about 1 and 2 M guanidine hydrochloride, barstar undergoes cold denaturation at low temperature as well as at the high temperature transition. One of the two prolines of barstar, Pro-48, has the cis conformation in the native fold. The effects of its isomerization to trans on folding and unfolding kinetics have been investigated in some detail (Schreiber and Fersht, 1993b; Shastry et al., 1994). If barstar is kept under unfolding conditions long enough for racemization of the cis-Pro-48 to mostly the trans conformation, and then taken to folding conditions, all of it returns rapidly to a functional folded conformation, but some 75% initially contains the misfolded trans-Pro-48, as revealed by its faster kinetics on reunfolding. Under folding conditions the isomerization of the misfolded trans-proline to cis-proline goes at about the same rate as an unconstrained proline for barstar(CCAA) (Schreiber and Fersht, 1993b), but at three times the rate for wild-type barstar (Shastry et al., 1994). Although the complete thermodynamic description of folding is most straightforward for proteins undergoing the strictest two-state unfolding, the lack of readily observed intermediates complicates detailed observation of folding pathways. As discussed above for barnase, this limitation can be mitigated by mutations that destabilize the native state relative to intermediate states, leading to relatively stable intermediates. Nath and Udgaonkar (1995), identifying a buried tertiary hydrogen bond connecting separate units of secondary structure, mutated the single histidine of barstar, His-17, to glutamine, eliminating the hydrogen bond between
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Barnase and Barstar
83
its imidazole ring and the phenolic hydroxyl of Try-30. Disturbance of the native fold is minimal but unfolding now clearly involves more than one relatively stable intermediate. In 1.5 M guanidine hydrochloride one such intermediate converts to a second with a time constant of several hours. At low pH barstar converts to a stable A state (Khurana and Udgaonkar, 1994; Khurana et al., 1995), resembling to some extent the A state seen under more restricted conditions for some barnase mutants. This A form, which has disrupted tertiary structure and increased exposure of hydrophobic surfaces to solvent but with much intact secondary structure, exists as large soluble aggregates. Conversion from the N to A forms with decreasing pH appears to follow the titration curve of a single group with a pK of 5.0. This titration is not affected by the H17Q mutation, although the buried His-17 has an abnormally low pK (6.2 or lower). Swaminathan et al. (1994), in time-resolved decay measurements of fluorescence intensity and anisotropy, found aggregation and greater surface accessibility and mobility for the tryptophans in the A form. With either increasing temperature or increasing denaturant, the A form gradually loses spectral evidence of secondary structure until, under conditions more severe than for the unfolding transition of the N form at pH 7-8, a final cooperative unfolding takes place coincident with disaggregation. Further complexity of these transitions is indicated by the noncoincidence of different spectral probes. DSC at pH 3 reveals no peak of excess heat capacity. The relationship of the A state to the intermediates seen in the unfolding of barstar(H17Q) is not clear, though both have some resemblance to the molten globule model. A clear difference between the two is that ANS reports accessible hydrophobic patches on the A form but none on the (H17Q) mutant at pH 7 at any stage of unfolding. Above pH 8 the stability of wild-type barstar declines. This is not true for the double Cys-to-Ala mutant even up to pH 10 (Khurana et al., 1995), suggesting that titration of one of the sulfhydryls removes a stabilizing hydrogen bond. Schreiber et al. (1994) found that mutation to alanine of any of four acidic residues involved in binding to barnase decreased the stability of the complex but increased that of barstar itself. Thus, as with barnase, optimization of function combined with a d e q u a t e stability appears to be the evolutionary goal.
84
R obe rt W.
Hartley
C. Barnase-Barstar Reaction
Schreiber and Fersht (1993a) measured both on and off rates, and hence equilibrium constants and Gibbs free energies, of the barnasebarstar interaction for both wild-type and mutant proteins using stoppedflow and chased label techniques. Measurements of equilibria and off rates, in reasonable agreement with these (Hartley, 1993), were based on ribonuclease activity measurements of the free active barnase in mixtures in which an active barnase competes with an inactive one for a barstar. A number of mutations in both proteins that were identified in these papers as affecting binding were of those residues, including several involved in barnase activity, later shown to be directly involved in the interface between the two proteins (Guillet et al., 1993a). Jones et al. (1993) also identified many of these residues on barnase by observing the effect of barstar binding on the NMR spectra of ~SN-labeled barnase and on deuterium exchange. At pH 8 and low salt, an association rate of 6.0 x 108 sec -~ and a dissociation rate of 8.0 x 10 -6 s e c -l M -1 combine to give a dissociation constant (Kd) of 1.3 • 10 -~4 M -l, from which we get a free energy of complex formation of 19 kcal mol -~. Increasing salt decreases stability by decreasing the on rate and especially by increasing the off rate, resulting in a Ka value of about 10 -12 M -1 at 0.5 M NaCI. For the barnase mutations that decrease binding, an increase in off rate is again mainly responsible. Indeed, for such mutations that increase the net positive charge on barnase, the decreased stability is in spite of an increased on rate. Mutation of any of several basic residues in the active site of barnase to alanine reduces the free energy of binding by 5 or 6 kcal mol -~ and a large increase in off rate with decreasing pH follows the titration of His-102 of barnase, the pK of which drops from 6.2 in free barnase to less than 5 in the complex (Schreiber and Fersht, 1995). Applying transition state theory to the measured on rates for a number of mutants, these authors conclude that the transition state for binding depends only on electrostatic interactions. Schreiber and Fersht (1995) also applied the double-mutant cycle analysis of AAG pioneered by the Cambridge group for barnase unfolding (see section IV,A) to the barnase-barstar intersection, allowing them to better estimate the contribution to binding energy of individual pairs of interacting residues and the cooperativity between neighboring pairs. Having determined the coupling energies for all of the pairs that might
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Barnase and Barstar
85
be expected to contribute the most to binding, they conclude that it is not yet possible to predict with any accuracy the contributions of particular pairs from structural information alone. Of the six most energetic coupling pairs, only three had been recognized as being of major importance from structural observations, those between Asp-39 of barstar and Argo83, Arg-87, and His-102 (6.7, 6.1, and 4.9 kcal mol-1). Unrecognized were those between barstar Asp-39, Asp-35, and Tyr-29 and barnase Lys-27, Arg-59, and Hisol02, respectively (4.8, 3.4, and 3.3 kcal mol-1). Even uncharged residues interact significantly at ranges up to 7 A and the Asp-39-Lys-27 pair, contributing 4.8 kcal mo1-1, are separated by 4.5 and a water molecule. Subtleties of the local environment are clearly important. The energetic contributions of mutants separated by more than 8 A, however, appear to be additive, suggesting that none of these mutants produce major rearrangements. It is notable that the sum of the contributions to coupling energy (41 kcal mo1-1) listed by Schreiber and Fersht (1995) alone add up to more than twice the AG of the whole interaction (19 kcal mol-1). Compensating negative contributions presumably include those arising from the distortion and expansion of the barstar hydrophobic core seen on binding (Buckle et al., 1994). DSC studies (Makarov et al., 1994; Martinez et al., 1995) have shown that barnase is markedly stabilized in its complex with barstar, its melting temperature increasing by some 20~ to approximately that of barstar alone. Thermal unfolding of wild-type barstar without a reducing agent such as DTT, alone or in complex with barnase, is largely irreversible. The melting of barstar(CCAA) and of its complex, on the other hand, is simple and is at least 80% reversible between pH 6 and pH 11 (Martinez et al., 1995). The dramatic stabilization of the lower melting component in such a tightly bound complex is explainable on the basis of mass action. At pH 3, the only peak of excess heat capacity is that of unbound barnase. The barnase mutant H102K, as noted earlier, has some residual activity. It binds so poorly to barstar, furthermore, that in a barnase expression vector it is much more toxic than wild-type barnase. This has prompted the development of a vector, related to that used to detect residual activity in barnase mutants, that allows the detection or, indeed, the selection, of barstar mutants that suppress, by improved inhibition, such barnase mutations (Jucovic and Hartley, 1996). Suppressors can be graded, depending on the amount of inducer of the tac promoter (on barnase) needed for lethality. Selection in this fashion from a library in which the six amino acids of barstar that contact His-102 of barnase in the complex were randomized, turned up a number of such suppressors.
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All involved mutation of either Tyr-29 or Tyr-30 or both. Two were single mutants, Y29P and Y30W. Of six double mutants, all but one included Y30W. Tyr-29 is on the barstar surface and interacts strongly with the imidazole of His-102, and its substitution in the suppressor is not surprising, although the range of substitutions is remarkable (Pro, Ala, Asp, Asn, and Arg as double mutants with Y30W). The more effective suppressor mutation, however, is the Y30W. The phenyl group of Y30 is completely buried in barstar, with only its main chain in contact with barnase. Its replacement by the larger indole must cause significant rearrangement of surrounding residues in such a manner as to adapt the barstar surface to that of barnase(H102K). The availability of these suppressor barstars made possible the production and isolation of barnase(H102K) for in vitro work, which confirmed its residual ribonuclease activity, its weak binding to wild-type barstar, and its much stronger binding to a suppressor barstar.
V. B A R N A S E A N D B A R S T A R H O M O L O G S
A . Bacillus
The closer homologs of barnase, from various strains of Bacillus, can be divided into two groups: those which differ from barnase at no more than two or three positions and those bearing a similar relation to binase, the enzyme from B. intermedius (strain 7P). Binase and barnase share about 85% sequence identity, with an almost superimposable structure. These ribonucleases are all inhibited by barstar (Yakovlev et al., 1995) and several have been cloned onto plasmids carrying its gene. Where it has been looked for, evidence for barstar homologs has been found in these strains, including B. intermedius, but despite vigorous efforts in several laboratories they have been cloned only from a few of the strains with ribonucleases very close to barnase. Sequences of barnase and those available for other strains of Bacillus and Streptomyces are compared in Fig. 6. Genes for equivalents of barnase and barstar have been cloned from three other strains of B. amyloliquefaciens (Hartley and Fitzgerald, 1989). Barnase and barstar from strain K (IAM1523) are identical to those of strain H. Barnase (IAM 1193) has a single substitution and barnase(IAM 1170) has that substitu-
2
1)
2)
87
Bamase and Barstar
I
im n
I L k S S D W - L I Y K T T D H Y Q T F T K I R I L Y s s D w - L I Y K T T D H Y Q T F T X I R
Fig. 6. Sequences of ribonucleases secreted by Baciilus and Streptomyces. Bold boxes surround residues conserved through both genera; thin boxes outline those conserved within each genus. (1) Barnase (B. amyloliquefaciens, strain H . IAM 1521); ( 2 ) barnase 1193 ( ( B . amyloliquefaciens, IAM 1193): ( 3 ) barnase 1170 (B. amyloliquefaciens, IAM 1170); ( 4 ) barnase Bci ( B . circulans): (5) binase ( B . inrermedius, strain 7P); ( 6 ) binase Bth ( 3 . rhuringiensis var. subtoxicus) or Beg (3. coagulans, strain BCF256); ( 7 ) RNase Sa (3. aureofaciens, strain BMK): ( 8 ) RNase Sa3 (S. aureofaciens, strain CCM 3239); (9) RNase Sa2 (S. aureofaciens, strain R8126); (10) RNase St (S. eryrhreus, strain NRRL 2338).
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tion and two others. The gene for the former has 7 base substitutions and the latter 19, only two of which coincide. All three amino acid substitutions are on the surface and only position 104 is near the active site. Note that still another amino acid (alanine) occupies this position in binase. For barstar, only strain I A M l l 7 0 differs from strain H, with three substitutions, K1Q, I13M, and R54S. There are 16 additional silent mutations. The enzyme from a strain reported as Bacillus circulans (Fedorova et al., 1994) was found to differ from barnase at only three positions, with substitutions Q15L, G65A, and Q104K (Dementiev et al., 1993a). Note that two of these substitutions are also found in barnase(IAM 1170), so that this enzyme, which might be called barnase Bci, differs from barnase ( I A M l l 7 0 ) only by Q15I and M109I. Binase Bco (Shlyapnikov and Dementiev, 1993) from Bacillus sp. BCF256 (tentatively identified as Bacillus coagulans), isolated from Siberian permafrost, differs from binase only by T106A. Mature binase Bp (Znamenskaya et al., 1995) from Bacillus pumilis strain KMM62 is identical to binase but with 10 silent mutations and three effective mutations in its propeptide. Binase Th (Dementiev et al., 1993b) from Bacillus thuringiensis var. subtoxicus is identical to binase Bco.
B. Streptomyces
It has been shown that the more distantly related extracellular ribonucleases from Streptomyces, with only 20 to 25% sequence identity to barnase but with the same basic fold (Hill et al., 1983), are also inhibited by barstar and protected by it in vivo (Hartley et al., 1996). Of these enzymes, four sequences (Shlyapnikov et al., 1986; Homerova et al., 1992; Nakamura et al., 1982) and two structures (Nakamura et al., 1982; Sevcik et al., 1991) are available. There are 16 rigorously conserved residues across the two genera (Fig. 6), including those known to be most involved in catalysis. With a more liberal definition of conservative substitution, the homology is clear from the sequences alone (Hartley, 1980), and the structures of the central/3 sheet and adjacent active site regions are highly conserved (Hill et aL, 1983). Among themselves, the four Streptomyces enzymes share sequence identities ranging from 49 to 69%. As noted above, these enzymes are all inhibited by barstar. Placed on a plasmid vector also carrying a functional barstar gene, each of their genes has been expressed in E. coli. RNases Sa, Sa2, and Sa3 are produced at
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20-100 mg/liter, RNase St at a much lower level. It is hoped that the recent cloning of Streptomyces barstar homologs (D. Krajcikova, personal communication) will allow large-scale production of RNase St as well. The toxicity of these enzymes is much lower than that of barnase, perhaps because of their disulifide bond and the low oxidation potential of the cell. They can, therefore, be carried alone in E. coli, but, without an inhibitor, only with very low expression. These ribonucleases are now available for the sort of folding studies being carried out on barnase and barstar. Note also that these enzymes form a bridge between barnase and the eukaryotic ribonuclease T1. They are much closer in sequence similarity to each than barnase and T1 are to each other. Parallel studies on all three subfamilies should help us identify the fundamental determinants of the microbial ribonuclease fold. As noted above, inhibitors from Streptomyces should soon be available. With the inhibition of the Sa and St ribonucleases by barstar, this adds still another dimension to the study of these protein-protein reactions. Of evolutionary interest, similarities of sequence have been reported, with varying degrees of conviction, between barnase and bacterial colicins (Lau et al., 1984) and prokaryotic and eukaryotic RNA polymerases (Shirai and Go, 1991) and between barstar and a yeast mitochondrial targeting domain (Pao and Saier, 1994).
IV. O T H E R P R A C T I C A L A P P L I C A T I O N S O F B A R N A S E
A number of workers have taken advantage of the intracellular toxicity of barnase. One approach is to insert the barnase gene into a eukaryotic genome on a promoter that is turned on only in specific tissues or under certain conditions. The first successful application of this was by Mariani et al. (1990), who produced male-sterile tobacco by incorporating the gene on a promoter that is activated only in a pollen-supporting tissue. On the same tobacco promoter the gene has produced male sterility, of wide interest in relation to hybrid seed production, in a number of commercial crops, most notably oilseed rape, but including cereals as well. Furthermore, male-fertile plants carrying both barnase and barstar genes were obtained as progeny of male-sterile plants pollinated by plants carrying the barstar gene on the same promoter (Mariani et al., 1992). With another promoter, Goldman et al. (1994) have investigated
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Robert W. Hartley
the physiology of the floral stigma and incidentally produced female sterility. The barnase and barstar genes are being used, with at least preliminary success (Strittmatter et al., 1995), to increase resistance to Phytophthora infestans in potato plants. The great Irish potato famine of 150 years ago was caused by P. infestans, and in 1995 a new strain became a serious threat to the American potato crop. Natsoulis and Boeke (1991) have suggested an antiviral strategy by successfully countering the action of a viruslike yeast retrotransposon by means of a transposon-induced barnase-capsid protein hybrid. Prior et al. (1991, 1992) showed that a hybrid of barnase and Pseudomonas exotoxin A, produced as a recombinant protein by E. coli, is cytotoxic to murine cells in culture. Although this suggests clinical use of barnase on cell-targeted carriers, the immunogenicity of barnase and the availability of more specific toxins make its practical application in this manner unlikely. Yazynin et al. (1996) have developed an E. coli plasmid cloning vector with positive selection, based on the conditional lethality of a barnase gene carrying an inserted multiple cloning site.
VII. C O N C L U D I N G R E M A R K S
As a simple linear peptide, with an amino acid sequence that determines how, in an aqueous environment, it folds to a specific conformation, barnase has proved an ideal subject. From its study a general outline of its folding pathway has been determined in some detail and considerable information has been accumulated about the energetics of various interactions in the folded protein. Much more of the latter type of information is needed for the design and testing of better potential functions for energy minimization schemes. At present such information is best acquired by protein engineering studies of small proteins such as barnase and barstar and their complex. In the not too distant future better data might come from similar work on even simpler proteins designed for the purpose. There seems to be a growing consensus that most proteins begin their in vitro folding by a rapid collapse to something like Ptitsyn's molten globule state, with a hydrophobic center or centers and most secondary structures formed but absent much native tertiary structure. As the
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general topology of each protein must also be largely established in the molten globule, it is during the initial collapse from a set of r a n d o m conformations that the difficult pathway decisions must be made. It is not certain that a few simple generalizations will help us in predicting such decisions for a particular protein. It must be r e m e m b e r e d that protein folds and folding pathways are products of Darwinian evolution and that what works is not necessarily simple. A clear understanding of the problem will come only with the unraveling and comparing of pathways within and between different protein families. With the start made on barnase and with the availability of r e c o m b i n a n t proteins from three widely spaced subfamilies of microbial ribonucleases, this family seems well placed for comparative folding studies. Because the S t r e p t o m y c e s enzymes have a disulfide bond and RNase T1 has two, a beginning might be to look for conditions under which these proteins, minus their ability to form disulfides, might take up their native folds. Barstar, with study of its folding well u n d e r w a y and with homologs in S t r e p t o m y c e s , represents another family available for such investigations. As a source of barnase expression plasmids, this author is aware of other researchers, mostly developmental biologists and plant virologists, who are attempting to use the lethality of the barnase gene on specific promoters. Indeed, their n u m b e r appears to exceed that of those interested in barnase for itself by a factor of about five.
REFERENCES
Agashe, V. R., and Udgaonkar, J. B. (1995). Thermodynamics of denaturation of barstar: Evidence for cold denaturation and evaluation of the interaction with guanidine hydrochloride. Biochemistry 34, 3286-3299. Agashe, V. R., Shastry, M. C. R., and Udgaonkar, J. B. (1995). Initial hydrophobic collapse in the folding of barstar. Nature (London) 377, 754-757. Arcus, V. L., Vuilleumier, S., Freund, S. M., Bycroft, M., and Fersht, A. R. (1994). Toward solving the folding pathway of barnase: The complete backbone 13C,~SN,and IH NMR assignments of its pH-denatured state. Proc. Natl. Acad. Sci. U.S.A. 91, 9412-9416. Bastyns, K., Froeyer, M., Volckaert, G., and Engelborghs, Y. (1994). The role of Glu-60 in the specificity of the recombinant ribonuclease from Bacillus amyloliquefaciens (barnase) towards dinucleotides, poly (A) and RNA. Biochem. J. 300, 737-742. Baudet, S., and Janin, J. (1991). Crystal structure of a barnase-d(GpC) complex at 1.9 A resolution. J. Mol. Biol. 219, 123-132. Brandts, J. F., Halvorson, H. R., and Brennan, M. (1975). Consideration of the possibility that the slow step in protein denaturation reactions is due to cis-trans isomerism of proline residues. Biochemistry 14, 4953-4963.
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Shastry, M. C. R,, and Udgaonkar, J. B. (1995). The folding mechanism of barstar: Evidence for multiple pathways and multiple intermediates. J. Mol. Biol. 247, 1013-1027. Shastry, M. C. R,, Agashe, V. R., and Udgaonkar, J. B. (1994). Quantitative analysis of the kinetics of denaturation and renaturation of barstar in the folding transition zone. Protein Sci. 3, 1409-1417. Shirai, T., and Go, M. (1991). RNase-like domain in DNA-directed RNA polymerase-II. Proc. Natl. Acad. Sci. U.S.A. 88, 9056-9060. Shlyapnikov, S. V., and Dementiev, A. A. (1993). Amino acid sequence and catalytic properties of the extracellular ribonuclease of Bacillus coagulans. Dokl. Acad. Nauk (Transl.) 332, 150-154. Shlyapnikov, S. V., Both, V., Kulikov, V. A., Dementiev, A. A., Sevcik, J., and Zelinka, J. (1986). Amino acid sequence determination of guanyl-specific ribonuclease Sa from Streptomyces aureofaciens. FEBS Lett. 209, 335-339. Smeaton, J. R., Elliott, W. H., and Coleman, G. (1965). An inhibitor in Bacillus subtilis of its extracellular ribonuclease. Biochem. Biophys. Res. Commun. 18, 36-42. Strittmatter, G., Janssens, J., Opsomer, C., and Batterman, J. (1995). Inhibition of fungal disease development in plants by engineering controlled cell death. Biotechnology, 13, 1085-1089. Stryer, L. (1965). The interaction of naphthalene dye with apomyoglobin and apohemoglobin. A fluorescent probe of nonpolar binding sites. J. Mol. Biol. 13, 482-495. Swaminathan, R., Periasamy, N., Udgaonkar, J. B., and Krishnamoorthy, G. (1994). Molten globule-like conformation of barstar: A study by fluorescence dynamics. J. Phys. Chem. 98, 9270-9278. Udgaonkar, J. B., and Baldwin, R. L. (1990). Early folding intermediate of ribonuclease A. Proc. Natl. Acad. Sci. U.S.A. 87, 8197-8201. Vuilleumier, S., and Fersht, A. R. (1994). Insertion in barnase of a loop sequence from ribonuclease Tl--Investigating sequence and structure alignments by protein engineering. Eur. J. Biochem. 221, 1003-1012. Vuilleumier, S., Sancho, J., Loewenthal, R., and Fersht, A. R. (1993). Circular dichroism studies of barnase and its mutants: Characterization of the contribution of aromatic side chains. Biochemistry 32, 10303-10313. Welker, N. E., and Campbell, L. L. (1967). Comparison of the a-amylase of Bacillus subtilis and B. amyloliquefaciens. J. Bacteriol. 94, 1131-1135. Wintrode, P. L., Griko, Y. V., and Privalov, P. L. (1995). Structural energetics of barstar studied by differential scanning microcalorimetry. Protein Sci. 4, 1528-1534. Yakovlev, G. I., Moiseyev, G. P., Struminskaya, N. K., Romakhina, E. R., Leshchinskaya, I. B., and Hartley, R. W. (1993). Increase of specificity of RNase from Bacillus amyloliquefaciens (barnase) by substitution of Glu for Set57 using site-directed mutagenesis. Eur. J. Biochem. 215, 167-170. Yakovlev, G. I., Moiseyev, G. P., Struminskaya, N. K., Borzykh, O. A., Kipenskaya, L. V., Znamenskaya, L. V., Leschinskaya, I. B., Chernokalskaya, E. B., and Hartley, R. W. (1994). Mutational analysis of the active site of RNase of Bacillus intermedius (Binase). FEBS Lett. 354, 305-306. Yakovlev, G. I., Moiseyev, G. P., Protasevich, I. I., Ranjbar, B., Bocharov, A. L., Kirpichnikov, M. P., Gilli, R. M., Briand, C. M., Hartley, R. W., and Makarov, A. A. (1995). Dissociation constants and thermal stability of complexes of Bacillus intermedius RNase and the protein inhibitor of Bacillus amyloliquefaciens RNase. FEBS Lett. 366, 156-158.
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Yanagawa, H., Yoshida, K., Torigoe, C., Park, J. S., Sato K., Shirai, T., and Go, M. (1993). Protein anatomy: Functional roles of barnase module. J. Biol. Chem. 268, 5861-5865. Yazynin, S. A., Deyev, S. M., Jucovic, M., and Hartley, R. W. (1996). A plasmid vector with positive selection and directional cloning based on a conditionally lethal gene. Gene 169, 131-132. Yoshida, K., Shibata, T., Masai, J., Sato, K., Noguti, T., Go, M., and Yanagawa, H. (1993). Protein anatomy: Spontaneous formation of filamentous helical structures from the N-terminal module of barnase. Biochemistry 32, 2162-2166. Znamenskaya, L. V., Gabdrakhmanova, L. A., Chernokalskaya, E. B., Leshchinskaya, I. B., and Hartley, R. W. (1995). Phosphate regulation of biosynthesis of extracellular RNases of endospore-forming bacteria. FEBS Lett. 357, 16-18.
3 RNase Ta/RNase T2 Family RNases M A S A C H I K A IRIE Department of Microbiology Hoshi College of Pharmacy Tokyo 142, Japan
I. RNase T1 Family RNases A. Structure B. Mechanism of Action II. RNase T2 Family RNases A. Structure B. Identification of Active Site Amino Acid Residues C. Mechanism of Hydrolysis of Nucleotidyl Bonds by RNase Rh D. Base Preference of RNase Rh; Modification by Protein Engineering of RNase Rh References
I. RNase T~ FAMILY RNases
Since the last review of RNase T1 by Takahashi and Moore (1982), many RNases that have specificity and molecular size similar to RNase T1 (RNase T1 family RNases) have been found and their primary structures have been elucidated. RNase T1 is known to be a guanylic acidspecific RNase and its molecular weight is around 11,000. In this review, we will summarize recent progress in this field with special reference to the structure-function relationship of these RNases and their mechanisms of action. RIBONUCLEASES: STRUCTURES AND FUNCTIONS
I01 Copyright 9 1997by Academic Press, Inc. All rights of reproduction in any form reserved.
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Masachika Irie
A . Structure
1. P r i m a r y
Structure
The primary structures of 16 R N a s e s belonging to R N a s e T~ family have been reported. These structures are s u m m a r i z e d in Fig. 1. T h e R N a s e T1 family R N a s e s are divided into two subfamilies according to their sequences. O n e is the R N a s e T1 subfamily (typified by R N a s e T1) and the other is the barnase subfamily. T h e latter includes R N a s e Sa, R N a s e St, R N a s e Bi, and barnase. The subfamily barnase is reviewed in C h a p t e r 2, this volume. The R N a s e s of the R N a s e T~ subfamily are divided into five groups according to the location of the disulfide bond. The most c o m m o n group is r e p r e s e n t e d by R N a s e T~, which has two disulfide bonds, b e t w e e n Cys-2 and Cys-10 and b e t w e e n Cys-6 and Cys103 ( R N a s e T~ numbering). T h e latter disulfide bond was conserved in all known R N a s e T~ family enzymes except for the bacterial R N a s e s (barnase subfamily). In the second group, R N a s e F~ and R N a s e Th~ contain a disulfide group b e t w e e n Cys-24 and Cys-84 in addition to the c o m m o n disulfide bond. The third group, R N a s e U 1, has a disulfide bond b e t w e e n Cys-52/53 (i.e., the inserted amino acid b e t w e e n residues 52 and 53 and Cys-87/88 in addition to the c o m m o n disulfide bond. T h e
Fig. 1. Amino acid sequences of RNase T~ fmaily RNases. Half-cystine residues are expressed by shaded letters. Amino acid residues of the active site are boxed. The box from the 42nd to the 46th residue indicates the base recognition site amino acids. The numbers at the top of the matrix are those of RNase T~ numbering. Pol, RNase from Pleurotus ostreatus (Nomura et al., 1994); U~, RNase from Ustilago sphaerogena (Takahashi and Hashimoto, 1988); U2, RNase from U. sphaerogena (Kanaya and Uchida, 1986); FI, RNase from Fusarium moniliforme (Hirabayashi and Yoshida, 1983); Fll, RNase from Fusarium lateritium (Bezborodova et al., 1988); Thl, Trichoderma harzianum (Polyakov et al., 1988); Ms, RNase from Aspergillus saitoi (Watanabe et al., 1982); T~, RNase from Aspergillus oryzae (Takahashi, 1965); C2, RNase from Aspergillus clavatus (Bezborodova et al., 1983a); Ap~, RNase from Aspergillus pallidus (Bezborodova et al., 1983b); NI, RNase from Neurospora crassa (Takahashi, 1988); Pch~, RNase from Penicillium chrysogenum (Shlyapnikov et al., 1986a); Pbl; RNase from Penicillium brevi-compactum (Shlyapnikov et al., 1984); Sa, RNase from Streptomyces aureofaciens (Shlyapnikov et al., 1986b); St, RNase from Streptomyces erythraeus (Yoshida et al., 1976); Bi, RNase from Bacillus intermedius (Aphanasenko et al., 1979); Ba, Barnase from B. amyloliquefaciens (Hartley and Barker, 1972).
20 ETG
i: ; T h l Ms T 1 C2 A P ~ N1 P c h l P b l Sa
F1 F11 T h l
A p l N1 Pch Pb1 Sa S t Bi
Ba
1
EA AVINTFDGVADYLI AQVINTFDGVADYLQ
-PV---nGPY0
I
E FPLVYNGPYYSSRDNYVSPGP
n
Q T T V E K
YYDYNNY YNNY YNNY YHDY YNNY Q YRNY Q YRNY R YNNY
EGF EGF EGF EGF EGF EGF EGF EGF
T
v
EGI
E FPIKSGG-VYTG----GSPGA E FPIRTGG-VYSG----GSPGA
R R R R R
R R
R R D R D M
u FENR
D V F S N R EGR D IF S N R E G K
80
D R VIYDQ-SGR D R VYYDSNDGT
D D D D D
EAS
H H H H H H H H H
111 I
TPTFF
u2
TDV-TNAIRSAR-TQV-NRAINNAK-D D I - N T A I Q G A L D D VANGDR--PDNYP SQV-RAAANAA QY YQNDDSAGSTTYP SQV-RAAANAA QY Y Q N D D T A G S T T Y P SAV-SAASNAA NYVRAGSTAGGSTYP SDV-SAAKAKGYSL YESDGTI--DDYP SDV-STAQAAGYQL HEDGETVGSNSYP SAV-SDAQSAGYQL ESAGQSVGRSRYP SAV-SDAQSAGYQL YSAGQSVGRSRYP SAI-SAALNKGYSY YEDGATAGSSSYP SAI-SAAQEAGYDL YSANDDV--SNYP SAI-SSAQAAGYNL YSTNDDV--SNYP SAL-PPEATDTLNL IASDGPFPYSQDG ADL-PPEATDTYEL IEKGGPYPYPEDG QASALGWVASKGDLAEVA-P-GKSIGG EAQALGWVASKGNLADVA-P-GKSIGG
A-GVIT .- . . .. - - A - G V I T VVFNDNDELA-GLIT V V F N N N D E L A - G L IT VIFDSHGNL D-MLIT VVFNGNDQL A-GVIT VIFNDDDELA-GVIT II~GEATQE DYYTGD FVVGDGGE- YFYTED LVYSSDWLI I L Y s s D w L IY-KTTD
H H H H H H H H H
TGAPSTNGFVE TGA-SGNNFVQ TGAASYDGFTQ TGA-SG TGA-SG TGA-SG TGA-SG TGA-SG TGA-SG TGA-SG NGA-SG TGA-SG TGA-SG N N F V A Y A T F S L -1 YESFRLT I V N ATFTRIR QTFTKIR
Y-KTTDIHI H
I RF SY S
7
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Masachika Irie
enzyme of the fourth group, RNase U2, has three disulfide bonds: one is the common disulfide bond, one is the same as RNase U1 (Cys-52/ 53-Cys-87/88), and one is between Cys-3 and Cys-52/53. The fifth group, RNase P0, has three disulfide groups: one in common with RNase T1, one the same as the disulfide group of RNase U2, and one different from RNase U2. Among them, RNase represented by RNases U~, U2, and RNase P0 are found in a species of the Basidiomycotina. The other two groups have been found in the Ascomycotina. These RNases have some structural similarity with the four RNases from Streptomyces and Bacillus (barnase subfamily).
2. Amino Acid Residues Responsible for Catalysis The amino acid residues responsible for the enzyme activity of RNase T1 were first determined by Takahashi and colleagues (Takahasi and Moore, 1982) to be two histidine residues, one glutamic acid residue, and one arginine reside. The details are given in their review. Similar experiments were performed for the other RNases, such as RNase St, RNase Sa, and RNase Ms.
3. Three-Dimensional Structure The three-dimensional structures of RNases belonging to the RNase T~ family enzymes were studied by X-ray crystallography and NMR. Rnases already analyzed with and without an inhibitor are RNase T~, RNase U2, RNase F~, RNase FI~, RNase Ms, RNase Pb~, RNase C2, and RNase The. The details of X-ray analysis of RNases will be given in Chapter 10, this volume. For convenience in discussing the mechanism of action, the three-dimensional structure of RNase T~ will be summarized here. RNase T~ consists of an c~ helix (Ser-13-Glu-28) and four antiparallel/~ sheets, /31 (Pro-39-Tyr-42),/32 (Pro-55-Pro-60),/33 (Asp-76-Phe-80), and/34 (Leu-86-Thr-91). The c~ helix is almost perpendicular to the /3 sheets (Sugio et al., 1985a,b; Arni et al., 1988; Koepke et al., 1989). The amino acid residues of the active site components are located on/31 and/34 sheets. The base recognition sites are composed mainly of Asn-43-Glu46 and Tyr-42 and Tyr-45. As shown in Fig. 2a, the base moiety of the 2'-GMP molecule is stacked with Tyr-45 and covered by Tyr-42 and also hydrogen bonded to amide or carbonyl groups of the main chain from Asp-43-Glu-46. The amino group of guanine interacts with the side
105
3 RNase T1/RNase T2 Family 30
70
Lys'41
Tyr.42
,O
~''-'~- - ~ '
=~
Tyr 45
q,
Ns~~p~0~ ~ _
,..,, , ~
PheSO
Gly 4"7
Glu 46
~ Asr,~
-
-.2'GMP
~176176
Arg77
Fig. 2. (a) Stereo view of the RNase T1 main chain (from Sugio et al., 1988). Circles indicate the positions of C c~ carbon atoms. (b) View of the guanine binding site found in the RNase T~-2'-GMP complex; Ca carbon atoms and the main-chain bonds are solid. Bonds of the 2'-GMP molecule are also solid (from Hakoshima et aL, 1988, by permission of Oxford University Press). (c) Stereo view of the active site looking parallel to the guanine base plane (from Sugio et aL, 1988).
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Masachika Irie
chain of Glu-46 and with the carbonyl group of Asn-98; the N-7 nitrogen of the base interacts with the side chain amido of Asn-43 and the C-6 carbonyl interacts with the Asn-44 amido. The loop from Phe-50, Asp49, Phe-48, and Gly-47 forms a network with the side chain of Asn-44 directly or via water, thus helping the fixation of the base moiety (Fig. 2b). This relationship was confirmed by the kinetic studies on the Glu46 and the Asn-98 mutant (Steyeart et al., 1991a). X-Ray crystallographic data showing the mutual relation of the catalytic site amino acids are shown in Fig. 2c. The side chains of Glu-58 and His-40 are close and His-92, Arg-77, and the two groups described above are located in very close proximity to the phosphate atom. The X-ray data of other RNase Tl-inhibitor complexes show that mutual locations of active site amino acid residues are somewhat different in detail, but seem to indicate essentially a similarity. For the subsite structure of RNase T1, Irie, and more precisely Waltz and colleagues (cited by Takahashi and Moore, 1982), presented kinetic evidence of the presence of a subsite (B2 site). Koellner et al. (1991) analyzed the crystal structure of the Y57W mutant of RNase T1 and found that the guanine recognition site was not much altered from the native enzyme, but that the adenine moiety of 2'-AMP interacts with C = O of Gly-74 and also stacks with His-92. The same type of experiment was performed by Hakoshima et al. (1990). Experiments suggesting the location of the B2 site were performed by Steyeart et al. (1991b). They analyzed the kinetic constants of GpN in comparison with those of GpMe with N36A, Y38F, H92Q, and N98A mutants. The enzyme affinity of GpMe was not very much different from those of the GpNs, that is, the contribution of the N moiety to the binding is not large. Among the mutants tested, the enzymatic activities of N36A and N98A against GpN, but not against GpMe, were markedly decreased. Therefore, it was suggested that Asn-36 and Asn-98 are constituents of the B2 site.
B. Mechanism of Action
Takahashi (1970) proposed a mechanism in which Glu-58 and one of two histidine residues, His-40 and His-92, work as a general acid-base catalyst (Fig. 3a), based on the results of chemical modifications. In the first step of the reaction, Glu-58 works as a general base and withdraws the proton from the 2'-OH of guanylic acid, and the proton on His-40
o
a
.D ~ o
|
..
o
e
e o~p/ o~C-- "----" H--N/~~ ..... H._O"" "~OxR
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....
..
b
~
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~
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0 Hisg2
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Hist,0
...... H - - N ~ , , N H
.... ..~\ Gtu58
..... H " H
Fig. 3. (a) A proposed mechanism for RNAse Tt (from Takahashi and Moore, 1982, reprinted with permission). Glu-58 acts as a base and one of the His residues acts as an acid in the catalysis. (b) A mechanism proposed by Nishikawa et al. (1987). His-40 and His-98 work as an acid and a base in the catalysis. Figure 3(b) reprinted with permission from Biochemistry 26:8623, 1987, American Chemical Society.
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Masachika Irie
or His-92 is transferred to the leaving nucleotides, thus working as an acid catalyst. In the second step, His works as a base and activates a water molecule, then Glu works as a proton donor. As discussed previously, X-ray crystallographic data revealed the location of His-40 and His-92. In these data, His-40 is in proximity to Glu58; therefore, the counterpart of Glu-58 in acid-base catalysis might be His-92. In this sense, the Takahashi mechanism is essentially an acid-base catalysis between Glu-58 and His-92. However, Nishikawa et al. (1987) prepared several mutant enzymes for Glu-58, His-40, and His-92 and measured their enzymatic activities against pGpC. The enzymatic activities of E58D, E58Q, E58A, H40A, and H92A were 10, 1, 5, 0.02, and 0% of that of the wild-type enzyme, respectively. E58A had about 5% of the native enzyme activity. Thus they concluded that the two His residues are indispensable for the catalysis, but Glu-58 is not. They also proposed an alternative mechanism, a general acid-base catalysis by two His residues (Fig. 3b). Shibata et al. (1988) reported evidence from the ~H NMR spectrum on the RNase T~ 2'-deoxy-2'-fluoroguanylyl3',5'-uridine complex that His-40 is important for substrate binding. Because Glu-58 and His-40 are very close together it is difficult to conclude which mechanism is the correct one using X-ray crystallography. Grunert et al. (1991) also prepared similar mutant enzymes, E58D, E58Q, H40T, H92F, and H92A, and their enzymatic activities were measured with GpA as a substrate. Kinetic constants, the Vmax/Km of these mutant enzymes, are 10, 7, 2, <0.1, and <0.1%, respectively. These results seem to be very similar to those of Nishikawa et al. (1987), except for H40T. The residual activity of H40T is critical. They also considered the primary structure of bacterial T~-like RNases, such as RNase St and RNase Sa. In these RNases, the His residue corresponding to His-40 is missing. They proposed a mechanism like the Takahashi mechanism for the first step reaction, in which Glu-58 works as a general base and His-40 assists by increasing the basicity of Glu-58. To solve this problem, Steyeart et al. (1990) and Steyeart and Wyns (1993) analyzed the pH profiles of the Vmax/Km Of RNase T1 and Glu-58-Ala RNase T~, and found that four and two ionizable groups participate in the catalysis, respectively; that is, His-92 and His-40 participate in the catalysis in the absence of Glu58. Nonaka et al. (1989,1993) determined the three-dimensional structure of an RNase Ms (an RNase T~ analog)- 2'-deoxy-2'-fluoroguanylyl-3',5'cytidine complex, and the locations of His-40, His-92, and Glu-58 (RNase T~ numbering) were compared with those of the other members of the RNases, such as RNase A, RNase St, and RNase Rh. The mutual location
3
RNase T1/RNase T2 Family
109
of His-92 and Glu-58 is best fitted to His-12-His-119 of RNase A, and His-46-His-109 of RNase Rh, but that of His-92 and His-40 was less well suited. Thus the acid-base catalysis by Glu-58 and His-92 is most possible for RNase T1.
II. RNase T2 FAMILY RNases
RNase T2 is the trivial name of an RNase of Aspergillus oryzae. The enzyme was discovered in the commercial digestive Taka-Diastase and identified as an adenylic acid-preferential RNase by Sato and Egami (1957). Further studies on this enzyme by Egami and collaborators showed that RNase T2 is a glycoprotein releasing 3'-adenylic acid preferentially from yeast R N A and its molecular mass is ~>36,000. The optimal pH of RNase T2 was found to be around pH 4.5. These studies are summarized in "The Enzymes" (Uchida and Egami, 1971) and "Microbial RNases" (Egami and Nakamura, 1969). RNase T2-1ike enzymes are defined as (1) having no absolute base specificity, unlike RNase A (a pyrimidine base-specific RNase) and RNase T1 (a guanine base-specific RNase), (2) having a molecular weight of more than 24,000, and (3) being an acid RNase. RNase T2-1ike RNases from various origins have been reported and we will describe progress in the biochemistry of RNases belonging to RNase T2 family RNases with special reference to the structure-function relationship. In 1989, the primary structures of two RNases, RNase T2 and RNase Rh, were determined by Kawata et al. (1989) and Horiuchi et al. (1989). Since these findings, the primary structures of fungal RNases similar in nature to RNase T2 have been determined. These studies indicated that the RNases of this family contain two typical sequences consisting of 9 and 12 amino acid residues in common (boxed residues in Fig. 4); therefore, enzymes with these common sequences can be classified as RNase T2 family enzymes. As will be discussed later, almost all of the amino acid residues constituting the active site are involved in these two segments, like cassettes. Another typical property of the enzymes of this family is that they are fundamentally base-nonspecific RNases and the molecular mass of their protein moiety is around 24 kDa. They are present various viruses, bacteria, fungi, amebas, plants, and animals. The RNase T2-1ike RNases found in the literature that are well characterized are shown in Table I.
1 Rh T2 Try Irp ATytttt Phyb S2 P1 LE WC ~m 3y Bspl CL1
(~) Rh T2 M Try Irp Le2 Phyb S2 P1 LE MC Dm Oy Bspl CL1
(~) Rh T2 M Try Irp Le2 Phyb S2 P1 LE MC Om Oy Bspl CL1
~
MKAVLALATLIGSTLASSCSST-ALSCSNSAN-SDTCC--SPEYGLVVLNMQ-WA--PGYG-PDN EFPSCPKDIPFSCQNSTAVADSCCFNSP--GGALLQTQFWDTNPPSG-PSD TIDTCSSDSPLSCQTDNE--ASCCFNSP--GGSLLQTQFWOYOPSDG-PSD ASKTCPSNTPLSCHNTTVVQDTCCF-IP--SGQLLQTQFWOTDPSTG-PSD VNSGCGTSGAESCSNSOO--GTCCFEAP--GGLLLQTQFWDTDPSTG-PSD ISSGCGTTGALSCSSNAK--GTCCFEAP--GGLILQTQFWOTSPETG-PTD STE KSTSFDFFIFVTEWNA-SI .... MSKSQLTSVFFILLC-ALSPIYGAFEYMQLVLTWPITFC-----RIKH C---ERT-PT.... KR-PAK .... FRPK-NIC SVFFIFLF-SFSPVYGNFEYLQLVLTWPASFC
PAVC---SF-QKSGSC-FDSFWFVQQW NWDVLIFTOOWPVTTCYHWREENPDQECSLP--Q-KKE KDWNYFTFAQQW~IAVC----AEHK----SCFIP---O--SVVt LEWSKLIMVHHW ATu .... DCQ .... EWSKLYLAHHWPVTu
Im m N N N V IF W S H E W S K H G T C s KSK-OSSLYNSML TYWPSNQG ...... ...... s D D E E IF W E H E W N K H G T C .... ADEDESJl ;WEHEWNKHGTC s TS ILEAQDRTELLSYMKEYWPDYEG NDE S'I FWEHEWGKHGTC I TD I L TAMGADDT LQYMQ TYWK DYQG ...... Q N E Q i I~ S E H E W S K H G T C [ G D L L T A Q G A S D T L D F M N Q Y W V D [ NG . . . . . . .... D G S N E E IL W E H E W A T H G T C [ SSLL TAQGASDTLQFMNQFWLNDPD ST ....... I SDL I DTM-QVWP S F TG ...... O N A S , I; W S H E W S K H G T C TK TKFDSLDKOA IF W K O E Y V K H G T C ~T ....... DGKKKNDLDERWPDL F D E N Y A K Y H O P IL W S Y E Y R K H G M C =K . . . . . . . . DNN I VDYLERHWVQMK S G S G S T IF W S H E W E K H G T C 5Q . . . . . . . I SDL I SSMQQNWP T LAC---P .... A - - N N Q Q ! IF W S H E W T K H G T C rK ....... ISHLQSQLNTLWPNVLR ,K ....... LNPI EDRt. ETFWPOLKG .... MOS TEWll LWKHEWQKHGTC MN . . . . . . . VMPLVPE LKK PYWPNLY .... DTKANS IF W E H E W S K H G T C SNCSLO I= W S H E W K K H G T C 4E . . . . . . . I KOL L POMKMYWPOL LH---P 1H---S S L N R T Q I; W K H E W E K H G T C rE ....... IKDLMSOMRRYWPOV I ~]
SG-LR-O-PPN! O-PPE
I
AFTLHGLWP SWTIHGLWP SWTIHGLWP SWTIHGLWP SWTIHGLWP SWTIHGLWP YFTIHGLWP NFTIHGLWPI NFTIHGLWP' DFGIHGLWP' TFTIHGLWP! FWTIHGIWP GWGIHGLWP' YWTIHGLWP YWTIHGLWP C2
j
DKCSGAYAPSGGCOSNRASSS DNCOGSY-GQF-COKSREYSN DNCOGSY--QEYCOOSREYSN DNCDGSF-PQT-COASRAYTN DNCDGSF-ESS-CDASRAYTG DNCDGSF-SEO-COPSROYTG ENSOGSY-PSG-CSSGK-FST ONHTTML-NY--CORSKPYNM EITGFRL-EF--CTGDPKYET NNNOGTY-PSN-COPNSPYOQ QG-SGTS-LTN-CPQGSPFOZ TKLHOM-GPNF-CNNSANFDP SSOTESKGPEN-CNGSWPFOI OK-SEA ...... CNRSWPFNP OK-GEE ...... CNRTWHFNV
140 120 VSTYOPOC YDNY E E-GED IVDYFQKAMOLRSQYNVYKAFSSNGI T PG X NT I EPSCYKDYS P-QKEVGDYLQKTVDLFKKGLDSYALAKAGIVPD TPS I NT I OPSCYTDYYA-QEEVGDFFQQVVDLFKTLDSYTALSDAGI TPD I TTLDPGCYDDYVP-T EEAADFFSKTVSLFKTLPTYQWLADAGI YSTLETSC L P SGSPKGAEAVAFFQQVVT LFKTLPTYQWLAKAGI TPO YSTLQTSC KPS LPEGSPKGAEAVAFFEQVVTLFKTLPTYEWLTNQGI SG Y . . . . A E-HDF FA TVL S L YDQYNVYKA LDNGG I E PG ....... C SOK F---OR LRDKFDLLSSLRNHGI SRG E--QYFDLAMT ....... C SKI Y .... T PG NQ-KAYFLLATRLKEKFOLLTTLRTHGI ....... A E SV L T---NQ-HA Y F KKALDLKNQI OLLS I LQGAOI H PO S E ST F .... ....... NQ-AAY F K LAVOMRNTLOSYTALRPHAAGPN ........ ILOASOIHPO AMLVEELDNE-LKYFEQGLTWREEYIMSR ........ AT S L PAT S N E-LKY FGMGLKLHAKYN I SR I L V N Q G I L P S ........ EPS AAQLN---SQ-RKYFGKSLDLYKALALTSMLQKLGI ........ KPG AAT LP I LNSQ-KKYFNKTLELYQLVNLNGFLLKAGI I J I C4
G--TYTATEMQSAIESYF-GAKAKIDCSSG-TLSOVALY-FYV-RGR--D-TYV-ITDALS SSKTYKRSEIESALAAIHOGKKPYISCEOG-ALNEZ-WY-FYNIKGNAITGEYQPIDTLTS ..... EDATYKLSOIEDALAAIHOGYPPYVGCEOG-ALSOLYYY-FNV-KGSAIGGTYV-ASERLE GSKSYALDDIOSALSQOH-GAEVTLGC-DGKTLNEV-WYHFNV-KGSLQDGQFV-AAEPOGA ....... SSKTFTLSEITSALKSAA-GVTPALDC-DGKNLNOISWY-FHL-KGSLLDGKFVPIDAPK SSTTHTYSALTAALEAEA-GVIPALNC-OGSOLDEIYWY-FHL-RGSVIDGEFEPISAPE-.... T-FASVGL-C--IT--KNLELROCPONMGS SS-SVSSDSLISVITONI-GGVPVLNC-EG-S .... SR--LRELKEIGI-C--FOE-TVKNVIDCPN-PKT F--SYTVQNLNNTIKAIT-GGFPNLTC T--KHTFGOIQKAIKTVTNOVOPOLKCVEH-IK-GVQELNEIGI-C--FNP-AADNFYPCHH-SYTGE-SYOLVNIRNAIKSAI-GYTPWIQCNVO-QS-GNSQLYQVYI-C--VDG-SGSSLIECPIFPGG GR-TKSRQAIKGFLKAKF-GKFPGLRCRTO-PQTKVSYLVQVVA-C--FAQ-OGSTLIDCTR--OT ~--NNTVAAINNAIvKAL-GKNPSIHCLYD-GKHGIs~LSEIRI-C--FS--KSLELI~CDGIKQGDAvPVGvPGGTIITNCHIGSLvH-YPsLvPPLQ (TAGYMINETEAAVKREL-GVOAVIECVYOKEKTKKOLLYEISI-C--LT--KEFELISCNKKEVS rDHYYQVSDIROALVTVY-KVVPKVQCFLLEKGQEVQLLGQVEL-C--FS--KDLQLONCSHAD . . . . . . . . . . . PK ..... CIEFSFTKELELRTCTE ~TTYYOMAAIKEVLTEFY-GITPKIQCLPPEEGEEAQTLG i J CS
TGSCSGD--VE-YPTK PG-CSTS-GIK-YLPKKSEN DSNCKGS-GIK-YPPKSSS KSTCPDO--VY-YOPKK AGSCASS-GLK-YPPKSG KGDCPSS-GIK-WLPKNNEK FWOCPAK---V----YYRNN -CKPTNKGVMFP CDETD-SKMILFR K--CGTS--IE-FPTF CGAN-FI---F ETTCPRKEPFF-YPPVHON LLVCNDG-PVL-YPPLL EELSVCNOTLPVY-YPSQVK
3
RNase T~/RNase T2 Family
111
A. Structure
1. Primary Structure In 1989, the primary structures of two RNases of this family, RNase T2 (Kawata et aL, 1989) and RNase Rh (Horiuchi et al., 1989), were reported. The structure of the former enzyme was determined by protein sequencing and that of the latter by a combination of protein sequencing and gene cloning. Further sequencing studies of fungal RNases were performed with Aspergillus saitoi (Watanabe et al., 1990), Trichoderma viride (Inada et aL, 1991), Lentinus edodes (Kobyashi et al., 1992), Irpex lacteus (a kind of mushroom) (Watanabe et al., 1995), and Physarum polycephalum (an ameba) (Inokuchi et al., 1993). The two typical active site cassettes were conserved for all enzymes studied. In 1989, McClure et al. found that these two cassettes were also found in self-incompatible factors (s-gene products of Nicotiana alata) and that the s-gene products have RNase activity. On the other hand, Jost et al. (1991) also found that extracellular RNase excreted from cultured tomato cells (Lycopersicon esculentum) has these cassettes. These findings revealed that RNase T2 family RNases are also present in the plant kingdom. Many s-gene products or s-RNase-like RNases have now been found in plants. These RNases will be discussed in Chapters 5 and 6 of this volume. Meador and Kennel (1990) and Favre et al. (1993) reported that the RNases from Escherichia coli and A e r o m o n a s hydrophila, respectively, are RNases of this family. In the animal kingdom, two RNases of this
Fig. 4. Amino acid sequences of RNase T2 family RNases. Rh, RNase from Rhizopus niveus; T2, RNase from Aspergillus oryzae; M, RNase from Aspergillus saitoi; Try, RNase from Trichoderma viride; Le2, RNase from Lentinus edodes; Phyb, RNase form Physarum polycephalum; $2, S-gene product from Nicotiana alata; P1, S-gene product from Petunia inflata; Le, RNase from tomato cells (Lycopersicon esculentum); Mc~, RNase from bitter gourd (Momoridica charantia); Dm, RNase from Drosophila melanogaster; Oy, RNase from oyster (Crassostrea gigas); Bsp~, bovine spleen acid RNase; CI~, chicken liver acid RNase. The numbers at the top of the matrix are those of RNase Rh numbering. Two conserved amino acid sequences characteristic of the RNase T2 family RNases are boxed. C1-C5 are sequences well conserved in plant RNases (Ioerger et al., 1991). Asterisks indigate the amino acid residues involved in the active site.
TABLE I RNases Belonging to the RNase T2 Family Origin a
Aspergillus oryzae (T1) Aspergillus saitoi (M) Trichoderma viride (Trv) Rhizopus niveus (Rh) Lentinus edodes (Le2) lrpex lacteus (Irpl) ( Irp 2) Physarum polycephalum (Phyb) L ycopersicon esculentum (Le) (Lx) Nicotiana alata (s2) Petunia inflata (p2)
Optimal pH 4.5 4.0 4.5 4.5 4.0 4.5 4.5 4.5-4.75 5.6 5.9 7.0
Amino acid residues b or molecular mass 239 238 244 222 239
p, g p p p, g p
260 p 235 p 180 p 205 213 192 199
p, g p g g
Carbohydrate + + +
Specificity~
Refs. d
+
A>G A>G>C>U A>G>U>C A>G>C,U A>G>C>U
1,2 3 4 5 6
+
A>G->C,U
7
+
G>A>U>C
8
G>U=A>C
9 10 11 12
+ +?
1 Po
UI
u2 F i
ii1
T h l
Ms T1
cz A p l N1 P c h l P b l
Sa St
B i Ba
U2 F1 F11 T h l MS T1
C2
A p l N1 P c h l P b l Sa
st
B i Ba
E
10
20
50
TDV-TNAIRSAR-TQV-NRAINNAK-DDI-NTAIQGALDD SQV-RAAANAA QY SQV-RAAANAA QY NY SAV-SAASNAA SDV-SAAKAKGYSL SDV-STAQAAGYQL SAV-SDAQSAGYOL
SF DF DQ DF
I
SAI-SSAQAAGYNL SAL-PPEATDTLNL IASDGPFPYSQDG ADL-PPEATDTYEL IEKGGPYPYPEDG EA Q ASALGWVASKGEL AEVA-P-GKSIGG R V K R L P N D Y I T K S A V I N T F D G V A( D Y L I A Q V I N T F D G V A, D Y L Q T Y H K L P N D Y I T K S E A Q A L G W V A S K G N L A D V A - P - G K S I G G
70 DIRIVIY D Q - S G R T P T F F E F P V F R - G SVYSG----GSPGA DGPYK E Y P L K T S S SGYTG----GSPGA P Y Y S S R D N Y V S P G P I T L GSGPWS E F P L V Y N G E F P I K S G G- V Y T G - - - - G S P G A -PV---DGPYC E F P I R T G G- V Y S G - - - - G S P G A -AV---NGPYC E F P I L S S GKTYTG----GSPGA -KG--LSKPFY -PV---SGTYY E Y P I M S D Y DVYTG----GSPGA S PGA -SV---SSPYY E W P I L S S G D V Y-S G----G E W P I L S S G STYNG----GGPGA -PV---SGNYY A-GLIT -PV---SGNYY E W P I L S S G STYNG----GSPGA A-GLIT D R VIFDSHGNL D-MLIT E F P I L S S GRVYTG----GSPGA -PT---AKPWY -PV---SGTYY D R VVFNGNDQL A-GVIT E F P I L R S GAVYSG----NSPGA D R VIFNDDDELA-GVIT E F P I L K S GKVYTG----SS-GA -PV---SGTYY R R II~GEATQE -PT-QSYGYYH E Y T V I T - - P---GA---RTRGT DYYTGD R R FVVGDGGE- P D - ~ A E G Y Y H E Y T V K T - - P--SG----DDRGA D R LVYSSDWLI -PS-AGSRTWR E A D I N Y - - -V-SG-----FRNA - P G - K S G R T W R ~ EJAD IN Y - - - T - S G - - - - - F R N S U
DF DF DF V T D D
V FQNR E S V-L V F E N R E G I-L
V F S N R EGR -L I ~ F S N R ~ E G-KL
100 TGAPST TGA-SG TGAASY TGA-SG TGA-SG TGA-SG TGA-SG TGA-SG H TGA-SG H TGA-SG H NGA-SG H TGA-SG HTGA-SG H YATFSL
U
TN ST T S
TIVN R R
114
Masachika Irie
family have been found, in oysters (Watanabe et al., 1993) and bullfrog (Yagi et al., 1995). A similar RNase has been found in D r o s o p h i l a m e l a n o g a s t e r (Hime et al., 1995). The primary structures of two animal RNases, bovine spleen RNase and chicken liver RNase, have also been determined (H. Narumi et al., 1995, and T. Uchida et al., 1995, unpublished data, respectively). [Spleen RNase was previously reported by Ohgi et al. (1988) and Hayano et al. (1993).] These RNases are members of the RNase T2 family. The latter enzyme was previously reported by Levy and Karpetzky (1980) and Miura et al. (1984) as a poly(U)-specific RNase. In addition to the above findings, Schneider et al. (1993) found that the surface proteins gp 53 (Collett et al., 1989) (from bovine viral diarrhea virus) and gp 441-48 (Rtimentnapf et al., 1991) (from hog cholera virus) have RNase activity and their sequences indicate that they are RNase T2 family enzymes. RNase T2 RNases are widely distributed in nature and comprise a very large family among the RNase superfamily. The primary structures of all known RNases except for some plant RNases are shown in Fig. 4. Although the primary structures of these RNases include two active center cassettes, there is little structural similarity between bacterial and virus RNases and fungal and plant RNases, except for the cassettes. Judging from the sequences of some plant RNases, they might contain free SH groups. Fungal RNases, such as RNase T2, RNase M, and RNase Rh, contain five disulfide bridges whose locations were determined by X-ray crystallography in the case of RNase Rh to be Cys-3-Cys-20, Cys-10-Cys-53, Cys-89-Cys-120, Cys-63-Cys112, and Cys-182-Cys-213 (Kurihara et al., 1992). The sequence of RNase T2 determined by Kawata et al. (1989) consisted of 239 amino acid residues, and the C-terminal Asn was glycosylated. However, the sequence deduced from the gene structure indicated that 25 amino acid residues were added further at the C terminus. By addition of this C-terminal sequence, it became possible to build up the consensus sequence for N-glycosylation, Asn-X-Thr/Ser (Ozeki et al., 1991). Thus it is quite reasonable that RNase Y2 found in Taka-Diastase is a processed product. Similar C-terminal processing was found in RNases Irpl and Irp2 from Irpex lacteus. The latter is a simple protein consisting of 235 amino acid residues; the former contains 248 amino acid residues, and its C terminus was 13 amino acid residues long. Furthermore, the C terminus of RNase Irpl is O-glycosylated (Watanabe et al., 1995). To understand the physiological role of such processing, further research is needed. By studying all the available sequences of plant RNases, three relatively conserved sections were found in addition to the two cassettes
3
RNase Tl/RNase Tz Family
115
(Ioerger et al., 1991). They are marked C1, C4, and C5 in Fig. 4. C2 and C3 are the same as the two typical cassettes. The C1 locus was relatively conserved in both fungi and plants. When we extend the comparison to other kingdoms, it is interesting that the C1 locus of lower animals, such as oysters and Drosophila melanogaster, is very similar to that of plants and amebas. However, in higher animals the sequence of this locus differs somewhat from that of the lower animals and plants.
2. Three-Dimensional Structure of RNase Tz Family Enzymes The CD spectra of fungal RNases in the shorter wavelength region (200-240 nm) are very similar to each other, indicating similar backbone conformation (Inokuchi et al., 1993; Kobayashi et al., 1992; Ohgi and Irie, 1977). However, the CD spectrum of tomato RNase Le is somewhat different from those of fungal RNases, indicating some sort of change in conformation, probably a less ordered structure than that of fungi (A. LOftier and M. Irie, 1993, unpublished data). X-Ray analysis of the structure of plant RNase is needed to determine the three-dimensional structures of plant RNases. Although most of the RNases of this family are glycoproteins, some of them are simple proteins (Table I). Among these proteins, crystallization has been reported for RNase Rh (Kurihara et al., 1992), RNase MCl (Amitabha De and Funatsu, 1992), and E. coli RNase I (Lim et al., 1993). At present, only the three-dimensional structure of RNase Rh has been elucidated, by Kurihara et al. A stereo view of RNase Rh structure is shown in Fig. 5a. It is an (c~ +/3)-type structure and consists of six a helices and seven/3 strands. They are aA, 72-79; aB, 82-90; c~C, 98-108; c~D, 128-143; c~E, 146-151; c~F, 163-174; 131, 25-33, ~2, 44-52; /33, 138-162; /34, 179-183; /35, 186-197; /36, 201-204; and /37, 217-220. The amino acid residues consisting of the active site are clustered on the/3 strand on three antiparallel strands and also on an c~ helix located parallel to the central three/3 sheets. The locations of these active site components are shown in Fig. 5b. Trp-49 is located in the middle of the active site amino acids, with hydrogen bonding to the carboxyl group of Glu-105 and partial stacking to His-ll9. A water molecule is located in the middle of Glu-105, His104, and His-ll9. As shown in Fig. 5b, the location of this water will be occupied by phosphate groups of nucleotides in the RNase R h - 2 ' - A M P complex (Nakamura et al., 1993; Hamashima, 1994).
116
Masachika Irie
N-te r m i n u s ~ ' ~
N-te rminus~N~
180
C-terminus
~
~
C-terminu~
60
b Asp51 Wat28
-9
Tr
Lysl08
Hisl04
!
":: ,
T
LysI08 His46
~~r
His46
Hisl04
Fig. 5. (a) Three-dimensional structure of RNase Rh (from Kurihara et al., 1992). aCarbon chain of the RNase Rh. (b) Active site of the RNase Rh-2'-AMP complex (from Nakamura et al., 1993; Hamashima, 1994). Figure 5(a) reprinted with permission from Elsevier Science.
3 RNase Tl/RNase T2 Family
117
B. Identification of Active Site A m i n o Acid Residues
1. Chemical Modification Chemical modification of RNase T2 family RNases have been carried out on RNase T2, RNase M, and RNase Rh (Kawata et al., 1991; Irie et al., 1986; Sanda et al., 1985). These three RNases are modified by iodoacetic acid with a concomitant loss in enzymatic activity. The carboxymethylation of RNase T2 was located at His-53 and His-115 (His-46 and His119 in RNase Rh numbering) at a ratio of 40:60 (Kawata et al., 1991). For RNase M, sequences of two peptides containing carboxymethylated histidine were determined (Harada and Irie, 1973) and their locations were identified by elucidation of the complete sequence as His-46-His109 (RNase Rh numbering) at a ratio of 40:60. The site of modification of both histidine residues is the N-3 position and it differs from the case of RNase A (Harada and Irie, 1973; Kawata et aL, 1991). For both RNases, 1 mol of the carboxymethyl group was introduced until complete inactivation occurred, thus carboxymethylation of either one of the two histidines causes inactivation. Photooxidation of RNase M in the presence of methylene blue inactivated the enzyme with loss of about 1 mol of histidine (Irie, 1969b). From these studies and the protein engineering studies described below, it is clear that His-46 and His-119 (in RNase Rh numbering) are crucial for the enzymatic activity of RNase Rh, and the modification of one of them results in loss of activity. RNase M and RNase Rh were inactivated by a water-soluble carbodiimide (Watanabe et al., 1983; Sanda et al., 1985). When RNase Rh was inactivated with ~4C-labeled water-soluble carbodiimide in the presence of the competitive inhibitor cytidine, enzymatic activity decreased to 80% of that of the native enzyme. The RNase was remodified after removal of cytidine. RNase activity was lost with the concomitant incorporation of ~4C-labeled water-soluble carbodiimide. This experiment showed that a carboxyl group is very important for enzymatic activity, but the location of this carboxyl group was not identified, probably because of instability of the modified enzyme. The tryptophan residues of RNase M were resistant to N-bromosuccinimide (NBS) oxidation. However, in acidic medium, where RNase M was partially and reversibly denatured, RNase M was inactivated by oxidation of a single tryptophan residue. This experiment showed that this partially buried tryptophan is very important for enzymatic ac-
118
Masachika Irie
tivity. When a mixture of RNase M and a 20-fold M excess of 2'(3'),5'thioguanosine was excited by light at 360 nm, a new excitation peak appeared at 350 nm. This indicated that at least one tryptophan residue is located near the thioguanosine base (Irie et al., 1972).
2. Kinetic Studies In order to gain insight into the mechanism of RNase T2 family RNases, kinetic studies were performed on RNase Rh and RNase M in an early stage of the research. The pH profiles of the kinetic constants for RNase M indicated that at least four dissociable groups with pKa values of 3, 4.5, 5.8, and 6.7 influenced the activity (Irie, 1969a). Reinvestigation of the pH profile of RNase Rh with UpU as substrate again showed that at least four dissociable groups with pKa values of 3, 4, 5.8, and 6.7 participated in activity. The first two values may be acidic amino acids and the latter two histidine residues. Because the transition at pH 4.0 became unclear in the hydrolysis of UpU by the E105Q mutant, this pKa value might be assigned to El05 of RNase Rh. The assignment of pKa 3.0 is still unknown (Fig. 6) (Irie et al., 1994).
3. Determination of Active Site Amino Acids by Protein Engineering Following the identification of two histidine residues important for enzymatic activity in RNase T2 and RNase M, it was thought that RNase T2 family RNases catalyze RNA hydrolysis by general acid-base catalysis, as does RNase A. Ohgi et al. (1991) found that yeast cells could express the cDNA of RNase Rh protein. They constructed a shuttle vector derived from pYE2211 in which the RNase Rh gene was inserted just after the promoter of D-glyceraldehyde 3-phosphate dehydrogenase with a prepro sequence of R h i z o p u s niveus aspartic protease. The N terminus of the expressed RNase was processed heterogeneously at the pro sequence of aspartic protease. To obtain homogeneous protein, the excreted enzyme was treated with a limited amount of S t a p h y l o c o c c u s aureus V8 protease. By this treatment of RNase Rh with three extra amino acids, Ala-Ser-Gly, at the N terminus (RNase RNAP Rh) was obtained in a yield of 40 mg/liter. The specific activity toward RNA and the CD spectrum of this enzyme were practically the same as those of native RNase Rh. To identify the active site histidine residues Ohgi et al. (1992) compared the amino acid sequences of fungal RNases and found that
3
119
RNase T1/RNase T2 Family o
9
Q
"!
OO0
O0
"!
Og
I
I
log V / K m
9
9 00
O0
Q
99
,,
9 9& , L A
log
V
A A A
A
Inllum
Ii Im um I
nl
A
mmIm him Ii II
AA
~..,',~m 1
I
I
2
3
4
i
4
'
0 0
O0
9 9, I
1
l
Oo
>
l
7
!
Rh
o O O O o
-~ 3 E v o
o
6
"'|
RNAP
0
O 0
,,
5
0
RNAPRhE105Q
2
QQ
o
QOOQ
oo
tOQOI
O
Q
1
O
QaQ QO
9
O
Q
0 I t
2
I
I
I
3
4
5
, I
6
t
o
_
7 pH
Fig. 6. (a) The pH profile of the pgm, log V. . . . and log(Vmax/Km)of RNase Rh toward UpU at 20~ (b) The pH profile of the initial velocity of E105Q toward UpU (40/xM) at 20~ (from Irie et al., 1994).
His-46 and His-109, both of which are carboxymethylated by iodoacetic acid, were conserved completely, and His-104 was also fairly well conserved. Mutant enzymes in which these His residues are replaced by Phe were prepared by Kunkel's method (Kunkel et al., 1987). The enzymatic
120
Masachika Irie
activities of three mutant enzymes, H46F, H104F, and H109F, are summarized in Table II. H46F and H109F were almost completely inactivated, but H104F is about 0.3% as active as the native enzyme. The enzyme activities toward a low molecular weight substrate (ApU) were very similar to those obtained with R N A as substrate. However, the Km value of H104F and the Ki value toward 3'-AMP were increased markedly. Therefore, His-104 was thought to be the binding locus of the phosphate group. The extremely low activities of the H46F and H109F mutants and inactivation of both histidine residues by iodoacetate showed that these two histidine residues are involved in the general acid-base catalysis of RNase Rh. From the 1H NMR of these mutant enzymes and RNase Rh, three histidine residues are easily assigned. By pH titration of the three His peaks, the pKa values His-46, His-104, and His-109 were determined to be 6.7, 5.9, and 6.3, respectively. Titration of His-109 is somewhat unusual. However, the pK~ value of His-109 is slightly lower than that of His-46, and the authors have tentatively assigned His-109 and His-46 as the general base and general acid partners of the catalytic action of RNase Rh. This assignment is also supported
TABLE !! Enzymatic Properties of Mutant Enzymes of RNase Rh" Relative activity (%)
Enzyme RNase Rh H46F H 104F H109F D51N D51E E 105Q E105D K 108R K 108T K108L
Kinetic constant: ApU
RNA
ApU
UpU
Km (M x 1 0 4 )
V. . . . (min i)
100 0.02> 0.3 0.02> 66 19 0.96 1.74 34 7.0 3.1
100 0.19 0.88 0.24 0.19 0.12 1.84 2.01
100
1.10
8130
2.53
56
2.07
122
31 1.0
m
m
n
" At pH 5.0. From Ohgi et al. (1992, 1993a,b).
m
Inhibition constant: 2'-AMP (M x 105) 0.37 0.55 11.0 0.63 5.2 0.23 0.27
3 RNase Tl/RNase T2 Family
121
by the reactivity of the histidine residues toward the iodoacetate of RNase T2 and RNase M. In order to identify the important carboxylic acid in RNase Rh, Ohgi et al. (1993b) compared several fungal RNases and some of selfincompatibility factors of Nicotiana alata with RNase activity. In this comparison the authors found that only two acidic amino acid residues are conserved among them. One was Asp-51 and the other was Glu-105. The substitution of Glu-105 for Gin (E105Q) caused marked inhibition of RNase activity up to 0.96% of the native enzyme. This mutant was also inactive toward dinucleoside phosphate. The pH profile of UpU hydrolysis by E105Q showed that the inactivation was marked in the acidic pH region (Fig. 6b). Therefore this protonated glutamic acid residue might act to polarize the P = O bond or to stabilize the pentacovalent intermediate, as Lyso41 in RNase A does (Douzou and Petzko, 1984; Trautwein et al., 1991). The D51N was fairly active when RNA was used as substrate. However, its activity toward ApU was markedly decreased, but not toward UpU. From this it was deduced that Asp-51 is not related directly to the hydrolysis mechanism, but seems to be related to the adenine base recognition. X-Ray crystallographic data of the RNase R h - 2 ' - A M P complex clearly showed that Asp-51 hydrogen bonded with the amino group of adenine (Fig. 5b) (Nakamura et al., 1993). Lys-108 was involved in the active site cassette. However, in the crystal structure of RNase Rh, the side chain of Lysol08 was directed away from the active site (Kurihara et al., 1992). This residue is conserved relatively well, but in some RNases, such as RNase Le2 (L. edodes) and the P. inflata P2 gene product, they are substituted by Thr and Arg, respectively. In a complex of RNase R h - 2 ' - A M P (Nakamura et al., 1993), and in another type of RNase Rh crystals (T. K. Nakamura and M. Hamashima (1994), unpublished data), the side chain of Lys-108 was located very close to the phosphate group of the inhibitor. Thus, in order to obtain some information on the role of the Lys-108 residue, several mutants at this position were prepared and their enzymatic properties were investigated. The results are shown in Table II. K108R, K108T, and K108L showed 33.5, 7, and 3% activity toward RNA, respectively (Ohgi et al., 1995). From these data, it was thought that a positive charge at this position is preferable, and that a hydroxy group and aliphatic side chain is less preferable for the enzymatic activity. Therefore, Lys
122
Masachika Irie
is not crucial, but is preferable for enzymatic activity. Possibly it polarizes the P = O bond or stabilizes the intermediate, as does Glu-105.
C. Mechanism of Hydrolysis of Nucleotidyl Bonds by RNase Rh From the results of chemical modification, protein engineering studies, and X-ray crystallography, a tentative mechanism for the action of RNase Rh (shown schematically in Fig. 7) was proposed by Irie and associates (Ohgi et aL, 1993a; Irie et al., 1994). His-46 and His-109 participate in general acid-base catalysis, and thus His-109 withdraws the hydrogen from the 2'-OH of the ribose moiety and forces it to attack the positively polarized P atom. His-46 might be a hydrogen donor to the leaving nucleoside. His-104 is the anion-binding site of the phosphate group and probably has some role in polarizing the phosphate moiety. Glu-105 also acts to polarize the P----O bond or to stabilize the pentacovalent intermediate. Lys-108 should also have some role in the activation step, but not in binding of the substrate. The contribution of Lys-108 to catalysis may be less than that of Glu-105. Asp-51 recognizes the amino group of adenine (it constitutes the so-called B1 site). The results of X-ray crystallography indicated that Trp-49 is partially buried in the molecule, but stacks with the base moiety of the nucleotide. Tyr-57, which is located on the surface of the enzyme, also stacks with the base moiety of the nucleotide (Nakamura et al., 1993). Therefore, these three groups constitute the B1 base recognition site. However, modification of amino acid residues of the base recognition site (vide infra) has a marked effect on enzymatic activity. This is probably due to the fact that these amino acids are located so close together, both spatially and in the primary sequence, and a change in the base recognition site markedly affects the mutual location of each component and hence, the efficiency of hydrolysis.
D. Base Preference of RNase Rh; Modification by Protein Engineering As shown in Table I, although RNase T2 family RNases are basenonspecific, the release of mononucleotides from RNA, the rates of
A
i --CH2"- CXO ~o H
Asp51
l_c.__~ ~ ~' \oH
NH 2
NH 2
\o- ~o<"~" I~.-~ l_c.~_c.~-c-, ,~ a
Olul05
OH
~. ~+ +~,,,, --0-"- ..........HN NH
~___~ ........
!_CH"2
His46
R2 - H
O=
"2 t 239
........"'-c~ ....~"./..
H. + ~
c.2-[
HIs 104 )H
H
Asp51
oo --C H 2--C'~OH
Fig. 7. Possible mechanism of action of RNase Rh. In this mechanism, Glu-105 and Lys-108 may work to polarize the P = O bond or stabilize the pentacovalent intermediate in the transition state. R1, R2, and R3 are His-104, Glu-104, or Lys-108 (from Ohgi et al., 1995).
:H
HIsI09
124
Masachika Irie
hydrolysis of four homopolynucleotides, and the rates of hydrolysis of dinucleoside phosphates by RNases, all showed unique base specificity. They are classified as (1) adenylic acid preferential, like many fungal RNases, (2) guanylic acid preferential, like RNase Le (tomato) and RNase phyb (ameba), and (3) uridylic acid preferential, like chicken liver RNase (Levy and Karpetzky, 1980; Miura et al., 1984). Precise principles to regulate these base specificities are not yet known. A few trials to find an empirical rule for base recognition were made by Ohgi et al. (1993b). As stated before, mutant D51N seems to lose adenylic acid preference and acquire guanylic acid preference. In the sequences in Fig. 1, position 51 was sometimes substituted by Glu, Asn, and Thr. The base specificities of such enzymes found in natural sources were mostly guanylic acid preferential. K. Ohgi, M. Takeuchi, M. Iwama, M. Irie (1995) prepared several mutant enzymes at this position and checked their base specificity (unpublished results). D51E and D51N are guanylic acid-preferential enzymes. The base specificity of D51Q is slightly different in that it is cytidylic acid preferential. Ohgi et al. (1993b) also prepared a mutant at Trp-49 and this mutant was markedly inactivated, indicating the importance of this Trp. However, although W49F was about 16% active, the rates of hydrolysis of CpG and UpG remained unchanged and those for ApG and GpG decreased, and it became more pyrimidine base-preferential than RNase Rh. The replacement of Tyr-57 by Phe, Trp, and Leu causes another modification in base specificity. Y57F enzymes are adenylic acid preferential, as is the wild type, but Y57W is more purine base preferential, probably due to the strong stacking of the Trp ring with the purine base. These experiments showed that we can modify the base specificity of RNases appreciably. Further studies will be necessary to establish what governs the base specificity or to obtain an empirical rule.
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3 RNase T~/RNase T2 Family
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Aphanasenko, G. A., Duddkin, S. M., Kaminir, L. B., Leshchinskaya, I. B., and Severin, E. S. (1979). Primary structure of ribonuclease from Bacillus intermedius. FEBS Lett. 97, 77-80. Arni, R., Heinemann, U., Tokuoka, R., and Saenger, W. (1988). Three-dimensional structure of the ribonuclease T~-2'-GMP complex at 1.9 ,~ resolution. J. Biol. Chem. 263, 15358-15368. Bezborodova, S. I., Khodova, O. M., and Stepanov, V. M. (1983a). The complete amino acid sequence of ribonuclease C2 from Aspergillus clavatus. FEBS Lett. 159, 256-258. Bezborodova, S. I., Ermkbaeba, L. A., Shlyapnikov, S. V., Polyakov, F. M., and Bezbordova, A. M. (1983b). Ribonuclease Ap~ of Aspergillus pallidus. Purification, determination of the primary structure, and crystallization. Biokhimia 53, 965-973. Bezobordova, S. I., Chepurnova, N. K., and Shlyapnikov, S. V. (1988). Ribonuclease FI, from Fusarium lateritium. Isolation, substrate specificity and amino acid sequence. Bioorg. Khim. 14, 893-904. Collett, M. S., Larson, R., Gold, C., Strick, D., Anderson, D. K., and Purchio, A. F. (1989). Molecular cloning and nucleotide sequence of the pestivirus bovine viral diarrhea virus. Virology 165, 191-199. Douzou, P., and Petzko, G. A. (1984). Stop action pictures at sub-zero temperature. Adv. Protein Chem. 36, 245-361. Egami, F., and Nakamura, K. (1969). "Microbial Ribonucleases." Springer-Verlag, Berlin. Favre, D., Ngai, P. K., and Timmis, K. N. (1993). Relatedness of a periplasmic broadspecificity RNase from Aeromonas hydrophila to RNase I of Eschrichia coli and to a family of eucaryotic RNase. J. Bacteriol. 175, 3710-3722. Grunert, H., Zouni, A., Beinecke, M., Quaas, R., Georgalis, Y., Saenger, W., and Hahn, U. (1991). Studies on RNase T~ mutants affecting enzyme catalysis. Eur. J. Biochem. 19"/, 203-207. Hakoshima, T., Toda, S., Sugio, S., Tomita, K.-I., Nishikawa, S., Morioka, H., Fuchimura, S., Kimura, T., Uesugi, S., Ohtsuka, E., and Ikehara, M. (1988). Conformational properties of the guanine-binding site of ribonuclease T~ inferred from X-ray structure and protein engineering. Protein Eng. 2, 55-61. Hakoshima, T., Itoh, T., Tomita, K.-I., Nishikawa, S., Morioka, H., Uesugi, S., Ohtsuka, E., and Ikehara, M. (1990). Crystallization and preliminary X-ray investigation of nonspecific complexes of a mutant ribonuclease T~(Y45W) with 2'-AMP and 2'-UMP. J. Mol. Biol. 216, 497-499. Hamashima, M. (1994). Ms. Thesis, Nagaoka University of Technology, Nagaoka, Japan. Harada, M., and Irie, M. (1973). Alkylation of a ribonuclease from Aspergillus saitoi with iodoacetate and iodoacetamide. J. Biochem. (Tokyo) "/3, 705-710. Hartley, R. W., and Barker, E. A. (1972). Amino acid sequence of extracellular ribonuclease (Barnase) of Bacillus amyloliquefaciens. Nature (London) New Biol. 235, 15-16. Hayano, K., Iwama, M., Sakamoto, H., Watanabe, H., Sanda, A., Ohgi, K., and Irie, M. (1993). Characterization of poly C preferential ribonuclease from chicken liver. J. Biochem. (Tokyo) 114, 156-162. Hime, G., Prior, L., and Saint, R. (1995). The Drosophila melanogaster genome contains a member of the Rh/T2/S-glycoprotein family of ribonuclease-encoding genes. Gene 158, 203-207. Hirabayashi, J., and Yoshida, H. (1983). The primary structure of ribonuclease F~ from Fusarium moniliforme. Biochem. Int. 7, 255-262.
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Horiuchi, H., Yanai, K., Takagi, M., Yano, K., Wakabayashi, E., Sanda, A., Mine, S., Ohgi, K., and Irie, M. (1989). Primary structure of a base nonspecific ribonuclease from Rhizopus niveus. J. Biochem. (Tokyo) 103, 408-418. Ide, H., Kimura, M., Arai, M., and Funatsu, G. (1991). The complete amino acid sequence of ribonuclease from the seeds of bitter gourd (Momoridica charantia. FEBS Lett. 284, 161-164. Inada, Y., Watanabe, H., Ohgi, K., and Irie, M. (1991). Isolation, characterization and primary structure of a base nonspecific and adenylic acid preferential ribonuclease with higher specific activity from Trichoderma viride. J. Biochem. (Tokyo) 110, 896-901. Inokuchi, N., Koyama, T., Sawada, F., and Irie, M. (1993). Purification, some properties and primary structure of base nonspecific ribonuclease from Physarum polycephalum. J. Biochem. (Tokyo) 113, 425-432. Ioerger, T. R., Gohike, J. R., Xu, B., and Kao, T.-H. (1991). Primary structural features of the self-incompatibility protein of Solanaceae. Sex Plant Reprod. 4, 81-87. Irie, M. (1969a). pH-Profile of the kinetic parameters of ribonuclease from Aspergillus saitoi. J. Biochem. (Tokyo) 55, 133-140. Irie, M. (1969b). Photooxidation of ribonuclease from Aspergillus saitoi. J. Biochem. (Tokyo) 66, 565-572. Irie, M., Harada, M., and Sawada, F. (1972). Studies on the state of tryptophan residues in ribonuclease from Aspergillus saitoi. J. Biochem. (Tokyo) 72, 1351-1359. Irie, M., Watanabe, H., Ohgi, K., and Harada, M. (1986). Site of alkylation of the major ribonuclease from Aspergillus saitoi with iodoacetate. J. Biochem. (Tokyo) 99, 527-533. Irie, M., Ohgi, K., Watanabe, H., lwama, M., Nakamura, T. K., Kurihara, H., Nonaka, T., Mitsui, Y., Horiuchi, H., and Takagi, M. (1994). pH-Profile of kinetic constants of RNase Rh from Rhizopus niveus and its mutant enzymes towards UpU and possible mechanisms of RNase Rh. J. Biochem. (Tokyo) 115, 1083-1087. Jost, W., Bak, H., Glund, K., Terpstra, P., and Beintema, J. J. (1991). Amino acid sequence of an extra-cellular, phosphate starvation induced ribonuclease from cultured tomato (Lycopersicon esculentum). Eur. J. Biochem. 198, 1-6. Kanaya, S., and Uchida, T. (1986). Comparison of the primary structure of ribonuclease U2 isoform. Biochem. J. 260, 164-170. Kawata, Y., Sakiyama, F., and Tamaoki, H. (1989). Amino acid sequence of ribonuclease from Aspergillus oryzae. Eur. J. Biochem. 176, 683-697. Kawata, Y., Sakiyama, F., Hayashi, F., and Kyogoku, Y. (1991). Identification of two essential histidine residues of ribonuclease T2. Eur. J. Biochem. 187, 255-262. Kobayashi, H., Inokuchi, N., Koyama, T., Watanabe, H., Iwama, M., Ohgi, K., and Irie, M. (1992). Primary structure of a base nonspecific and adenylic acid preferential ribonuclease from the fruit bodies of Lentinus edodes. Biosci. Biotechnol. Biochem. 55, 2003-2010. Koellner, G., Grunert, H. P., Landt, O., and Saenger, W. (1991). Crystal structure of the Tyr45Trp mutant of ribonuclease TI in a complex with 2'-adenylic acid. Eur. J. Bioo chem. 201, 199-202. Koepke, J., Maslowska, M., Heinemann, U., and Saenger, W. (1989). Three dimensional structure of ribonuclease T~ complexed with guanylyl-2',5'-guanosine at 1.8 ,~ resolution. J. Mol. Biol. 206, 475-488. Kunkel, T. A., Robert, J. D., and Zakcus, R. A. (1987). Rapid and efficient site-specific mutagenesis without phenotypic selection. In "Methods in Enzymology" (R. Wu, and L. Grossman, eds.), Vol. 154, pp. 415-423 Academic Press, Orlando, Florida.
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Kurihara, H., Mitsui, Y., Ohgi, K., Irie, M., Mizuno, H., and Nakamura, K. T. (1992). Crystal and molecular structure of RNase Rh, a new class of microbial ribonuclease from Rhizopus niveus. FEBS Lett. 306, 189-192. Levy, C. C., and Karpetzky, T. P. (1980). The purification and properties of chicken liver RNase. An enzyme which is useful in distinguishing between cytidylic and uridylic acid. J. Biol. Chem. 255, 2153-2159. Lim, L. W., Mathur, S., Canistrano, V. J., and Kennel, D. (1993). Preliminary X-ray crystallographic studies of ribonuclease I from Escherichia coli. J. Mol. Biol. 234, 499-500. L6ffler, A., Glund, K., and Irie, M. (1993). Amino acid sequence of an intracellular ribonuclease from cultured tomato (Lycopersicon esculentum) cells. Eur. J. Biochem. 714, 627-633. McClure, B. A., Haring, V., Ebert, P. R., Anderson, M. A., Simpson, R. J., Sakiyama, F., and Clarke, A. E. (1989). Style self-incompatibility gene products of Nicotiana alata as ribonuclease. Nature (London) 342, 955-958. Meador III, J., and Kennel, D. (1990). Cloning and sequencing the gene encoding Escherichia coli ribonuclease I exact physical mapping using the genomic library. Gene 95, 1-7. Miura, K., Inoue, Y., Hashimoto, Y., Inoue, A., and Ueda, T. (1984). Purification of chicken liver ribonuclease by affinity chromatography with UMP-Sepharose CL. Chem. Pharm. Bull. 32, 4054-4060. Nakamura, T. K., Ishikawa, N., Hamashima, M., Kurihara, H., Nonaka, T., Mitsui, Y., Ohgi, K., and Irie, M. (1993). Protein nucleic acid recognition in ribonuclease Rh-2'adenylic acid complex. Third International Meeting, Ribonucleases, chemistry, biology, biotechnology p. 5. Narumi, H., Ogawa, Y., lwama, M., Kusano, A., Sanda, A., Ohgi, K., Irie, M. (1995). Unpublished data. Narumi, H., Ogawa, Y., Uchida, T., Watanabe, H., Iwama, M., Sanda, A., Ohgi, K., and Irie, M. (1995). The primary structure of bovine spleen acid ribonuclease. Abstracts of the Japanese Biochemical Society (September, Sendai). Nishikawa, S., Morioka, H., Kim, H.-J., Fuchimura, K., Tanaka, T., Uesugi, S., Hakoshima, T., Tomita, K.-I., Ohtsuka, E., and Ikehara, M. (1987). Two histidine residues are essential for ribonuclease Tt activity as is the case of ribonuclease A. Biochemistry 26, 8620-8624. Nomura, H., Inokuchi, N., Kobyashi, H., Koyama, T., Iwama, M., Ohgi, K., and Irie, M. (1994). Purification and primary structure of a new guanylic acid specific ribonuclease from Pleurotus oseteatus J. Biochem. (Tokyo) 116, 26-33. Nonaka, T., Mitsui, Y., Watanabe, H., Ohgi, K., and Irie, M. (1989). Three dimensional structure of ribonuclease Ms-3'-guanylic acid at 2.5 ,A resolution. FEBS Lett. 283, 207-209. Nonaka, T., Nakamura, K. T., Uesugi, S., Ikehara, M., and Irie, M. (1993). Crystal structure of ribonuclease Ms (as a ribonuclease T~ homologue) complexed with a guanylyl(3',5')-cytidine analogue. Biochemistry 32, 11825-11837. Ohgi, K., and Irie, M. (1977). Circular dichroism studies on the N-bromosuccinimide oxidation of ribonuclease from Aspergillus saitoi. J. Biochem. (Tokyo) 81, 1031-1039. Ohgi, K., Sanda, A., Takizawa, Y., and Irie, M. (1988). Purification of acid ribonuclease from bovine spleen. J. Biochem. (Tokyo) 103, 267-273. Ohgi, K., Horiuchi, H., Watanabe, H., Takagi, M., Yano, K., and Irie, M. (1991 ). Expression of RNase Rh from Rhizopus niveus in yeast and characterization of the secreted proteins. J. Biochem. (Tokyo) 109, 776-785.
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Watanabe, H., Hamid Fauzi, Iwama, M., Onda, T., Ohgi, K., and Irie, M. (1995). Primary structure of RNases from Irpex lacteus. Biosci. Biotechnol. Biochem. 59, 2092-2103. Yagi, H., Kobayashi, H., Inokuchi, N., Koyama, T., and Irie, M. (1995). Purification and properties of a base nonspecific acid ribonuclease from bullfrog (Rana catesbeiana). Biol. Pharm. Bull 18, 219-223. Yoshida, N., Sasaki, A., Rashid, A., and Ohtsuka, H. (1976). The amino acid sequence of ribonuclease St. FEBS Lett. 64, 122-125.
4 Structure and Mechanism of Action of the Cytotoxic Ribonuclease c -Sarcin IRA G. WOOL Department of Biochemistry and Molecular Biology The University of Chicago Chicago, Illinois 60637
I. II. III. IV. V. VI. VII. VIII. IX. X. XI. XII.
Discovery of a-Sarcin and Determination of the Basis of Its Cytotoxicity Substrate Specificity of a-Sarcin a-Sarcin Cleavage Site Sequence Structure of a-Sarcin and of Related Aspergillus Toxins Mechanism by Which ot-Sarcin Enters Cells a-Sarcin Recognition Elements in 28S r R N A Conformation of the Sarcin Domain in 28S r R N A Effect of Mutations of the Analog of G4319 in 28S r R N A on Recognition of Oligoribonucleotides by ct-Sarcin Phenotype of Mutations of G2655 in Escherichia coli 23S r R N A Relationship of the ot-Sarcin R N A Identity Element to Selection by the Toxin of the Unique Site of Phosphodiester Bond Hydrolysis Three-Dimensional Structure of Restrictocin, a Homolog of ct-Sarcin. Binding to a-Sarcin Domain R N A and the Catalytic Mechanism Coda References
I. D I S C O V E R Y OF a - S A R C I N A N D D E T E R M I N A T I O N OF T H E BASIS OF ITS C Y T O T O X I C I T Y
In 1956 Birger Olson and colleagues were asked to expand the scope of the antibiotic screening program in the Michigan Department of RIBONUCLEASES: STRUCTURES AND FUNCTIONS
131 Copyright 9 1997by AcademicPress, Inc. All rights of reproduction in any form reserved.
132
Ira G. Wool
Health to include a search for anticancer agents. Soon afterward they obtained a sample of soil from a Michigan farm and isolated from it the mold Aspergillus giganteus (MDH18894) (Olson and Goerner, 1965; Olson et aL, 1965). The mold was found to produce a substance that was marvelously effective in inhibiting the growth of tumors in rats, particularly of sarcoma 180 and carcinoma 755. Birger Olson christened the basic protein, a-sarcin, for antisarcoma, a-Sarcin had a clinical trial, but sad to say it proved to be insufficiently effective and toxic. For more than a decade a-sarcin languished. Its renaissance is an oft-told tale of a potential therapeutic agent whose discovery created great excitement and commensurate expectations, that failed in the clinic, and was buried in disappointment, only to be resurrected by biochemists and molecular biologists because of its utility in analyzing the structure and function of ribosomes and of ribonucleoprotein complexes. Vazquez (1979; Fernandez-Puentes and Vasquez, 1977; Conde et al., 1978) and Cundliffe (Hobden and Cundliffe, 1978), and associates, established that the mechanism of the cytotoxicity of a-sarcin, and of the related Aspergillus toxins restrictocin and mitogillin, was an inhibition of protein synthesis, and that this inhibition was, in turn, the result of an effect on ribosomes. Then in the most important single experiment, Schindler and Davies (1977) showed that a-sarcin inactivated ribosomes by cleaving a fragment, called the a fragment, from the RNA in the large subunit of yeast ribosomes. It was established that only a single fragment was produced by the toxin and that the fragment was probably derived from the 3' end of 28S rRNA (Schindler and Davies, 1977). Restrictocin and mitogillin have the same mechanism of action as asarcin (Fernandez-Luna et al., 1985; Fando et al., 1985). Moreover, the primary structure of restrictocin differs from that of mitogillin by only one residue, and they share 86% amino acid sequence identity with a-sarcin.
II. S U B S T R A T E SPECIFICITY OF a - S A R C I N
When ribosomes are the substrate, the cleavage of RNA by a-sarcin is remarkably specific (Endo and Wool, 1982; Endo et al., 1983). Not only are 5S, 5.8S, and 18S rRNAs spared, but only one phosphodiester bond in 28S rRNA is hydrolyzed (Endo et al., 1983). The specificity cannot be attributed to the unavailability of other phosphodiester bonds
4
Structure and Mechanism of a-Sarcin
133
because, if ribosomes are treated with other ribonucleases, all species of rRNA are progressively digested and no specific oligoribonucleotides are formed (Endo and Wool, 1982). The c~ fragment can be generated from 60S ribosomal subunits as well as from 80S ribosomes but, of course, not from 40S particles. Ribosomes are extremely sensitive to the toxin. A concentration as low as 3 x 10 -8 M will produce the a fragment (Endo et al., 1983). The Vmax for a-sarcin (at 3 x 10 -8 M) with ribosomes is 1.6/xM/min; the Km is 5.3/zM and the kcat is 55 min -1. What is striking about c~-sarcin is that when the substrate is RNA rather than ribosomes, and the concentration of the toxin is higher, its specificity is entirely different, albeit still unusual. If, for example, the substrate is free 28S rRNA, the toxin causes extensive digestion of the nucleic acid (Endo et aL, 1983). It is important that at lower, cytotoxic, concentrations c~-sarcin retains its specificity for the single phosphodiester bond even in free 28S rRNA (Miller and Bodley, 1988). Experiments with homopolymers established that a-sarcin action is preferentially on purines in RNA; experiments with 5S rRNA revealed a most unusual property of the enzymemthat a-sarcin cuts on the 3' side of nearly every adenine and guanine without regard to whether the nucleotide is in a single- or double-stranded region. Hydrolysis by a-sarcin of RNA substrates requires neither monovalent nor divalent cations; divalent cations are inhibitory at any concentration whereas monovalent cations are only inhibitory when the concentration exceeds 0.1 M. Catalysis generates a 3'-phosphate and a 5'-hydroxyl at the site of cleavage (Endo and Wool, 1982). The reaction is not markedly affected by pH but the enzyme is most active at pH 7. a-Sarcin, like many other nucleases, hydrolyzes both RNA and DNA. Digestion of DNA, however, requires about two orders of magnitude more of the toxin for hydrolysis of 50% of the substrate (Endo et al., 1983). Moreover, cleavage of DNA requires magnesium whereas hydrolysis of RNA is inhibited by the cation. Finally, when D N A is the substrate, both purines and pyrimidines are attacked.
III. a-SARCIN CLEAVAGE SITE SEQUENCE
The site of cleavage by c~-sarcin is on the 3' side of the guanosine at position 4325 in 28S rRNA (Fig. 1) (Chan et al., 1983). G4325 is embedded in a purine-rich stretch of 12 nucleotides that is near universal in the
134
Ira G. W o o l
U--A C.-O C--G OAc -'GAu A CGGA / AUCAGCGGGGAAA IIIII .III II oUAOUC AucccA U~
ACUGGcUU cAAGCG GUGGcGGc Uu .III.I C U CGCUGc A
~uuuUU
A
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/
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'
c C
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AGeA
Fig. 1. The a-sarcin domain of 28S rRNA. A portion of rat 28S r R N A with the asarcin domain" the sites of a-sarcin and ricin action are indicated and the stretch of 12 bases that is near universal is underlined.
R N A of cytoplasmic ribosomes. Indeed, this is the longest consecutive sequence of conserved nucleotides in rRNA. It should be noted that c~sarcin has neither antimicrobial nor antifungal activity because it enters neither bacterial nor fungal cells; however, isolated yeast and Escherichia coli ribosomes are sensitive to the toxin. Indeed, all ribosomes that have been tested, including those of the producing organism, are sensitive to the toxin. Obviously, the ribosomes of the producing organism are not inherently resistant to the toxin. How then does the Aspergillus survive the synthesis
4
Structure and Mechanism of a-Sarcin
135
of the toxin? a-Sarcin is synthesized as a protoxin with a 27-aminoacid signal sequence (Lamy and Davies, 1991) that is removed during translation. Prepro-ct-sarcin has a molecular weight of 22,5000 and is processed in the endoplasmic reticulum during translation to the 18,500 molecular weight pro-a-sarcin, which is further processed to the mature form that is packaged into vacuoles for secretion (Endo et al., 1993a,b). Thus protection of the producing organism is the reflection of the biochemistry of the secretory system, which confines the toxin to the endoplasmic reticulum before its synthesis is complete; the endoplasmic reticulum (like the digestive tract) is physiologically outside the cell and the mature toxin cannot regain entry to the cell because the cell membrane of Aspergillus, like that of other fungi, is not permeable to the toxin. In depictions of the secondary structure of the large ribosomal subunit RNA (Wool, 1986; Gutell and Fox, 1988), the a-sarcin cleavage site is in a single-stranded loop (Fig. 1) and the site must, of course, be accessible on the surface of the ribosome. Moreover, the domain was deemed from the start to be important for the function of the ribosomemfirst, because the nucleotide sequence is conserved. The rule for rRNAs is conservation of secondary structure, not primary sequence, so retention of the a-sarcin sequence is impressive. This is not to say that there are no other nucleotide sequences in rRNA that are conserved; there most certainly are, and most of these sequences are important for function. Second, it is surprising that cleavage of a single phosphodiester bond inactivates the ribosome because ribosomes survive mild treatment with nucleases despite many nicks in the rRNA; indeed, intact rRNA is not essential for protein synthesis. Until recently little was known of the function of individual ribosomal components or even of ribosomal domains. As a rule, neither the ribosomal proteins nor the nucleic acids have activity when separated from the particle. In the beginning it was thought that the rRNAs only provided a scaffolding to support the ribosomal proteins that catalyzed the partial reactions of protein synthesis. However, the pendulum has swung in the other direction. Now the rRNAs are envisioned as being responsible for the basic biochemistry of protein synthesis: for the binding of aminoacyl-tRNA, mRNA, and the initiation, elongation, and termination factors; for peptide bond formation; and for translocation. The ribosomal proteins, which are presumed to be a later evolutionary embellishment, are though to facilitate the folding and the maintenance of an optimal configuration of the rRNA, perhaps, in this way conferring on protein synthesis speed and accuracy (Stern et al., 1989). The value that derives from an analysis of the mechanism of action of antibiotics and of toxins that affect ribosomes is in concentrating
136
Ira G. Wool
attention on regions where efforts to comprehend functional correlates of structure are likely to be rewarded. The catastrophic effect of a-sarcin on protein synthesis is ample evidence that the sarcin domain is crucial for ribosome function. This is almost certainly because the domain is involved in EF-1 (or EF-Tu)-dependent binding of aminoacyl-tRNA to ribosomes and EF-2 (or EF-G)-catalyzed GTP hydrolysis and translocation. The conclusion comes from a series of compelling observations: first, that these are the partial reactions of protein synthesis that are most adversely affected by a-sarcin (Vazquez, 1979); and, second, that cleavage at the a-sarcin site in E. coli 23S rRNA interferes solely with the binding of EF-Tu and EF-G (Hausner et al., 1987). Moreover, EFTu and EF-G footprint in the a-sarcin domain (Moazed et al., 1988). In E. coli 23S rRNA EF-Tu protects four nucleotides against chemical modification and these correspond in eukaryotic 28S rRNA to G4319, A4324, G4325, and A4329; G4325 is at the a-sarcin cut site and the other three are in the universal sequence. EF-G also protects four nucleotides: three are the same as the ones protected by EF-Tu, they correspond to G4319, A4324, and G4325.
IV. S T R U C T U R E OF ct-SARCIN A N D OF R E L A T E D Aspergillus TOXINS
The specificity of a-sarcin is likely to be a reflection of the structure of the toxin and of a domain in the ribosome. The protein, whose amino acid sequence has been determined (Sacco et al., 1983), has 150 residues and a molecular weight of 16,987 (Fig. 2). The toxin is very basic, having 20 lysyl, four arginyl, and eight histidyl residues; there are four cysteines and they are linked by disulfide bridges between the residues at positions 6 and 148 and 76 and 132. c~-Sarcin, apart from the disulfide bridges, at least superficially resembles a ribosomal protein; i.e., it is small and basic. Although the amino acid sequences of a-sarcin (Sacco et al., 1983), and of the related ribotoxins mitogillin (Fernandez-Luna et al., 1985) and restrictocin (Lopez-Otin et al., 1984), were first determined directly from the proteins, they have been confirmed from the sequences of nucleotides in cDNAs (Henze et al., 1990; Lamy and Davies, 1991; Lacadena et al., 1994; Wnendt et al., 1993). Comparison of amino acid sequences (Sacco et al., 1983) revealed that a-sarcin is related to certain other nucleases; for example, the toxin has 24% amino acid identity with T1 and 34%
4
Structure and Mechanism of a-Sarcin
137
Alpha-sarcin Restrictocin
1 20 40 60 AVTWTCLNDQKNPKTNKYETKRLLYNQNKAE SNSHHAPLS DGKTGS SYPHWFTNGYDGDG A- T W T C I N Q Q L N P K T N K W E D K R L L Y S Q A K A E S N S H H A P L S D G K T G S SY P H W F T N G Y D G N G
Alpha-sarcin Restrictocin
80 KLPKGRTPI KFGKSDCDRPPKHSKDGNGKTDHYLLEF KL I K G R T P I K F G K A D C D R P P K H S Q N G M G K D D H Y L L E F
Alpha-sarcin Restrictocin
140 RVI Y T Y P N K V F C G I I A H T K E N Q G E L K L C S H RVI Y T Y P N K V F C G I V A H Q R G N Q G D L R L C SH
i00 120 PTF P D G H D Y K F D S K K P K E N P G P A PTF P D G H D Y K F D S K K P K E D P G P A
Fig. 2. An alignment of the amino acid sequences of a-sarcin and restrictocin. The proteins share 86% amino acid identity. Restrictocin and mitogillin differ by only one residuemD25 in mitogillin and $25 in restrictocin.
with U2. Moreover, an alignment of the amino acid sequences had suggested that H49, E95, R120, and H136 in restrictocin might be in the catalytic center; these residues correspond to H50, E96, R121, and H137 in a-sarcin. Mutations, E95G and H136L, were constructed in restrictocin; the former had decreased activity whereas the latter was inactive (Yang and Kenealy, 1992), suggesting that H136 in restrictocin (H137 in c~-sarcin) is involved in catalysis. However, more important than the similarities are the differences between a-sarcin and ribonucleases T1 and U2, and more explicitly the differences in structure that account for the differences in properties, aSarcin is a cytotoxin in part because it has acquired a means of entering cells, no small evolutionary accomplishment for a nuclease. It is a property the related ribonucleases do not share. Moreover, a-sarcin selects and cleaves a single phosphodiester bond in ribosomes whereas ribonuclease U2 causes extensive digestion of the R N A in ribosomes. What one wants to know is the portions of the structure of a-sarcin, absent in related ribonucleases, that account for these properties. The Aspergillus toxins, c~-sarcin (Wawrzynczak et al., 1991), restrictocin (Orlandi et al., 1988), and mitogillin (Better et al., 1992) are candidate "magic bullets" for the treatment of cancer and of viral-infected cells, including those infected with human immunodeficiency virus (HIV), and for use in tissue and organ transplantation. In the most common scenario the toxin is coupled to a specific binding protein or to an antibody directed to an antigen on the surface of cancer cells or cells infected with virus. The intention is to deliver the catalytic toxin exclusively to these cells. Immunotoxins have been constructed and they are effective
138
Ira G. Wool
in vitro against cancer cells; moreover, they do not appreciably harm
nonmalignant cells. However, they are less effective when administered to patients and frequently produce intolerable side effects, although some limited success has been achieved recently. The lack of effectiveness has been attributed to the immunotoxins having poor pharmacodynamicsma short half-life, susceptibility to proteolytic degradation, or capture by the immune system.
V. M E C H A N I S M BY W H I C H a S A R C I N E N T E R S CELLS
a-Sarcin is a cytotoxin and is effective in restricting the growth of tumors, which implies that it is able to enter cells, but how it accomplishes this is a mystery. The toxin appears to have little activity on intact mammalian cells (A. Lin and I. G. Wool, 1983, unpublished data). There is no evidence that o~-sarcin has a specific domain that binds to a particular receptor to effect entry; indeed, a search for receptor-mediated translocation was unsuccessful (A. Lin and I. G. Wool, 1983, unpublished data). a-Sarcin is, on the other hand, an effective inhibitor of protein synthesis in intact virus-infected cells (Munoz et al., 1985; Otero and Carrasco, 1985), in cells of some tumor lines (Turnay et al., 1993), and in cells whose membrane permeability has been modified by phospholipase C (Otero and Carrasco, 1988). What these studies imply is that the entry of a-sarcin into cells requires alteration or damage to the cell membrane by viral infection, by malignant transformation, or by some toxic agent. An extension of this reasoning suggests a physiological function of asarcin: that the toxin acts to limit damage to an organism by mediating the death of cells that have, for example, been infected by virus.
VI. a - S A R C I N R E C O G N I T I O N E L E M E N T S IN 28S r R N A
Early on an effort was made to determine whether the specificity of a-sarcin is dependent on the complex ordered structure of 28S rRNA that derives from the presence of the nucleic acid in a particle containing proteins. A contingent aim was to establish the minimal substrate for c~-sarcin. For this purpose, an oligoribonucleotide (a 35-mer) was synthesized using a synthetic D N A template and the phage T7 RNA polymerase
4
Structure and Mechanism of ~-Sarcin
139
(Endo et al., 1988). This oligomer (Fig. 3A) has the nucleotide sequence and the secondary structure of the domain in eukaryotic 28S r R N A that is attacked by c~-sarcin and is referred to as wild type. Treatment of the synthetic oligoribonucleotide with lower concentrations of a-sarcin, i.e., 2.9 x 10 -8 M, led to the formation of two fragments (Fig. 3B, lane 2) (Endo et al., 1990). About 85% of the substrate is cleaved (Fig. 3C). Cleavage by a-sarcin is at the guanosine at position 21 in the synthetic oligoribonucleotide, which corresponds to G4325 in 28S rRNA and is precisely where the toxin hydrolyzes the nucleic acid in intact ribosomes (Endo et al., 1990). In contrast, a higher concentration of a-sarcin (2 x 10 -6 M) led to cleavage of the substrate at all, or nearly all, of the purines (Fig. 3B, lane 5). This conforms with the observation that higher concentrations of a-sarcin, i.e., higher than is necessary to inactivate ribosomes, lead to the hydrolysis on the 3' side of almost every purine in RNA. To define the features of the structure that prescribe binding to the RNA and that are necessary for the catalysis of hydrolysis, the nucleotides in the a-sarcin domain synthetic oligoribonucleotide (Fig. 4, I) were systematically altered (Endo et al., 1990). Cleavage by a-sarcin of a variant of the wild-type RNA with a transition of the guanosine at position 21 to an A (Fig. 4, II) was decreased to 35% of the control. Transversions of the wild-type G to U (Fig. 4, III) or to C (Fig. 4, IV) reduce hydrolysis to approximately 17 and to 1%, respectively. Thus, there is strong, but by no means absolute, dependence on preservation of the G at the site of covalent modification; the preference is G > A > U ~> C. What is notable is that a-sarcin, which had been presumed to be a purine-specific nuclease, has some activity with pyrimidines. The results prejudice one to consider structure rather than sequence as the more important determinant of specificity. The ct-sarcin domain RNA has a canonical protein-binding structure: a stem, a loop, and a bulged nucleotide (Fig. 4, I). The last occurs in a number of ribosomal protein-binding sites. The suspicion was that the bulged U at position 6 in the substrate (position 4310 in 28S rRNA) is not necessary for a-sarcin action because it does not occur at the comparable site in E. coli 23S rRNA, and the bacterial ribosomes are sensitive to the toxin; just as suspected, a variant that lacked the bulged U in the stem (Fig. 4, V) is as sensitive to the toxin as the wild type (Endo et al., 1990). To test the importance of the stem for a-sarcin action, a linear molecule (a 35-mer) was constructed (Fig. 4, XXIII) that retained the 17 nucleo-
140
Ira G. Wool
Fig. 3. The effect of a-sarcin on a synthetic oligoribonucleotide that mimics the toxin domain in 28S rRNA. The synthetic radioactive a-sarcin domain oligoribonucleotide (35-met) (A) was renatured and then (B) incubated at a concentration of 1.6/zM for 20 min at 20~ without ot-sarcin (lane 1) or with increasing concentrations of the toxin: 2.9 x 10 -8 M (lane 2); 5.9 x 10 8 M (lane 3); 2.9 x 10 7 M (lane 4); 2 x 10 -6 M
4
Structure and Mechanism of a-Sarcin
141
tides in the loop sequence but with the 5' 11 nucleotides and the 3' 7 nucleotides altered so that they would not pair (Endo et al., 1990). a-Sarcin did not cleave the linear substrate specifically. However, there was nonspecific digestion; indeed, the linear RNA appears to be more sensitive to this nonspecific effect of the toxin than the structured RNA. Having established that the stem is required, the number of base pairs in the helix that are needed was determined (Endo et al., 1990). Successive additional base pairs were deleted from the wild-type oligoribonucleotide (Fig. 4, V I - I X ) . A variant having only three pairs in the stem (Fig. 4, IX) was recognized by a-sarcin. It seems likely that the helix is necessary only to tether the ends of the loop so as to allow it to form a specific conformation. In nature the helix may be longer and hence more stable than is required for recognition by a-sarcin; we presume it is neither longer nor more stable than is required for its contribution to the function of this ribosomal domain in protein synthesis. Synthetic oligodeoxynucleotide analogs of E. coli tRNA Ph~ and of tRNA Ly~ are recognized by their cognate aminoacyl-tRNA synthetases. A deoxy analog of the wild-type a-sarcin RNA (Fig. 4, XIV), on the other hand, was not cleaved specifically by a-sarcin (Endo et al., 1990). However, there was nonspecific digestion and, as we had observed before, the hydrolysis of the DNA required magnesium; nonspecific cleavage of RNA by a-sarcin is moderately inhibited by magnesium. The effect of the context of the a-sarcin site guanosine on recognition was evaluated (Endo et al., 1990). It was anticipated that alterations in
(lane 5). A radioautograph of the polyacrylamide-urea gel was used for the analysis. The radioactivity in each of the bands (of 35, 21, and 14 nucleotides) was determined for lanes 2-5, and (C) the percentage of the original substrate that was specifically cleaved was plotted as a function of the concentration of a-sarcin. ( 9 1 6 9 Wild-type 35-mer; cf. A. ( O - - O ) The 34-mer lacking the wild-type bulged nucleotide at position 6. In A, the S and the arrow designate the site of cleavage by a-sarcin. In B, the numbers on the left indicate the number of nucleotides in the RNA. The shadow bands that form a ladder are oligoribonucleotides that most probably were formed by degradation, perhaps due to radiation damage.
S S G
18
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xxm Fig. 4. The structures (I-XXII ) of variants of the a-sarcin domain oligoribonucleotide.
4
Structure and Mechanism of a-Sarcin
143
the 5' adjacent adenosine would not have an appreciable effect because depurination of A4324 by pretreatment with ricin did not affect subsequent cleavage by a-sarcin at G4325 in the same ribosomes. Nonetheless, a series of variants were constructed with alterations of the ricin site A to G, U, or C (Fig. 4, X V - X V I I ) . As expected, all were recognized by a-sarcin with specificity and with normal efficiency. The context was changed in another way: the tetranucleotide GAG(sarcin)A was left intact but the remainder of the universal portion of the loop sequence was engineered so that it was entirely uridines (Fig. 4, XVIII) or, in a second variant, uridines and guanosines (Fig. 4, XIX) (Endo et al., 1990). Neither oligonucleotide is a competent substrate for a-sarcin. Thus context either directly or indirectly influences the recognition of the R N A by c~-sarcin. The specific response to a-sarcin is also lost when the position of the tetranucleotide G A G A in the loop is changed (Fig. 4, X X - X X I I ) (Endo et al., 1990). The results with one of the variants is particularly instructive (Fig. 4, XXI). In this mutant there is still a guanosine at position 21, as in the wild-type substrate; nonetheless, there was no cleavage there. Thus, recognition is not merely of a guanosine at the correct position in the sequence, a-Sarcin appears to appreciate the structure of the loop and this structure is no doubt affected by the nucleotide sequence. The conclusion to this point was that specific recognition of rRNA by a-sarcin requires, in the first instance, a stem and a loop, but that a bulged nucleotide in the stem is not necessary. There is a strong preference for a guanosine at the site of covalent modification and the surounding context, i.e., the 12-base universal nucleotide sequence affects, either directly or indirectly, the binding of a-sarcin and catalysis. The exception is the immediate 5' adjacent adenosine, which has no influence on identification of the RNA. The stem is essential; however, only three of the seven base pairs found in 28S rRNA are needed. The helical stem appears to contribute to recognition only indirectly by tethering the ends of the loop and conferring on the latter a specific conformation. Finally, the position of the tetranucleotide GAG(sarcin)A in the loop conditions recognition, once again, either directly or indirectly by altering the context and hence its conformation. Concurrent experiments with ricin A chain (RA), a cytotoxin that inactivates ribosomes by depurinating the adenosine at position 4324 in 28S rRNA (Fig. 1), the nucleotide 5' adjacent to the a-sarcin cleavage site (Endo and Tsurugi, 1987, 1988), had indicated that RA has an absolute requirement for a G A G A tetraloop; moreover, there was strong
144
Ira G. Wool
evidence that the tetraloop exists, at least transiently, in intact ribosomes (Endo et al., 1991; G10ck et al., 1992). a-Sarcin, on the other hand, does not recognize oligoribonucleotide substrates that have only a G A G A tetraloop. If there is a tetraloop in the sarcin domain in 28S r R N A , it would have to be closed off by a base pair between the cytosine on the 5' side and the guanosine on the 3' side of the G A G A tetranucleotide (Fig. 5 I). A series of mutants was constructed to test this prediction (Gltick et al., 1994). a-Sarcin catalyzes covalent modification of a variant having a reversal of the 5' C and the 3' G (Fig. 5, If). This is a rare change in the G A G A context that is tolerated. In addition, a-sarcin modifies a mutant that has a transversion of the 3' G to a C (Fig. 5, I l l ) , albeit the
R
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A
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s
A\
Ac G A GA6~,A
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A
A
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G A
A
A
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s
Fig. 5. The structures (1-VII) of a set of oligoribonucleotide mutants of the a-sarcin domain RNA in which the 5'-guanosine and the 3'-cytosine that surround the GAGA tetranucleotide have been varied.
4
Structure and Mechanism of a-Sarcin
145
effect is moderately diminished. RA does not recognize this mutant. This is a discontinuity in the identity elements for RA and for a-sarcin. The decrease in cleavage by a-sarcin occasioned by the G-to-C mutation is almost entirely the result of an effect on catalysis. The kcat is decreased almost by an order of magnitude and the kcat/gm by 0.15, but the affinity of the toxin for the substrate is hardly changed. A mutant with simultaneous transversions of the 5' C to A and of the 3' G to U creating a potential A U (Fig. 5, V) pair is affected by asarcin and by RA (Gltick et al., 1994). However, simultaneous changes of the 5' C and the 3' G to A (Fig. 5, VI) or, in a separate mutant, to U (Fig. 5, VII) lead to loss of recognition by RA but not by a-sarcin, although, once again the response to a-sarcin is reduced. Obviously, RA cannot depurinate oligoribonucleotides that lack the capacity to shut off a GAGA tetraloop by forming a closing Watson-Crick pair; it is also clear that the inability to do so modestly impairs, but certainly does not abolish, recognition by a-sarcin.
VII. C O N F O R M A T I O N OF THE SARCIN D O M A I N IN 28S r R N A
From the beginning, experiments have addressed two separate structure-function problems (Wool et al., 1992). The first is the structure of the sarcin domain RNA when resident in the ribosome and the contribution of this structure to the biochemistry of protein synthesis. This is job one. The second problem is the identity elements for the recognition by a-sarcin of a rRNA domain and hence the chemistry of a particular RNA-protein association. The structural requirements for protein synthesis are far more stringent than for toxin recognition (Wool et al., 1992). An observation that may be important for understanding the contribution of the sarcin district of 28S rRNA to the biochemistry of protein synthesis is that the identity elements for RA and for a-sarcin are different (Wool et al., 1992). RA requires a G A G A tetraloop whereas asarcin does not. Moreover, if the capacity to form a G A G A tetraloop is abolished, modification of the R N A by RA is lost, but that by a-sarcin is not. Thus, the identity elements required by RA and by a-sarcin are different, yet the two ribotoxins catalyze covalent modifications of adjacent nucleotides in rRNA. The simplest rationalization of these observations is that the identity elements for a-sarcin lie outside of the G A G A tetraloop.
146
Ira G. Wool
The universal sequence of nucleotides in the sarcin domain did not evolve to maximize the efficiency of recognition by the toxin; the pressure presumably was to facilitate the binding of two related but dissimilar proteins, the elongation factors 1 and 2 (or Tu and G). The elongation factors bind to essentially the same site on the ribosome (Moazed et al., 1988) and a single round of peptide bond formation requires the displacement of the first (EF-1 or EF-Tu) to allow binding of the second (EF-2 or EF-G) and vice versa. Thus, it may be the necessity to bind two different proteins, in an ordered fashion, that explains the extraordinary conservation of the sequence of nucleotides in the loop. These nucleotides may be necessary also for reversible transitions in conformation, which may in turn be needed for protein synthesis. Obviously, what is essential to the resolution of these problems are the details of the structure of the sarcin domain. There is a model of the conformation of the domain derived from nuclear magnetic resonance (NMR) spectroscopy (Scewczak et al., 1993; Szewczak and Moore, 1995). The proposed three-dimensional conformation of the sarcin RNA (Fig. 6) satisfies all of the NMR spectroscopic data, is entirely consistent with the RNA mutants, and perhaps most importantly the hydrogen bonding pattern agrees precisely with chemical modification data obtained independently (H. F. Noller and D. Moazed, 1987, unpublished data; quoted in Szewczak et al., 1993 and Szewczak and Moore, 1995). The structure has, just as was predicted from analysis of mutants, a G A G A tetraloop shut off by a C.G pair (Fig. 6). However, there is a noncanonical, heteropurinic intraloop G - A pair; thus there are only two unpaired nucleotides in the loop. Below the closing C.G pair there is a second heteropurinic A - G pair just as in the tetraloop. This is followed by a reversed Hoogsteen U - A pair, just above which the stacking of the bases in the strands cross over. G10 is far and away the most interesting residue in this RNA. The sequential pattern of nuclear Overhauser enhancement (NOE) connectivities breaks at G10, giving every indication it is a bulged base. Moreover, there is a NOE between A9 and U l l , the only nonsequential aromatic-aromatic N O E in the molecule. The key to the structure, however, is a NOE between the imino proton of G10 and the sugar proton of G19, which implies they are close, and a model has been built in which G10 reaches across the U 1 l - A 2 0 pair toward the ribose of G19 and places its imino proton within hydrogen bonding distance of the phosphate oxygen between G19 and A20. There is next a symmetric homopurinic A - A pair; below this C8 and U7 are juxtaposed to C22 and C23 with no evidence of pairing. Finally, there is a six-base pair
4
Structure and Mechanism of a-Sarcin
147
Fig. 6. The structure of the a-sarcin domain RNA. (Left) Schematic of the secondary structure of the ot-sarcin domain. (Right) Diagram of the hydrogen bonding and base stacking interactions derived from NMR spectroscopy of the same oligoribonucleotide (derived from Szewczak et al., 1993). with permission). The open arrow designates the Nglycosidic bond cleaved by ricin; the filled arrow designates the phosphodiester bond cleaved by ot-sarcin.
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Ira G. Wool
canonical A-form R N A helix. Thus the R N A , which in secondary structure diagrams is depicted as having a 17-member loop (Fig. 6, on the left), in actuality has a compact conformation that approximates an extended helix (Fig. 7) with few unpaired nucleotides and most strikingly many noncanonical pairs (Fig. 6, on the right). The pattern of chemical modification of nucleotides in the sarcin loop in the free R N A differs from that in the ribosome (H. F. Noller and D. Moazed, 1987, unpublished data). Because the differences include reactivity increases, as well as the expected decreases due to interactions with proteins and neighboring R N A segments, the conformation of the sacrin loop in the ribosome cannot be exactly the same as the one just described. It has been suggested that conformational changes in the sarcin loop trigger translocation (Wool et al., 1992). Because both a-sarcin and R A are able to attack the sarcin loop in the ribosomes, and because a good case can be made that a conformation like the one discussed here is necessary for sensitivity to RA, it seems reasonable to suggest, as Szewczak and Moore (1995) have, that this structure is a conformation that is present in the ribosome at some stage during elongation.
VIII. E F F E C T OF M U T A T I O N S OF T H E A N A L O G OF G4319 IN 28S r R N A O N R E C O G N I T I O N OF O L I G O R I B O N U C L E O T I D E S BY a - S A R C I N
In the three-dimensional conformation of the sarcin domain R N A the guanosine at position 15 (G10 in the oligoribonucleotide used to derive the N M R conformation, G15 in the wild-type oligoribonucleotide described in this section, and G4319 in 28S r R N A ) is bulged (Fig. 6) and produces a prominent kink in the helical structure (Fig. 7) (Szewczak et al., 1993). Although, the R A recognition element can be described with some precision, it is a G A G A tetraloop, that for
Fig. 7. A three-dimensional conformation of the a-sarcin domain RNA. The conformation was obtained from the coordinates deposited in the Brookhaven data base with the program GRASP at a Silicon Graphics workstation. The bulged guanosine (gold) and the sites of action of a-sarcin (blue) and of ricin (purple) are designated. The phosphodiester backbone is represented by a red ribbon. Derived from Szewczak et al. (1993).
4
Structure and Mechanism of a-Sarcin
149
a-sarcin at first could only be d e f i n e d n e g a t i v e l y u m o s t definitively a G A G A t e t r a l o o p is not a c o m p e t e n t substrate. B e c a u s e the c o n f o r m a tion of G15 is such a p r o m i n e n t f e a t u r e of the N M R structure, a t t e n t i o n was d i r e c t e d to it (Gltick a n d W o o l , 1996). F o u r m u t a t i o n s of G15 w e r e constructed: the d e l e t i o n of the n u c l e o t i d e , a t r a n s i t i o n to a d e n o s i n e , a n d t r a n s v e r s i o n s to cytidine a n d to uridine (Fig. 8). T h r e e of the four variants are n o t r e c o g n i z e d by a-sarcin at all; cleavage of the fourth, a G 1 5 A m u t a n t , is b a r e l y d e t e c t a b l e at the highest c o n c e n t r a t i o n of c~-sarcin a n d e v e n this m i n i m a l hydrolysis is n o t consistent. Thus, G4319 is the critical n u c l e o t i d e for c~-sarcin recognition.
IX. P H E N O T Y P E 23S r R N A
O F M U T A T I O N S IN G2655 IN Escherichia coli
O n e way, p e r h a p s the way, to define the function of the sarcin d o m a i n r R N A in p r o t e i n synthesis is by the analysis of the p h e n o t y p e of mutants. M u t a t i o n s have b e e n c o n s t r u c t e d in G2655 in E. coli 23S r R N A (Fig. 9) (M. M a c b e t h and I. G. W o o l , 1995, u n p u b l i s h e d data): the deletion of the n u c l e o t i d e (G2655A), a transition to a d e n o s i n e ( G 2 6 5 5 A ) , a n d t r a n s v e r s i o n s to cytosine ( G 2 6 5 5 C ) and to uridine ( G 2 6 5 5 U ) . G2655 is the b u l g e d g u a n o s i n e that is critical for c~-sarcin r e c o g n i t i o n a n d it is a n u c l e o t i d e that is p r o t e c t e d against chemical
Fig. 12. The structure of the active site in restrictocin. The five-stranded/3 sheet is twisted and forms a cleft; the catalytic residues (in red) are at the open end of the cleft. An inorganic phosphate (in yellow and red) is also at the active site. The two disulfide bonds are in orange. Fig. 13. A model of the complex of restrictocin and a-sarcin domain RNA. The model is from a docking experiment using the coordinates of the NMR conformation of the c~sarcin domain RNA (Szewczak et al., 1993) and those for restrictocin (Yang and Moffat, 1996). On the left, the positions of the bulged guanosine (G10 in Fig. 6) and the guanosine at the cleavage site (G16 in Fig. 6) in the RNA are in orange; the charge distribution on the surface of restrictocin is indicated by red (negative) and blue (positive). On the right, the complex has been rotated 90~to show the complementarity of the surfaces of restrictocin (blue, on top) and of the RNA (red, below).
150
Ira G. Wool R
R
s G
u cc"
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R
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.CG;o %
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R
~ /s
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C
~ /s
.CG;oG%
u
G
Co cc" C-G
to- G-C-3o
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U\ U
G
cc"
C-G
Jo- G-C-3o
U-A C-G C-G UA- U A'UoH 3' G G
5' pppG
Fig. 8. The structures of a set of oligoribonucleotide mutants of the a-sarcin domain RNA in which the guanosine at position 15, the analog of G4329 in 28S rRNA, was varied.
modification by EF-Tu and EF-G. The mutations were constructed in an rrnB gene in a high-copy-number plasmid that has an erythromycinresistance marker. In bacteria in which the plasmid 23S rRNA gene has a chromosometype PiP2 promoter, 70% of the cellular rRNA is derived from the plasmid. The G2655A, G2655C, and G2655U mutants do not grow; they have a dominant lethal phenotype (M. Macbeth and I. G. Wool, 1995, unpublished data). To our surprise and to our mystification, the G2655A mutant grows normally on agar plates and in liquid media. This was a surprise because, to our knowledge, no organism in the biosphere has an adenosine at this position and an adenosine was thought unlikely to reproduce the unusual bulged conformation found in the N M R structure. However, in a competitive growth experiment in which equal numbers of wild-type and G2655A mutant cells were mixed and grown together for approximately 150 generations, there was a definite phenotype: no G2655A mutant cells survived, i.e., they could not compete with the wild type. The plasmid 23S rDNA has silent mutations near the sarcin region (Fig. 9, inset) that allow the source of the R N A in ribosomes (plasmid or chromosomal) to be established; the determination is with specific oligodeoxynucleotide primers and reverse transcriptase. This has made it possible to test the sensitivity of the mutant ribosomes
4 Structure and Mechanism of a-Sarcin
151 6 5 5 - > A. U. C. A
2 U U
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Fig. 9. The a-sarcin domain of E. coli 23S rRNA. The mutants of G2655 that were constructed are designated. Inset: Region and identity of silent nearby mutations used to prime specifically for the mutant plasmid-encoded 23S rRNA.
to a-sarcin (M. M a c b e t h and I. G. W o o l , 1995, unpublished data). R i b o s o m e s with plasmid-derived 23S r R N A having a m u t a t i o n at G 2 6 5 5 are not affected by a-sarcin, w h e t h e r the m u t a t i o n is a deletion, a transition to a d e n o s i n e , or a transversion to cytosine or u r i d i n e - - a l l are resistant to the toxin. Thus, the results with r i b o s o m e s c o n f o r m precisely with the results obtained in vitro with o l i g o r i b o n u c l e o t i d e s (see Section VIII).
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Ira G. Wool
X. R E L A T I O N S H I P OF THE a - S A R C I N R N A I D E N T I T Y E L E M E N T TO SELECTION BY THE T O X I N OF THE U N I Q U E SITE OF PHOSPHODIESTER BOND HYDROLYSIS
How can one explain the relationship between the identity element, G4319, and the cleavage by this small enzyme (recall that c~-sarcin has 150 amino acids) of the phosphodiester bond on the 3' side of the relatively distant G4325? One possibility is that c~-sarcin recognizes or induces a conformation of the sarcin domain that is different than that derived from NMR spectroscopy; the assumption is that in this putative alternate conformation the recognition nucleotide and the hydrolytic site are closer. This should not be dismissed out of hand because, as has been pointed out, the sarcin loop R N A is not very stable thermodynamically; moreover, the interpretation would accord with the speculation that conformational changes in the domain drive translocation. Unfortunately the possibility is not easily tested. The bulged G10 (the G4319 analog) is about 15 A away but on the same side of the sarcin loop as the phosphate group of the diester that is cleaved. This suggests a second possibility: that c~-sarcin recognizes G4319 and then cuts at a fixed distance. The virtue of this proposal is that it can be tested. The test requires the construction of a suitable substrate: the basic operation is to insert two pairs of nucleotides between G14.A17 and C13.G18 so as to change the distance between G15 (G4319) and the wild-type hydrolytic site (Gltick and Wool, 1996). In preliminary experiments, promising results were obtained with this construct (Fig. 10A): c~-sarcin-catalyzed cleavage occurred at a new, equidistant site just as predicted. This was confirmed in separate experiments by identification of the source of the cleavage fragments with 5' and 3' end-labeled oligoribonucleotides. Nonetheless, the results were not decisive; it was clear that distance was not the only determinant. The nature of the inserted nucleotides was important, at least in a negative sense; they could not form Watson-Crick pairs. In the initial experiment, the inserted 3' side nucleotides were guanosine (17a) and adenosine (17b) to preserve the sequence of the wild-type cut site; the 5' side nucleotides, 13a and 13b, were randomized (Fig. 10A). When a construct was synthesized with uridine (13a) and cytosine (13b) inserted on the 5' side to complement the guanosine (17a) and adenosine (17b) inserted on the 3' side, thereby permitting the formation of two presumably stable pairs in the loop (i.e., C13b 9G17a and U13a 9A17b), no hydrolysis by
4
Structure and Mechanism of ~-Sarcin
153
c~-sarcin occurred at either the new or old sites. However, if the 5' insertion was either adenosine (13a) and guanosine (13b) or cytosine (13a) and adenosine (13b) (Fig. 10B), which are unlikely to form stable pairs with the nucleotides inserted on the 3' side, i.e., guanosine (17a) and adenosine (17b), there was cleavage by the toxin at the new site. This is a strong indication that c~-sarcin requires an open, or at least an unstable, structure in the sarcin loop; a requirement that was implied by earlier experiments. But even with these constructs there was a small amount of residual cleavage at the old, or wild-type, site. For this reason the wild-type G - A cleavage sequence was changed to C - C (Fig. 10C) (Gltick and Wool, 1996). Hydrolysis with this substrate was exclusively at the new site. It needs to be noted that hydrolysis by c~-sarcin at the new site in the variant oligoribonucleotide is not as efficient as at the old site in the wild-type RNA. An assessment of the extent of the decrease in efficiency as a function of c~-sarcin concentration indicates that cleavage at the new site is only about 25% of that at the old site. What is particularly important, however, is that cleavage at the new site retains its dependence on the G4319 analog; just as with the wild-type substrate, deletion of that guanosine or its transversion to a cytidine abolishes hydrolysis. The results suggest that G4319 is the identity element for c~-sarcin recognition, that the toxin binds to G4319 or at least in close proximity to the nucleotide, and that binding allows c~-sarcin to orient itself and to catalyze hydrolysis of a phosphodiester bond at a fixed distance and at a fixed position in space relative to G4319. Manipulation of the structure of the R N A substrate has uncovered this aspect of how recognition is related to catalysis. That cleavage at the new site in the variant R N A is never as efficient as at the old site in the wild-type R N A indicates that not all of the structural features of the relationship have as yet been defined.
XI. T H R E E - D I M E N S I O N A L S T R U C T U R E OF R E S T R I C T O C I N , A H O M O L O G OF a~-SARCIN. B I N D I N G TO a - S A R C I N D O M A I N RNA AND THE CATALYTIC MECHANISM
The three-dimensional structure of restrictocin, refined to a resolution of 1.7 .A,, has been determined by X-ray diffraction of single crystals
154
Ira G. Wool
Fig. 10. Analysis of the relationship of the distance between the a-sarcin identity guanosine, the analog of G4319 in 28S rRNA, and the site of cleavage. The structure of the wild-type oligoribonucleotide is in Fig. 6. (A) A mixture of 16 mutant oligomers in which the nucleotides at positions 13a and 13b were randomized (they are designated N).
( Y a n g and Moffat, 1996). Recall that restrictocin shares 86% a m i n o acid identity with a - s a r c i n and has the s a m e m e c h a n i s m of action. Restrictocin has two a helices and two a n t i p a r a l l e l / ~ sheets (Fig. 11). T h e structural core is c o m p o s e d of a p e r p e n d i c u l a r ct helix of t h r e e turns p a c k e d against a five-stranded antiparallel fl s h e e t that is stabilized by
4
Structure and Mechanism of a-Sarcin
155
Fig. 10. (Continued). (B) The residues at 13a and 13b are cytosine and adenosine.
hydrophobic residues. The NH2 terminus has a long two-stranded antiparallel/3 sheet. A particularly distinctive feature of restrictocin is the large connecting loops between/3 strands. The structural core forms a cleft that has the putative catalytic residues and an inorganic phosphate believed to be derived from the crystallization buffer (Fig. 12). These five residues in restrictocin (Y47, H49, E95, R120, and H136) exactly reproduce five in the catalytic center of RNase T1. Although restrictocin and T1 share 24% amino acid identity, the latter has only 104 residues whereas restrictocin has 149. That the structure
156
Ira G. Wool
Fig. 10. (Continued). (C) the residues at positions 16 and 17 are changed to cytosine. The open arrow designates the site of cleavage in the wild-type substrate; the filled arrow designates the new site.
of the core in restrictocin closely resembles that in T1 strongly suggests that the mechanism of phosphodiester bond cleavage is the same. Studies of the T1 catalytic mechanism had suggested a two-step reaction (cf. Yang and Moffat, 1996): the first is a phosphoryl transfer in which the base-acid couple H49 and E95 (the residues and positions are in restrictocin but identified by analogy to T1) act as the general base to abstract, a proton from the 2'-hydroxyl of a ribose and H136 serves as
4
Structure and Mechanism of ~-Sarcin
157
Fig. 11. The secondary structure elements in restrictocin. A ribbon diagram of the secondary structure of restrictocin in which the elements are designated. The core of the enzyme consists of a five-stranded antiparallel/3 sheet (strands B3, B4, B5, B6, and B7) stabilized by an c~ helix (HI) having three turns that is perpendicular to the/3 sheet. The NH2 terminus has a second antiparallel/3 sheet with two strands (BI and B2) linked by loop L1, which has six residues that are not resolved. There is a helix (H2) with a single turn in the loop (L3) that links strands B3 and B4.
a general acid to protonate the 5'-oxygen of the leaving nucleotide of the RNA (which now has a 5'-hydroxyl) yielding a 2',3'-cyclic phosphate intermediate. The inorganic phosphate in the restrictocin structure may occupy the site used by the phosphorus of the 2',3'-cyclic intermediate. In the hydrolysis reaction, the roles of the catalytic residues are reversed; H136 serves as the general base and activates a water molecule; H49 and E95 donate a proton to the 2'-oxygen. The activated water attacks the phosphorus in the cyclic intermediate and forms the second product, an RNA with a 3'-phosphate. The role of R120 is not clear; it may electrostatically stabilize the phosphate group or it may support the
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Ira G. Wool
geometry of the active site through a network of hydrogen bonds. It needs to be noted that there is some disagreement about the T~ catalytic mechanism, particularly about the assignment of the residues that serve as general base and general acid. Thus, the analysis of site-directed mutants of restrictocin designed to probe the catalytic mechanism will be helpful. Restrictocin has secondary structural elements very much like those in T~ and some other ribonucleases; what sets restrictocin apart from T~ and the other ribonucleases is the extent and the complexity of its loops (Figs. 11 and 12). Presumably it is amino acid sequences and tertiary structures in one or more of the loops that endow restrictocin with its specificity for a single nucleotide in 28S rRNA and that, perhaps, mediate entry of the toxin into the cell. In this interpretation it is the loops that give restrictocin its toxic phenotype. Loop 4, which spans residues 99 to 117, is exposed to solvent, is mobile and rich in lysine residues, and is critically positioned to play a role in the specificity of substrate binding. Restrictocin has a dome and a planar side with the catalytic residues at one end of a shallow cleft that is extended by a platform formed by loops L2 and L4; the concave surface of the platform is a possible domain for the binding of RNA (Fig. 13). Using the coordinates for the NMR conformation of the sarcin domain RNA, a docking experiment was done with restrictocin (Yang and Moffat, 1996). The program GRASP was used and advantage was taken of the fit of the concave surface of restrictocin and the crown (the G A G A tetraloop) of the RNA (Fig. 13). The lysine-rich loop L4 was found to be positioned in the major groove of the helical stem. This fit has the phosphorus of the phosjghodiester bond between G16 and A17 at the cleavage site only 0.93 A from the phosphate in the restrictocin crystal structure. In the model derived from the docking experiment, a platform formed by loops L2 and L4 provides most of the interface with the RNA: K42 in loop L2 and K106, Kll0, K l l l , and Kl13 in loop L4 could form salt bridges with phosphates in the RNA. The side chain of D108 or of S109 might form hydrogen bonds with bases in the RNA and facilitate discrimination. The positively charged ridge formed by K60, K63, K80, K88, and R65 in loop L3 is close to the sugar-phosphate backbone of G13-G16 in the R N A to further facilitate the interaction. Perhaps the most important feature of the model is that the bulged G10, the critical c~-sarcin recognition element in the RNA, is close to the lysine-rich loop L4. Thus, the structural elements in restrictocin that account for the specificity of RNA recognition are the platform formed by loops L2 and L4, and the positively charged ridge in loop L3.
4
Structure and Mechanism of a-Sarcin
159
XII. C O D A
Willie Sutton, when asked why he robbed banks, replied in what has become a cliche, "because that is where the money is." We have studied the mechanism of action of c~-sarcin because we want information on the molecular basis of the function of ribosomes in protein synthesis, on the chemistry of a very specific p r o t e i n - R N A interaction, and on the mechanism of catalysis by a special ribonucleasemand the c~-sarcin domain r R N A is where that information is. The studies of the mechanism of action of c~-sarcin have been fruitful and there is every indication they will continue to be.
REFERENCES
Better, M., Bernhard, S., Lei, S. P., Fishwild, D., and Carroll, S. (1992). Activity of recombinant mitogillin and mitogillin immunoconjugates. J. Biol. Chem. 267, 1671216718. Chan, Y. L., Endo, Y., and Wool, I. G. (1983). The sequence of the nucleotides at the c~sarcin cleavage site in rat 28S ribosomal ribonucleic acid. J. Biol. Chem. 258, 1276812770. Conde, F. P., Fernandez-Puentes, C., Montero, M. T. V., and Vazquez, D. (1978). Protein toxins that catalytically inactivate ribosomes from eukaryotic microorganisms. Studies of the mode of action of alpha sarcin, mitogillen and restrictocin: Response to alpha sarcin antibodies. F E M S Microbiol. Lett. 4, 349-355. Endo, Y., and Tsurugi, K. (1987). The mechanism of action of ricin and related toxic lectins on eukaryotic ribosomes. The site and characteristics of the modification in 28S ribosomal RNA caused by the toxins. J. Biol. Chem. 262, 5908-5912. Endo, Y., and Tsurugi, K. (1988). The RNA N-glycosidase activity of ricin A-chain. The characteristics of the enzymatic activity of ricin A-chain with ribosomes and with rRNA. J. Biol. Chem. 263, 8735-8739. Endo, Y., and Wool, I. G. (1982). The site of action of c~-sarcin on eukaryotic ribosomes. The sequence at the c~-sarcin cleavage site in 28S ribosomal ribonucleic acid. J. Biol. Chem. 257, 9054-9060. Endo, Y., Huber, P. W., and Wool, I. G. (1983). The ribonuclease activity of the cytotoxin c~-sarcin. The characteristics of the enzymatic activity of c~-sarcin with ribosomes and ribonucleic acids as substrates. J. Biol. Chem. 258, 2662-2667. Endo, Y., Chan, Y. L., Lin, A., Tsurugi, K., and Wool, I. G. (1988). The cytotoxins c~sarcin and ricin retain their specificity when tested on a synthetic oligoribonucleotide (35-mer) that mimics a region of 28S ribosomal ribonucleic acid. J. Biol. Chem. 263, 7917-7920.
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Endo, Y., Gltick, A., Chan, Y. L., Tsurugi, K., and Wool, I. G. (1990). RNA-protein interaction. An analysis with RNA oligonucleotides of the recognition by c~-sarcin of a ribosomal domain critical for function. J. Biol. Chem. 265, 2216-2222. Endo, Y., Gl0ck, A., and Wool, I. G. (1991). Ribosomal RNA identity elements for ricin A-chain recognition and catalysis. J. Mol. Biol. 221, 193-207. Endo, Y., Oka, T., Tsurugi, K., and Natori, Y. (1993a). The biosynthesis of a cytotoxic protein, c~-sarcin, in a mold of Aspergillus giganteus I. Synthesis of prepro- and proot-sarin in vitro. Tokushima J. Exp. Med. 40, 1-6. Endo, Y., Oka, T., Tsurugi, K., and Natori, Y. (1993b). The biosynthesis of a cytotoxic protein, ot-sarcin, in a mold of Aspergillus giganteus. II. Maturation of precursor form of ot-sarcin in vitro. Tokushima J. Exp. Med. 40, 7-12. Fando, J. L., Alaba, I., Escarmis, C., Fernandez-Luna, J. L., Mendez, E., and Salinas, M. (1985). The mode of action of restrictocin and mitogillin on eukaryotic ribosomes. Eur. J. Biochem. 149, 29-34. Fernandez-Luna, J. L., Lopez-Otin, C., Soriano, F., and Mendez, E. (1985). Complete amino acid sequence of the AspergiHus cytotoxin mitogillin. Biochemistry 24, 861-867. Fernandez-Puentes, C., and Vazquez, D. (1977). Effects of some proteins that inactivate the eukaryotic ribosome. FEBS Lett. 78, 143-146. Gltick, A., and Wool, I. G. (1996). Determination of the 28S ribosomal RNA identity element (G4319) for alpha-sarcin and the relationship of recognition to the selection of the catalytic site. J. Mol. Biol. 256, 838-848. Gl0ck, A., Endo, Y., and Wool, I. G. (1992). Ribosomal RNA identity elements for ricin A-chain recognition and catalysis. Analysis with tetraloop mutants. J. Mol. Biol. 226, 411-424. Grtick, A., Endo, Y., and Wool, I. G. (1994). The ribosomal RNA identity elements for ricin and for c~-sarcin, mutations in the putative CG pair that closes a G A G A tctraloop. Nucleic Acids Res. 22, 321-324. Gutcll, R. R., and Fox, G. E. (1988). A compilation of large subunit RNA sequences presented in a structural format. Nucleic Acids Res. 16, r175-r269. Hausncr, T. P., Atmadja, J., and Nicrhaus, K. H. (1987). Evidence that the G2661 region of 23S rRNA is located at the ribosomal binding sites of both elongation factors. Biochimie 69, 911-923. Hcnzc, P. P., Hahn, U., Erdmann, V. A., and Ulbrich, N. (1990). Expression of the chemically synthesized coding region for the cytotoxin ot-sarcin in Escherichia coli using a secretion cloning vector. Eur. J. Biochem. 192, 127-131. Hobden, A. N., and Cundliffe, E. (1978). The mode of action of alpha sarcin and a novel assay of the puromycin reaction. Biochem. J. 170, 57-61. Lacadena, J., Pozo, A. M., Barbero, J., Mancheno, J. M., Gasset, M., Ofiaderra, M., LopezOtin, C., Ortega, S., Garcia, J., and Gavilanes, J. G. (1994). Overproduction and purification of biological active native fungal c~-sarcin in Escherichia coli. Gene 142, 147-151. Lamy, B., and Davies, J. (1991). Isolation and nucleotide sequence of the Aspergillus restrictus gene coding for the ribonucleolytic toxin restrictocin and its expression in Aspergillus nidulans: The leader sequence protects producing strains from suicide. Nucleic Acids Res. 19, 1001-1006. Lopez-Otin, C., Barber, D., Fernandez-Luna, J. L., Soriano, F., and Mendez, E. (1984). The primary structurer of the cytotoxin restrictocin. Eur. J. Biochem. 143, 621-634.
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Miller, S. P., and Bodley, J. W. (1988). ot-Sarcin cleaves ribosomal RNA at the ot-sarcin site in the absence of ribosomal proteins. Biochem. Biophys. Res. Commun. 154, 404-410. Moazed, D., Robertson, J. M., and Noller, H. F. (1988). Interaction of elongation factors EF-G and EF-Tu with a conserved loop in 23S RNA. Nature (London) 334, 362-364. Munoz, A., Castrillo, J. L., and Carrasco, L. (1985). Modification of membrane permeability during Semliki Forest Virus infection. Virology 146, 203-212. Olson, B. H., and Goerner, G. L. (1965). Alpha sarcin, a new antitumor agent. I. Isolation, purification, chemical composition, and the identity of a new amino acid. Appl. Microbiol. 13, 314-321. Olson, B. H., Jennings, J. C., Roga, V., Junek, A. J., and Schuurmans, D. M. (1965). Alpha sarcin, a new antitumor agent. II. Fermentation and antitumor spectrum. Appl. Microbiol. 13, 332-326. Orlandi, R., Canevari, S., Conde, F. P., Leoni, F., Mezzanzaniza, D., Ripamonti, M., and Colnaghi, M. I. (1988). Immunconjugate generation between the ribosome inactiving protein restrictocin and anti-human breast carcinoma MAB. Cancer Immunol. Immunother. 26, 114-120. Otero, M. J., and Carrasco, L. (1985). Proteins are cointernalized with virion particles during early infection. Virology 160, 75-80. Otero, M. J., and Carrasco, L. (1988). Exogenous phospholipase C permeabilizes mammalian cells to proteins. Exp. Cell Res. 177, 154-161. Sacco, G., Drickamer, K., and Wool, I. G. (1983). The primary structure of the cytotoxin ot-sarcin. J. Biol. Chem. 258, 5811-5818. Schindler, D. G., and Davies, J. E. (1977). Specific cleavage of ribosomal RNA caused by alpha sarcin. Nucleic Acids Res. 4, 1097-1110. Stern, S., Powers, T., Changchien, L. M., and Noller, H. F. (1989). RNA-protein interactions in 30S ribosomal subunits: Folding and function of 16S rRNA. Science 244, 783-790. Szewczak, A. A., and Moore, P. B. (1995). The sarcin/ricin loop, a modular RNA. J. Mol. Biol. 247, 81-98. Szewczak, A. A., Moore, P. B., Chan, Y. L., and Wool, I. G. (1993). The conformation of the sarcin/ricin loop from 28S ribosomal RNA. Proc. Natl. Acad. Sci. U.S.A. 90, 9581-9585. Turnay, J., Olmo, N., Jiminez, A., Lizarbe, M., and Gavilanes, J. (1993). Kinetic study of the cytotoxic effect of cz-sarcin, a ribosome inactivating protein from Aspergillus giganteus, on tumor cell lines: Protein biosynthesis inhibition and cell binding. Mol. Cell. Biochem. 122, 39-47. Vazquez, D. (1979). "Inhibitors of Protein Biosynthesis," pp. 80-81. Springer-Verlag, Berlin. Wawrzynczak, E., Henry, R. V., Cumber, A., Parnell, G., Derbyshire, E., and Ulbrich, N. (1991). Biochemical, cytotoxic and pharmacokinetic properties of an immunotoxin composed of a mouse monoclonal antibody Fib75 and the ribosome-inactivating protein c~-sarcin from Aspergillus giganteus. Eur. J. Biochem. 196, 203-209. Wnendt, S., Felske-Zech, H., Henze, P., Ulbrich, N., and Stahl, U. (1993). Characterization of the gene encoding o~-sarcin, a ribosomal-inactivating protein secreted by Aspergillus giganteus. Gene 124, 239-244. Wool, I. G. (1986). Studies of the structure of eukaryotic (mammalian) ribosomes. In "Structure, Function, and Genetics of Ribosomes" (B. Hardesty and G. Kramer, eds.), pp. 391-411. Springer-Verlag, New York.
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Wool, I. G., Gltick, A., and Endo, Y. (1992). Ribotoxin recognition of ribosomal RNA and a proposal for the mechanism of translocation. Trends Biochem. Sci. 17, 266-269. Yang, R., and Kenealy, W. (1992). Regulation of restrictocin production in Aspergillus restrictus. J. Gen. Microbiol. 138, 1421-1427. Yang, X., and Moffat, K. (1996). Crystal structure of the highly specific Aspergillus ribotoxin, restrictocin. Structure 4, 837-852.
5 Plant Ribonucleases P A U L I N E A. B A R I O L A A N D P A M E L A J. G R E E N Department of Energy Plant Research Laboratory Department of Biochemistry Michigan State University East Lansing, Michigan, 48824-1312
I. Introduction II. Classes of Plant RNases A. Classifications Based on Early Biochemical Work B. Plant RNases in the T2 Family C. Pathogenesis-Related Protein Group PR-10 D. Group V Allergens E. Plant Nucleases III. Regulation and Functions of Plant RNases A. Phosphate Remobilization B. Senescence C. Cell Death Pathways D. Defense against Pathogens E. RNA Processing and Decay IV. Conclusions and Future Prospects References
I. I N T R O D U C T I O N
During the past few years, the study of plant ribonucleases (RNases) has advanced considerably due to renewed interest in the field and to technological advances. Prior to the early 1980s, the regulation and biochemistry of plant RNases were actively investigated as reviewed by RIBONUCLEASES: STRUCTURES AND FUNCTIONS
163 Copyright 9 1997by AcademicPress, Inc. All rights of reproduction in any form reserved.
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Farkas (1982) and Wilson (1982), but subsequently interest in the field subsided because the methods available at the time were insufficient to elucidate the biological functions of individual enzymes. Much renewed interest was prompted by the discovery that genotype-specific ribonucleases, the S-RNases, are critical components of self-incompatibility (SI) mechanisms in some solanaceous plants (McClure et al., 1989b; Lee et al., 1994). These findings, which exploited gene cloning and transgenic plant technologies, emphasized that plant RNases could participate in diverse and unexpected processes. Moreover, the discovery of the SRNases led to the identification of a large number of related plant RNases that do not function in self-incompatibility, called the S-like RNases. The goal of this chapter is to highlight current knowledge of the Slike RNases and other RNA-degrading enzymes that are not associated with self-incompatibility in higher plants. The S-RNases are reviewed in Chapter 6 of this volume. Rather than attempt a comprehensive review, this chapter concentrates on findings of the past decade with emphasis on molecular analyses and work reported since the last review of this topic (Green, 1994). Our discussions are divided into sections dealing with the classes of plant RNases and with their regulation and functions. When possible, recent findings are related to those in the early literature and the most promising prospects for future study.
II. C L A S S E S O F P L A N T R N a s e s
A. Classifications Based on Early Biochemical Work
Before the widespread use of molecular biology and protein microsequencing, plant RNases were extensively characterized on the basis of their biochemical properties. An abundance of reports on RNases from a variety of plants facilitated the classification of plant RNases into four main groups: RNase I, RNase II, nuclease I, and exonuclease I (Farkas, 1982; Wilson, 1982). RNase I proteins, or acid RNases, are RNA-specific, soluble endonucleases with molecular masses from 20 to 25 kDa and pH optima between 5.0 and 6.0. They are insensitive to E D T A and produce 3'-phospho (3'-P) nucleotides as end products. RNase II enzymes are also RNA-specific endonucleases with molecular mass between 17 and 25 kDa. They, too, are EDTA-insensitive and
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produce 3'-P nucleotide end products, but unlike RNase I enzymes, they have pH optima of 6.0 to 7.0. They differ most from RNase I-type enzymes by their microsomal location. Both RNase I and RNase II enzymes preferentially cleave bonds adjacent to guanine. Nuclease I proteins degrade both RNA and ssDNA endonucleolytically, with a preference for bonds adjacent to adenine, and produce 5'-P nucleotide end products. Highly sensitive to EDTA, they have molecular masses of 31 to 39 kDa and pH optima of 5.0 to 6.5. Finally, exonuclease I enzymes are large exonucleases of more than 100 kDa, are capable of degrading both RNA and ssDNA, have pH optima of 7.0 to 9.0 and a high sensitivity to EDTA, and produce 5'-P end products. The above designations are not comprehensive and did not include all known RNases even at the time of their publication (Farkas, 1982; Wilson, 1982). The classification of RNases was further complicated by common problems such as proteolytic degradation and aggregate formation during purification, as well as inconsistency in the assay procedures used in different laboratories. In addition, not every defining characteristic is investigated for each RNase reported. Despite these problems, many RNases reported more recently fit well into one of the four groups. Both Arabidopsis thaliana (Yen and Green, 1991) and barley (Yen and Baenziger, 1993) appear to have representatives of the RNase I, RNase II, and nuclease I classes, based on the properties of the RNAdegrading enzymes observed using activity gels. Additional reports from barley describe RNase I-type enzymes (Prentice and Heisel, 1985; Kenetick and Blake, 1986) and nucleases (Prentice and Heisel, 1986; Brown and Ho, 1986). Other RNase I-type enzymes have been identified in wheat (Blank and McKeon, 1991a) and tomato (Abel and Glund, 1987). A protein from wheat is a recent addition to the RNase II family (Yen and Baenziger, 1993), and nucleases have been found recently in wheat (Kuligowska et al., 1988; Blank and McKeon, 1989; Yen and Baenziger, 1993), rye (Siwecka et al., 1989), zinnia (Thelen and Northcote, 1989), and lentil (Kefalas and Yupsanis, 1995).
B. Plant RNases in the T2 Family
The family of plant RNases best characterized is a subset of the family of enzymes typified by the fungal RNase T2 (reviewed in Chapter 3 of this volume). This family is the most widespread RNase family known,
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with representatives in viruses (Schneider et al., 1993; Hime et al., 1995), bacteria (Meador and Kennell, 1990; Favre et al., 1993), fungi (reviewed in Chapter 3 of this volume), slime mold (Inokuchi et al., 1993), D r o s o p h ila (Hime et al., 1995), oyster (Watanabe et aL, 1993), cow (Irie, 1993), and chicken (Irie, 1993). Plant members of this family were first identified when sequences of proteins linked to gametophytic self-incompatibility in the Solanaceae family (reviewed in Chapter 6 of this volume; see also Kao and Huang, 1994) were determined and found to be similar to fungal ribonucleases (McClure et al., 1989b). The identification of these proteins, termed S-RNases, led to the discovery of related proteins in a variety of self-compatible plant species. Based on their similarity to SRNases, the latter group of enzymes was referred to as "S-like RNases" (Taylor et al., 1993). The S-like RNases have the two conserved histidine residues shown to be necessary for catalysis in RNase Rh, a related fungal RNase (Ohgi et al., 1992), and the five boxes of conserved sequence characteristic of the S-RNases (Ioerger et al., 1991), although these boxes in the S-like RNases tend to be less highly conserved. In general, the S-like RNases have molecular masses between 21 and 29 kDa, and most have been shown or predicted to be secretory proteins. Although the S-RNases and S-like RNases share many structural features (Green, 1994), each group contains highly conserved residues not found in the other (Taylor et al., 1993). In fact, gene genealogies of these RNases indicate that most S-like RNases form a lineage distinct from that of the S-RNases (Bariola et al., 1994). Initially the term "S-like RNases" was used to describe RNases that are structurally similar to S-RNases but found in species not known to exhibit self-incompatibility. However, as more of these proteins were found in a wide variety of plant species, it became apparent that S-like RNases are likely to play roles distinct from selfincompatibility, but fundamental to plants in general. Based on this hypothesis, self-incompatible plants would be expected to contain proteins of both the S-RNase and S-like RNase families. This is now known to be the case, as discussed below. In addition, some species that exhibit self-incompatibility contain proteins very closely related to S-RNases that are not involved in the self-incompatibility of the plant, as will be discussed. These enzymes lack the distinguishing structural features associated with S-like RNases and appear to have arisen as part of the S-RNase lineage after it diverged from the S-like RNase lineage. To avoid confusion, we propose that S-like RNases be defined as proteins from self-incompatible or self-compatible plants whose structures allow their placement into the evolutionary lineage shown in Fig. 1.
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Plant Ribonucleases LC1 MC1 LC2 LX ZE1 LE NE RNS1 ZE2 RNS3 NNUC RNS2
Fig. 1.
Gene genealogy of the S-like RNases. The Genetics Computer Group program
PILEUP (Devereux et al., 1984) was used as described (Taylor et al., 1993) to create the dendrogram. RNase abbreviations are as follows: LC1 and LC2 are from Luffa cylindrica (Nakamura et al., 1996); MC1 is from Momordica charantia (Ide et al., 1991); LX (LOftier et al., 1993) and LE (Jost et al., 1991) are from Lycopersicon esculentum; NE is from Nicotiana alata (Dodds et al., 1996); ZE1 and ZE2 are from Zinnia elegans (Ye and Droste, 1996); RNS1, RNS2, and RNS3 are from Arabidopsis thaliana (Taylor et al., 1993; Bariola et al., 1994); and NNUC is from Nelumbo nucifera (G. Day, Z. Chen, and T. Chow, unpublished, GenBank accession number M83668).
The dendrogram shown in Fig. 1 is the result of a computer analysis of the relatedness of all known Tz-type proteins for which full sequences are available. Interestingly, the plant S-like RNases form a lineage separate from that of the S-RNases and other Tz-type proteins from viruses, bacteria, fungi, and animals, although only the S-like lineage is shown in Fig. 1. Three cucurbit S-like RNases, MC1, LC1 and LC2, have formed a separate sublineage within the S-like RNase group. It is possible that their divergence from the other S-like RNases represents a difference in function rather than evolutionary distance. This idea is based on a molecular systematic model of plant relatedness in which Arabidopsis is more closely related to the Cucurbitaceae than to the Solanaceae or Asteraceae (Meagher et al., 1989), two of the other families that contribute proteins to the S-like RNase lineage. Comparison of the amino acid sequences of the proteins that form the S-like RNase lineage to those of the S-RNases reveals numerous residues that are conserved in S-like RNases but not in S-RNases (Green, 1994). Conserved residues unique to the S-like RNases are mainly found between the conserved boxes
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that contain the two active-site histidine residues, as well as at the Nterminal end (see Green, 1994). The first S-like RNase genes to be identified were RNS1, RNS2, and R N S 3 of Arabidopsis thaliana (Taylor and Green, 1991). These were cloned via the polymerase chain reaction (PCR) based on their containing the conserved active site regions of the T2 family. The initial PCR products isolated in this reaction contained several residues conserved in the S-RNases. When the full sequences of their cDNAs were determined, however, several differences from the S-RNases were apparent (Taylor et aL, 1993; Green, 1994), laying the foundation for the designation of the S-like RNases as a separate group. One of the R N S genes, RNS2, is expressed in all organs examined, with highest expression in flowers (Taylor et al., 1993). Preliminary immunocytological evidence indicates that RNS2 is extracellular (P. A. Bariola and P. J. Green, unpublished); RNS1 and RNS3 are also expected to be extracellular because they appear to contain typical N-terminal signal sequences for entry into the secretory pathway (Bariola et aL, 1994). RNS1 and RNS3 are quite closely related to RNases of the selfcompatible tomato Lycopersicon esculentum, LE (Jost et aL, 1991) and LX (LOftier et al., 1993) (see Fig. 1), both of which were isolated from cell cultures due to their appearance on starvation for phosphate (Pi) (Ntirnberger et al., 1990; LOftier et aL, 1992). Three other Pi-starvation induced RNases from tomato (LOftier et aL, 1992) have also recently been sequenced at the protein level (K0ck et aL, 1995). These enzymes, designated LV1, LV2, and LV3, were isolated from tomato vacuoles. Interestingly, LV3 appears to be identical to RNase LE, an extracellular protein, and the regions of LV1 and LV2 that have been sequenced are identical to sequences of RNase LX. The latter enzyme has been shown to have an intracellular but extravacuolar location (LOftier et al., 1992). RNase LX contains a putative C-terminal endoplasmic reticulum (ER) retention signal (LOftier et al., 1993), which is missing in the LV2 protein sequence (K0ck et aL, 1995). This observation may indicate that in the absence of this putative ER retention signal the protein is targeted to the vacuole. It has not been reported whether these proteins sharing long sequences are products of the same genes. Whether or not this is the case, regulating the location of these RNases could be a novel mechanism for control of RNase activities and possibly RNase functions. Another member of the Solanaceae, the self-compatible species Nicotiana sylvestris, was found to contain a stylar RNase with homology to the T2 family (Golz et al., 1997). Tubers from the lotus species N e l u m b o
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nucifera gave rise to another S-like RNase, identified as a storage protein (G. Day, Z. Chen, and T. Chow, unpublished, GenBank accession number M83668). Two S-like RNases were identified in cultured zinnia leaf mesophyll cells undergoing xylogenesis (Ye and Droste, 1996). These genes, Z R N a s e I and ZRNaselI, are very closely related to the tomato and Arabidopsis RNases described above, but have different expression patterns; neither is induced in response to Pi starvation (Ye and Droste, 1996). Other studies have shown that the seeds of several cucurbit species contain S-like RNases. The first of these enzymes to be identified was RNase MC1 from the seeds of the bitter gourd Momordica charantia (Ide et al., 1991). Subsequently two RNases were isolated on the basis of their translational inhibitory properties in cell-free systems: cusativin, from cucumber seeds (Rojo et al., 1994a), and melonin, from seeds of the melon Cucumis melo (Rojo et aL, 1994b). Both of these enzymes were found to have sequences characteristic of S-like RNases. Cusativin is known to accumulate only in the coat and cotyledons of dry seeds (Rojo et al., 1994a). Two related S-like RNases, LC1 and LC2, have been cloned from the seeds of Luffa cylindrica, the sponge gourd (Nakamura et aL, 1996). Because all of the above cucurbit species are self-compatible it was suggested that the RNases play a role in protection of the seeds against pathogens (Rojo et al., 1994a; Nakamura et aL, 1996). It is likely that S-like RNases will be found to be widespread among monocotyledonous plants as well. This is evidenced by the recent identification in rice, which is self-compatible, of several cDNA sequences corresponding to S-like RNases by investigators participating in the Rice Genome Project (GenBank accession numbers D21885, D22272, D23641, and D24884). Based on these partial sequences it is not yet clear whether monocot S-like RNases will form their own sublineage within the S-like RNase lineage. Interestingly, self-compatible species have been found to contain proteins structurally similar to the S-RNases. Two RNases, Sx and So, have been identified in a self-compatible cultivar of Petunia hybrida (Ai et al., 1992). Both their structural similarity to the S-RNases and breeding behavior suggest that these RNases are defunct S-RNases, possibly selected for in the breeding process that generated this self-compatible cultivar from its self-incompatible ancestors (Ai et al., 1992). A selfcompatible variant of the potato Lycopersicon peruvianum contains a protein, So, whose sequence identifies it as a member of the S-RNase family, but which lacks RNase activity (Royo et al., 1994). A mutation
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of one of the highly conserved histidines in RNase Sc is thought to be responsible for the lack of activity and thus the self-compatibility of this normally self-incompatible plant. Like RNases Sx and So, the Sc protein is structurally more closely related to the S-RNases than the S-like RNases. Several T2-related enzymes other than the S-RNases have also been identified in self-incompatible plants. RNase NE, an S-like RNase with a sequence quite similar to those of RNase LE of tomato and RNS1 of Arabidopsis, was identified via PCR in anthers of Nicotiana alata (Dodds et al., 1996, GenBank accession number U13256). Similarly, styles of N. alata contain a member of the T2 family, RNase MS1, thought to be unassociated with self-incompatibility (Kuroda et al., 1994), but the sequence of this enzyme has not been reported so it is unclear if this protein is an S-RNase or an S-like RNase. Other members of the T2 family in self-incompatible plants bear stronger resemblance to the S-RNases, such as RNase X2 from Petunia inflata (Lee et al., 1992). Although abundant in pistils, the protein is not associated with selfincompatibility. RNase X2 may have diverged from the S-RNases, or alternately evolved from a common ancestor (Lee et al., 1992). Few of the plant RNases described in this section have been categorized according to the traditional biochemical classifications referred to in Section II,A. However, RNases LE, LX, LV1, LV2, and LV3 from tomato are all considered RNase I-type enzymes, based on their biochemical properties (NUrnberger et aL, 1990; Jost et al., 1991; LOftier et al., 1992, 1993). Most of the S-like RNases described in this section have molecular masses in the range of 20 to 25 kDa, as specified for RNase I-type enzymes, but RNS2 from Arabidopsis, with a deduced molecular mass of 27.1 kDa and two potential N-glycosylation sites (Taylor et al., 1993), is an exception. The tertiary structure of the S-like RNases has been investigated to a limited extent. Preliminary observations indicate that the tertiary structure of RNase LE (M. Irie, M. K0ck, and K. Glund, unpublished) exhibits several differences from that of RNase Rh from the fungus R h i z o p u s niveus (Kurihara et al., 1992), the only other member of the T2 family whose tertiary structure is currently available. In addition, RNase MC1 has been crystallized (De and Funatsu, 1992), so additional data on this enzyme may be forthcoming.
C. Pathogenesis-Related Protein Group PR-10 Pathogenesis-related proteins (PR proteins) are enzymes induced in plants upon pathogen attack or in related situations (van Loon et al.,
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1994), and presumably have roles in the defense of plants against pathogens. The numerous families of PR proteins include such diverse members as chitinases (Legrand et al., 1987), glucanases (Kauffmann et al., 1987), and proteinase inhibitors (Geoffroy et al., 1990). PR proteins have been linked to RNases with the report that a ginseng RNase with nonspecific activity (Moiseyev et al., 1994) has protein sequence homology to two PR proteins from parsley (Somssich et al., 1988; van de L6cht et al., 1990). The parsley proteins in turn are known to be members of a large PR protein family designated as PR-10 (van Loon et al., 1994; Swoboda et al., 1995b and references therein). The members of this family are present in a variety of plants, have molecular masses of 17 to 18 kDa, and are considered intracellular. In addition, proteins of a related group, the major pollen allergens in birch, the Bet v 1 family, are constitutively present at high levels in pollen (Swoboda et al., 1994). These pollen allergens constitute a large isoform family in birch and other plants (Swoboda et al., 1995a). Besides the ginseng RNase preparation (Moiseyev et al., 1994), which has been shown to be a mixture of two related proteins (G. Moiseyev and J. Beintema, unpublished), the only other protein in the PR-10 family that has been reported to have RNase activity is Bet v 1, reported in two separate papers (Swoboda et al., 1996; Bufe et al., 1996). Efforts to demonstrate activity for members of this family in potato (Constabel and Brisson, 1995), parsley (I. Somssich, unpublished), and asparagus (S. A. J. Warner and J. Draper, unpublished) have been unsuccessful. Until more proteins of this type are shown to exhibit RNase activity, it may be premature to designate this family as another major family of plant RNases. However, if activity can be demonstrated in other PR-10 proteins, this family would be the first group of intracellular, nonspecific RNases characterized molecularly in plants. In addition, it may then be possible to establish a direct link between specific RNase activities and plant defense against pathogens.
D. Group V Allergens
RNase activity has also been associated with a protein from timothy grass, P h i p Vb (Bufe et al., 1995), a member of a group of grass pollen allergens molecularly distinct from the Bet v 1 family described above. These proteins, the group V allergens, generally have molecular masses of 32 to 38 kDa, and include proteins targeted to the amyloplasts in rye
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grass (Singh et al., 1991; Knox, 1993). However, the RNase activity of Phl p Vb should be interpreted with caution, because the activity is inhibited by human placental RNase inhibitor, which was previously found to inhibit only animal RNase A-type proteins among RNase groups previously tested (Chapter 4 in this volume; also see Lee and Vallee, 1993). The sequences of group V allergens exhibit no readily apparent similarity to proteins in the RNase A superfamily. [Similar concerns about the use of human placental RNase inhibitor are also associated with the cucumber S-like RNase cusativin (Rojo et al., 1994a), as well as with one of the Bet v 1 studies (Bufe et al., 1996), because the PR-10 proteins also have no obvious sequence similarities to RNase.] Whether the group V allergens comprise another major RNase family has yet to be determined.
E. Plant Nucleases
Although many plant enzymes with the characteristics of nuclease I enzymes have been identified, little sequence information is available to confirm the relatedness of these proteins. However, there is a limited region of sequence identity between two nucleases from distantly related plants. One of these proteins is a 39-kDa nuclease I secreted from barley aleurone layers (Brown and Ho, 1987). The secretion of this nuclease is induced by gibberelic acid (Brown and Ho, 1986), a plant hormone that induces the secretion of a range of hydrolytic enzymes from aleurone layers to mobilize seed endosperm reserves for the germinating seedling (Jacobsen et al., 1995). Another nuclease, from zinnia, is induced during the differentiation of xylem elements (Thelen and Northcote, 1989). Although the zinnia nuclease cannot be strictly classified as a nuclease I (Wilson, 1982) due to its 43-kDa molecular mass, its N-terminal sequence is similar to that of the barley nuclease, implying that these two enzymes are related. It has been noted (Fraser and Low, 1993) that the partial barley nuclease sequence is similar to the amino termini of S1 and P1 nuclease from the fungi Aspergillus oryzae ( I w a m a t s u et al., 1991) and Penicillium citrum (Maekawa et al., 1991), respectively. These fungal nucleases are part of a large family of single-strand-specific bifunctional nucleases (Gite and Shankar, 1995) based on their biochemical properties. Mung bean nuclease (Laskowski, 1980) is also considered to be a member of this family. The biochemical characteristics that define this
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group (Gite and Shankar, 1995) are much broader than those that define the plant nuclease I enzymes (Wilson, 1982), and nuclease I enzymes could easily be classified as part of the single-strand-specific nuclease family. [See Gite and Shankar (1995), for an extensive discussion of the biochemical properties of the enzymes of this family.] In Arabidopsis, nucleases shown to have some of the properties of nuclease I enzymes have been identified using activity gels (Yen and Green, 1991). A doublet of about 33 kDa that appears on both RNase and DNase activity gels led to the suggestion that the same 33-kDa enzymes can degrade both RNA and DNA. This idea was confirmed through the analysis of altered RNase profile (arp) mutants of Arabidopsis. Six arp mutants that either lack or overproduce one or both of the 33-kDa doublet RNase activities were examined on DNase activity gels and shown to exhibit phenotypes identical to those on RNase activity gels (Abler and Green, 1997). This result demonstrates genetically that the 33-kDa doublet represents a pair of bifunctional nuclease activities. The availability of these mutants may help elucidate the biological functions of these nucleases and reveal whether they are encoded by the same gene. In addition to the nuclease I enzymes mentioned earlier (Section II,A), a number of activities with some similarities to a tobacco pollen nuclease I (Matousek and Tupy, 1984) have been identified in pollen from various species, with a nuclease from Pinus nigra being the best characterized (Matousek and Tupy, 1985). Likewise, a nuclease with similar properties was identified in tobacco anthers (Matousek and Tupy, 1987). Singlestrand-specific nucleases have recently been found in spinach (Strickland et al., 1991), scallion (Uchida et aL, 1993), wheat chloroplasts (Monko et al., 1994), and pea seeds (Naseem and Hadi, 1987) and chloroplasts (Kumar et al., 1995). It will be interesting to determine if the nucleases described in this section, the plant enzymes classified as nuclease I mentioned in Section II,A, and the single-strand-specific nucleases from fungi and plants are members of the same or multiple molecular families.
III. R E G U L A T I O N AND FUNCTIONS OF P L A N T R N a s e s
A. Phosphate Remobilization Several S-like RNases have been shown to be up-regulated in response to starvation for inorganic phosphate (Pi). The first of these reports
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showed that RNase LE from tomato is secreted in response to P~ limitation (Ntirnberger et aL, 1990). Subsequently, increased levels of the other tomato S-like RNases, LX, LV1, LV2, and LV3, all intracellular enzymes, were found during Pi starvation (L6ffler et al., 1992). Most of this work was done at the protein level, but recently starvation for this nutrient has been shown to induce RNases LE and LX at the m R N A level (K6ck et aL, 1995). These results mirrored previous reports of two Arabidopsis RNase genes, R N S 1 (Bariola et al., 1994) and R N S 2 (Taylor et al., 1993), which were also found to be Pi-starvation inducible. In particular, the R N S 1 m R N A is dramatically up-regulated from a low basal level (Bariola et al., 1994). The induction of R N S 1 also appears to occur at the protein level, because an RNase activity that comigrates with RNS1 produced in yeast increases in parallel with R N S 1 m R N A (Bariola et al., 1994; Howard et aL, 1997). In addition, P~ starvation has recently been found to induce the gene for N. alata RNase NE (Dodds et aL, 1996). However, at least one S-like RNase gene, R N S 3 of Arabidopsis, does not respond to Pi limitation, so Pi-starvation inducibility should not be considered characteristic of the S-like RNases. Limitation for P~ also leads to a large increase in the activity of a surface membrane-associated nuclease in the trypanosome Crithidia luciliae (Gottlieb et al., 1988). Although the sequence of this protein is not available, the nuclease is considered similar to plant nuclease I enzymes due to its enzymatic properties (Neubert and Gottlieb, 1990). It is conceivable that plant nuclease I genes could also be induced by Pi starvation. Indeed, extracts of plants grown on P~-deficient medium exhibit increased RNase activity on activity gels in the area of the 33-kDa nuclease doublet (refer to Bariola et al., 1994). It has been proposed that under P~-limiting conditions, RNases could degrade RNA in conjunction with phosphatases and phosphodiesterases to release P~, making it available for the plant to use in other processes (Glund and Goldstein, 1993). This response is likely only a part of a broader effort by the plant to optimize Pi availability under conditions of scarcity (Goldstein et al., 1989). RNases could increase the efficiency of the plant to scavenge P~ in several ways. RNases selected from roots into the soil could make Pi previously sequestered in R N A in organic matter in the soil available for uptake. The role of scavenging P~ from RNA in the growth environment is believed to be a main function of two fungal nucleases (Fraser and Low, 1993). Extracellular RNases within the plant could rescue Pi from RNA that arises in the extracellular space from cells that have lysed due to senescence, damage, or programmed
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cell death (discussed below). Finally, vacuolar RNases may participate in some aspect of intracellular RNA degradation, because short pieces of RNA exist in the vacuoles of plant cells (Abel et al., 1990). It is generally recognized that one main function of the plant vacuole is the turnover of cellular macromolecules, analogous to animal lysosomes (Boiler and Wiemken, 1986). Because autophagy of small amounts of cytoplasm may occur in plant cells (Boiler and Wiemken, 1986), this process may be a route for vacuolar uptake of cytoplasmic RNA. An increase in the RNase concentration in the vacuole could speed the process of the recycling of components of RNA. Remobilization of Pi may take place during normal plant development. In barley seeds, a nuclease I is secreted from the aleurone layer upon treatment with gibberellin (Brown and Ho, 1986), a hormone associated with seed germination (Jacobsen et aL, 1995). The aleurone layer lays outside the endosperm, which stores macromolecules such as proteins and carbohydrates. Germination triggers the secretion of several hydrolytic enzymes (Jacobsen et aL, 1995), which release nutrients in forms able to be used quickly by germinating seedlings. The nuclease I is proposed to degrade nucleic acids in endosperm, in conjunction with acid phosphatases, to release nucleosides and phosphate for use in new RNA synthesis in the seedling (Brown and Ho, 1986). The widespread increase in RNase activities in germinating seeds (Farkas, 1982) suggests that this may be a universal phenomenon in plants.
B. Senescence
The effect of senescence on plant RNase activity has been studied extensively. In general, RNase activities increase in plants during senescence, but in different systems the timing and extent of the increases vary (Farkas, 1982). Many of these variations are likely due to differences in systems and experimental design (e.g., studies in attached leaves or excised leaves), but different patterns of induction may also contribute (Farkas, 1982). It is now clear that senescence dramatically up-regulates individual RNase activities and genes. In wheat leaves, single-strandspecific nuclease activity increases during senescence (Blank and McKeon, 1989), as does the activity of three RNases of 20 to 27 kDa (Blank and McKeon, 1991b), monitored both biochemically and in activity gels. The three Arabidopsis S-like RNase genes, RNS1, RNS2, and
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RNS3, are each induced to different extents in leaves during senescence: R N S 1 mRNA levels increase only slightly (Bariola et al., 1994), whereas those of R N S 2 and R N S 3 increase more dramatically (Taylor et al., 1993; Bariola et al., 1994). R N S 2 is also known to be induced in senescing petals (Taylor et al., 1993).
During senescence, cellular structures are disassembled and macromolecules in certain plant organs are broken down, freeing nutrients for relocation to other organs (Stoddart and Thomas, 1982). This process is thought to occur to conserve minerals and nutrients for reuse. The breakdown and redistribution can occur both during vegetative growth, such as the rescue of minerals from senescing cotyledons, and also during reproductive growth, when in some plants all organs except the reproductive structures senesce and the vegetative organs serve as a source of nutrients for the reproductive structures (Nood~n, 1988). RNases are likely included in the variety of hydrolytic enzymes induced during senescence (Borochov and Woodson, 1989) that facilitate the breakdown of cellular components. The actions of RNases during senescence could lead to the recycling of P~.
C. Cell Death Pathways
Programmed cell death is an essential part of developmental patterns and physiological processes in many organisms (Vaux, 1993). In plants, cell death pathways are only beginning to be investigated (Greenberg, 1996). Programmed cell death is associated with sex determination in maize (DeLong et al., 1993), the hypersensitive response (HR) against plant pathogens (Greenberg et al., 1994), and possibly pollination, senescence, and various developmental processes (Greenberg, 1996). One of the best studied processes regarding cell death in plants is the differentiation of xylem, a major component of the plant vascular system. When isolated leaf mesophyll cells from zinnia are cultured in the presence of appropriate concentrations of the hormones auxin and cytokinin, they differentiate synchronously into elongated, lignified tracheary elements, the building blocks of the xylem (Fukuda, 1992). The last part of the differentiation process involves strengthening of the cell wall and hydrolysis of the end walls between two differentiating cells to form a tube. Vacuoles lyse several hours after the secondary cell wall is formed, leading to degradation of cytoplasmic macromolecules. Finally, lysis of
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the protoplast occurs so that the cell wall can serve as a link in the channel that forms the xylem (Fukuda, 1992). A single-strand-specific nuclease of 43 kDa with homology to a nuclease I of barley (Brown and Ho, 1986) appears during xylogenesis in the zinnia system (Thelen and Northcote, 1989). In addition, several RNase activities of 17 to 25 kDa appear in zinnia cell extracts, and a 37-kDa nuclease accumulates in the culture medium. One of these RNases may correspond to ZRNase I, an S-like RNase with a predicted molecular mass of 27 kDa, whose cDNA was isolated from zinnia mesophyll cells differentiating into xylem elements (Ye and Droste, 1996). In zinnia cultured mesophyll cells, its mRNA first appears at high levels after about 48 hr of culture in differentiation-inducing medium, relatively late in the differentiation process (Ye and Droste, 1996). This result was confirmed by tissue-print hybridization, in which Z R N a s e I m R N A appears in differentiating xylem elements of stems (Ye and Droste, 1996). A protease gene is also strongly induced at 48 hr of culture (Ye and Varner, 1993). It is possible that these and other hydrolytic enzymes are involved either in killing the cell directly or in degrading cytoplasmic components during and after lysis in order to clear the xylem channel and facilitate nutrient reutilization. Nucleases could be involved in RNA degradation as well as the fragmentation of DNA, which is one of the hallmarks of apoptotic cell death in animal systems (Zhivotovsky et al., 1994). Nuclear DNA fragmentation has been observed in pea root xylem cells undergoing cell death (Mittler and Lam, 1995a). Less is known about the association of RNases and nucleases with other cell death pathways in plants. However, recent studies indicate that anther nucleases are highest during the first microspore division in tobacco (Matousek et al., 1994). This led to the suggestion that anther nucleases could participate in tapetal cell degeneration (Matousek et al., 1994).
D. Defense against Pathogens Increases in RNase activities in diseased plants are well-documented (Farkas, 1982; Green, 1994). Several roles can be proposed for RNases in plant disease and defense. First, if RNases are involved in cell death pathways in plants, as discussed in Section III,D, they could play a role in the hypersensitive response of plants, which appears to involve programmed cell death (Greenberg et al., 1994). The HR involves death
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of plant cells shortly after pathogen infection in the immediate vicinity of the infection site, and the localized cell death is thought to contribute to the resistance of the plant to the disease (Keen, 1992). Recently, a DNase activity has been found to be induced in tobacco nuclei during cell death due to the HR (Mittler and Lam, 1995b). This DNase, NUCIII, has been characterized as an endonuclease that cleaves both singlestranded and double-stranded DNA, but it has not been reported whether the protein has RNase activity (Mittler and Lam, 1995b). It seems probable that D N A fragmentation occurs during the HR and the NUCIII could play a role in this process. In tobacco, an HR-like response that was for some reason induced by overexpression of a bacterial proton pump gene provided evidence for D N A fragmentation (Mittler et al., 1995). A second role for plant RNases during pathogen attack could be to act as defense proteins in tissues potentially susceptible to infection. For example, the pistil, which is penetrated by the pollen tube during the fertilization process, is rich in extracellular nutrients that should make it susceptible to pathogen invasion. However, it is rarely infected. This resistance may be due to defense-related proteins that are present extracellularly in the pistil, such as proteinase inhibitors (Atkinson et al., 1993). Several T2-type RNase genes have been shown to be expressed in pistils, so they are also candidate defense proteins. These include X 2 of Petunia inflata (Lee et al., 1992), R N S 2 of A r a b i d o p s i s (Taylor et aL, 1993), and N E of Nicotiana alata (Dodds et al., 1996). It has been proposed that gametophytic self-incompatibility may have arisen via the recruitment of defense-related pistil RNases (Lee et al., 1992). Extracellular RNases may also play a part in the defense by plants against RNA viruses. Finally, RNases are known to accumulate in plant vacuoles (Farkas, 1982; Wilson, 1982). The defense-related proteins chitinase and 13-1,3-glucanase, which can degrade fungal cell walls, are also known to be sequestered in vacuoles, increasing in abundance during pathogen attack (Mauch and Staehelin, 1989). Mauch and Staehelin (1989) propose a model in which accumulation of large amounts of these proteins in vacuoles is an advantage: when fungi invade, cells lyse due to either the HR or the direct pathogen invasion of the cell. On lysis, fungal hyphae would be flooded with defense-related proteins in high enough concentrations to lyse the hyphae. RNases may well be components of this onslaught of hydrolytic defense enzymes released from the vacuoles on cell lysis. Plants interpret mechanical wounding as a signal of attack, because pathogen infection and chewing by insects or other herbivores often
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result in wounding of tissues. Wounding induces defense-related genes in plants, such as those of proteinase inhibitors (Ryan, 1990), peroxidases (Bowles, 1990), and chitinases (Bowles, 1990). Wounding is known to induce rapid increases in RNase activities (Farkas, 1982). One zinnia Slike RNase gene, ZRNasell, is rapidly induced on mechanical wounding (Ye and Droste, 1996). In Arabidopsis, wounding of stems resulted in induction of an RNase activity of about 34 kDa (M. Saitoh, M. L. Abler, and P. J. Green, unpublished). Like the 33-kDa bifunctional nuclease activities discussed in Section II,D, the 34-kDa RNase comigrates with a DNase activity induced under the same conditions, so the protein is likely a bifunctional nuclease. Further examination of this enzyme and its wound inducibility may provide insights into the roles of RNases in defense responses.
E. RNA Processing and Decay 1. Nuclear Activities A number of events that take place in plant nuclei likely involve RNases, but very few such enzymatic activities have been identified. Presumably the major degradative process in the nucleus is the decay of introns and other sequences removed from precursors of mature mRNAs, rRNAs, and tRNAs. There are a few early reports of nuclearassociated RNase activities that might be involved in these processes in plants (reviewed in Farkas, 1982), but this area of research should be revisited because it is underdeveloped not only in plants but also in other eukaryotes (Stevens, 1993; Ross, 1995). The most convincing data for an RNase located in the nucleus comes from work on the 7-2/MRP RNA, which is known to be the RNA component of RNase MRP in mammalian cells. In plants and mammalian cells, 7-2/MRP RNA is found in nucleoli, where it is likely to be involved in rRNA processing (Kiss and Filipowicz, 1992; Kiss et al., 1992; Morrissey and Tollervey, 1995).
2. Chloroplast and Mitochondrial Activities Most of the organellar RNase activities that have been identified participate in the maturation of 5' and 3' ends of chloroplast and mitochondrial transcripts such as tRNAs. The ribonucleoprotein RNase P is responsible
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for the processing of the 5' end of pre-tRNAs (Altman et al., 1993); in plants, RNase P enzymes have been identified in chloroplasts (Gegenheimer, 1996) and mitochondria (Marchfelder and Brennicke, 1994). Interestingly, the spinach chloroplast RNase P may not contain an RNA component (Wang et al., 1988; Gegenheimer, 1996). Maturation of the 3' end of pre-tRNAs in eukaryotes involves an endonucleolytic cleavage in most cases, in contrast to the prokaryotic mechanism (Deutscher, 1993). 3' tRNA processing activities are detectable in plant nuclei, mitochondria, and chloroplasts (Oommen et al., 1992; Marchfelder and Brennicke, 1994; Gegenheimer, 1996). Preliminary characterization of RNase activities that affect 3' end maturation has also been achieved (Chen and Stern, 1991). These include one or more 3' ~ 5' exoribonucleases and an endoribonuclease. The latter has been shown to cleave the spinach petD mRNA at the termination codon and at the mature RNA 3' end. Interestingly, cleavage of the petD mRNA at the termination codon leads to rapid degradation of the upstream RNA. In maize, a nuclear mutation, crpl, has been isolated that blocks the processing of the polycistronic precursor of petD mRNA, which appears to inhibit translation (Barkan et al., 1994). It is possible that the C R P I gene encodes a processing RNase or a protein that regulates such an activity. 3. Activities Implicated in Cytoplasmic mRNA Decay
Little is known about the RNase activities that facilitate the degradation of most plant mRNAs, because most degrade without generating easily identifiable intermediates. Two exceptions are the soybean rbcS (Tanzer and Meagher, 1994, 1995) and the P H Y A (Higgs and Colbert, 1994) mRNAs. Discrete fragments of the rbcS mRNA are observed in vivo and are produced in an in vitro decay system. The structures of these fragments suggest that they are produced by a stochastic endonuclease followed by exonuclease digestion in the 5'--* 3' or 3'--* 5' direction (Tanzer and Meagher, 1994, 1995). In contrast, a continuous population of lower molecular weight fragments is observed for the P H Y A mRNA, rather than discrete intermediates. Characterization of these fragments indicates that they most likely arise through the action of 5'--* 3' and 3' ~ 5' exoribonucleases, although endoribonuclease models cannot be ruled out (Higgs and Colbert, 1994). In yeast, many mRNAs are known to be degraded by a pathway involving deadenylation, possibly by a poly(A) nuclease, followed by decapping and digestion by XRN1, a
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5' --~ 3' exoribonuclease. This type of pathway may explain the decay of about 25% of the P H Y A mRNA, but the remainder (Higgs and Colbert, 1994), as well as the rbcS mRNA (Tanzer and Meagher, 1995), appears to degrade independent of deadenylation. Deadenylationindependent mRNA decay pathways, some of which involve cleavage by sequence-specific endoribonucleases (Brown and Harland, 1990; Binder et al., 1994), also exist in yeast and animal systems (Beelman and Parker, 1995). Recent reports have identified sequences that can trigger rapid decay of reporter mRNAs in plants. These include the 3' untranslated region (UTR) of the Arabidopsis S A U R - A C1 transcript (Gil and Green, 1996), a dimer of the DST element (Newman et al., 1993), which is conserved among the 3' UTRs of unstable S A UR transcripts (McClure et al., 1989a), and A U U U A repeats (Ohme-Takagi et al., 1993), which also trigger mRNA decay in mammalian cells (Shaw and Kamen, 1986; Vakalopoulou et al., 1991). However, it is unknown whether these sequences serve as endonuclease-sensitive sites in plants or whether they trigger exonuclease digestion. By examining the effect of these instability elements in the Arabidopsis mutants that alter the RNase profile (arp mutants, described in Section II,D) and other mutants that may become available, it may be possible to identify some of the RNases involved in general and specific mRNA decay pathways. In addition, in vitro systems (Byrne et al., 1993; Tanzer and Meagher, 1994, 1995) may facilitate purification of RNases that act on specific mRNA transcripts, particularly in the case of the P H Y A and rbcS transcripts. Finally, it has been suggested that antisense RNA effects may be mediated in part by the action of a double-stranded RNA-degrading activity that is presumed to degrade the sense-antisense hybrid (Nellen and Lichtenstein, 1993). This is consistent with the observation that in many cases accumulation of the sense RNA is diminished in plants engineered to produce a corresponding antisense RNA (Nellen and Lichtenstein, 1993; Bourque, 1995). In one case, this effect has been shown to be due to rapid decay of the sense RNA (Jiang et al., 1994). Although a considerable amount of dsRNase activity in plants may be extracellular (Matousek et al., 1994), perhaps the enzyme corresponding to an Arabidopsis-expressed sequence tag (Genbank accession number Z18464; HOfte et al., 1993), which has homology to RNase III, a bacterial intracellular dsRNase (Robertson, 1982), is involved in this process.
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IV. C O N C L U S I O N S A N D F U T U R E P R O S P E C T S
The relationships among different RNases have become much clearer now that many have been cloned and sequenced. It seems apparent that the S-like RNases are among the major, if not the major, class of R N A degrading enzymes in higher plants. Based on their gene expression and activity levels, the S-like RNases are likely to participate in fundamental physiological processes such as senescence, phosphate starvation responses, and cell death pathways. The next few years should be even more exciting as new members of this and other plant RNase families are examined and compared at the molecular level. Genetic approaches that offer tremendous promise are now being applied to the study of RNase function. Mutants with altered RNase profiles are being examined for their effects on m R N A decay and responses to the endogenous and exogenous stimuli that are known to affect the expression of RNase genes. Conversely, as more mutants are isolated with defects in m R N A decay, phosphate-starvation responses, and senescence and cell death pathways, they can be tested for alterations in known RNases. New and emerging questions will include how and where individual RNases are localized within and outside of cells, and whether the localization of any RNase is differentially regulated, for example, the tomato LV enzymes. It will also be important to explore the possible existence of proteinacious RNase inhibitors that might specifically interact with major plant RNases in a fashion analogous to the RNase inhibitors of the vertebrate RNase A superfamily of enzymes (reviewed in Chapter 9 of this volume). Beyond addressing these basic questions, future studies with plant RNases could also have applied significance. Already, male sterile maize (Mariani et al., 1990) and corresponding restorer lines (Mariani et al., 1992) have been created by expressing heterologous RNase and RNase inhibitor genes in appropriate cell types. By manipulation of endogenous RNase genes, it may be possible to engineer plants with other desirable traits that would lead to crop improvement.
ACKNOWLEDGMENTS
We are grateful to many colleagues who have provided reprints, preprints, and unpublished data, and to Dr. Michael Abler for comments on the manuscript. Work in the
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author's laboratory was supported by grants from the National Science Foundation (IBN9408052) and the Department of Energy (FG02-91-ER20021) to PJG.
REFERENCES
Abel, S., and Glund, K. (1987). Ribonuclease in plant vacuoles: Purification and molecular properties of the enzyme from cultured tomato cells. Planta 172, 71-78. Abel, S., Blume, B., and Glund, K. (1990). Evidence for RNA-oligonucleotides in plant vacuoles isolated from cultured tomato cells. Plant Physiol. 94, 1163-1171. Abler, M. L., and Green, P. J. (1997). Manuscript in preparation. Ai, Y., Tsai, D.-S., and Kao, T.-H. (1992). Cloning and sequencing of cDNAs encoding two S proteins of a self-compatible cultivar of Petunia hybrida. Plant Mol. Biol. 19, 523-528. Altman, S. Kirsebom, L., and Talbot, S. (1993). Recent studies of ribonuclease P. F A S E B J. 7, 7-14. Atkinson, A. H., Heath, R. L., Simpson, R. J., Clarke, A. E., and Anderson, M. A. (1993). Proteinase inhibitors in Nicotiana alata stigmas are derived from a precursor protein which is processed into five homologous inhibitors. Plant Cell 5, 203-213. Bariola, P. A., Howard, C. J., Taylor, C. B., Verburg, M. T., Jaglan, V. D., and Green, P. J. (1994). The Arabidopsis ribonuclease gene RNSI is tightly controlled in response to phosphate limitation. Plant J. 6, 673-685. Barkan, A., Walker, M., Nolasco, M., and Johnson, D. (1994). A nuclear mutation in maize blocks the processing and translation of several chloroplast mRNAs and provides evidence for the differential translation of alternative mRNA forms. E M B O J. 13, 3170-3181. Beelman, C. A., and Parker, R. (1995). Degradation of mRNA in eukaryotes. Cell (Cambridge, Mass.) 81, 179-183. Binder, R., Horowitz, J. A., Basilion, J. P., Koeller, D. M., Klausner, R. D., and Harford, J. B. (1994). Evidence that the pathway of transferrin receptor mRNA degradation involves an endonucleolytic cleavage within the 3' UTR and does not involve poly(A) tail shortening. E M B O J. 13, 1969-1980. Blank, A., and McKeon, T. A. (1989). Single-strand preferring nuclease activity in wheat leaves is increased in senescence and is negatively photoregulated. Proc. Natl. Acad. Sci. U.S.A. 86, 3169-3173. Blank, A., and McKeon, T. A. (1991a). Three RNases in senescent and nonsenescent wheat leaves. Characterization by activity staining in sodium dodecyl sulfate-polyacrylamide gels. Plant Physiol. 97, 1402-1408. Blank, A., and McKeon, T. A. (1991b). Expression of three RNase activities during natural and dark-induced senescence of wheat leaves. Plant Physiol. 97, 1409-1413. Boiler, T., and Wiemken, A. (1986). Dynamics of vacuolar compartmentation. Annu. Rev. Plant Physiol. 37, 137-164. Borochov, A., and Woodson, W. R. (1989). Physiology and biochemistry of flower petal senescence. Hortic. Rev. 11, 15-43. Bourque, J. E. (1995). Antisense strategies for genetic manipulations in plants. Plant Sci. 105, 125-149.
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6 S-RNases and Other Plant Extracellular Ribonucleases SIMON K. PARRY,* ADRIENNE
YING-HONG
E. CLARKE,
*a AND
LIU,t ED
NEWBIGIN*
*Plant Cell Biology Research Centre School of Botany University of Melbourne Parkville, Victoria 3052 Australia tDepartment of Plant Science Waite Institute University of Adelaide Adelaide, South Australia 5005 Australia
I. Introduction II. Self-lncompatibility in the Solanaceae A. Role of Ribonucleases in Arrest of Pollen Tube Growth B. Glycosylation of S-RNases III. S-RNases from the Rosaceae IV. Extracellular Ribonucleases Associated with Stress A. Phosphate Starvation B. Senescence V. Seed RNases and Other RNases of Unknown Function and Location VI. Plant Extracellular RNases: Enzymes in Search of a Substrate References
1To whom correspondence should be addressed.
RIBONUCLEASES: STRUCTURES AND FUNCTIONS
191 Copyright 9 1997by AcademicPress, Inc. All rights of reproduction in any form reserved.
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I. I N T R O D U C T I O N
Ribonucleases (RNases) occur as soluble enzymes in most plant tissues (Farkas, 1982). Although many plant RNases are involved in "classical" activities such as the processing and turnover of cellular RNA, others are located either outside the cell or within the vacuole and probably do not take part in normal R N A metabolism. In some instances, these "nonclassical" RNases were first described as effectors of certain biological processes and only later found to be RNases; in other cases, RNase activities or RNase-related sequences were identified that accumulated in response to physiological or developmental signals or as a result of external signals such as nutrient deficiency or attack by pathogens. The number of known plant extracellular and vacuolar RNases is increasing, and D N A and protein sequences from several different examples are now available. Some plant RNases have been purified as enzymatically active proteins in quantities sufficient for performing detailed studies of their structure. Our biological understanding of these RNases, however, has lagged behind the recent molecular progress, and in only one case is RNase activity known to be required for the proper function of a plant protein. In no case have substrates for these RNases been unequivocally identified. In this review, we will discuss what is known about these enzymes, their structure, and the biological functions they are thought to fulfill in the plant. We start with discussion of a class of RNases found in the flowers of some plants of the family Solanaceae, and the role of these proteins in reproductive biology.
I1. S E L F - I N C O M P A T I B I L I T Y IN T H E S O L A N A C E A E
Self-incompatibility is a widespread and genetically determined mechanism that prevents pollen from one plant from fertilizing other flowers on the same plant. Several species in the large family of plants known as the Solanaceae have a type of self-incompatibility known as gametophytic self-incompatibility. In these plants, a single genetic locus with a large number of alleles, the S-locus, controls the self-incompatibility response. When the single S-allele borne by a haploid pollen grain from a self-incompatible species lands on a style expressing the same S-allele, the growth of the pollen tube through the pistil is slowed or arrested
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(see Fig. 1). Consequently, the pollen tube is unable to deliver its sperm cells to an ovule and fertilization does not take place. Self-incompatibility thus involves an exchange of information between the pollen grain or tube and the pistil, with a differential response leading to either pollen tube growth and fertilization (a compatible response), or the arrest of pollen tube development (an incompatible response). Because the information initiating this response is encoded by the S-locus, this locus must be expressed in both pollen and pistil, with the information presumably transferred between the two cell types when the S-locus products interact. Most of the early studies on gametophytic self-incompatibility in the Solanaceae were with perennial plants with large pistils that flower
Fig. 1. Behavior of pollen in the gametophytic self-incompatibility system. Selfincompatibility is controlled by a single genetic locus (S-locus). Pollen grains are from plants with either the S~$2, SIS3, or $3S4 alleles of the S-locus. The behavior of these pollen grains on an SIS2 pistil is shown. When the S-allele of the pollen matches either of the Salleles in the pistil, growth of the pollen tube is inhibited. Pollen with S-alleles not matching the S-alleles of the pistil will successfully grow through to the ovule at the base of the pistil. For example, the $3 pollen tube can grow through an S~$2 pistil but growth of the S~ pollen tube is inhibited (middle panel). An extracellular RNase (S-RNase) determines the S-genotype of the pistil in self-incompatible plants from the Solanaceae. The gene encoding this protein is expressed only in pistils and only at later stages of floral development. The hatching shows where S-RNase genes are expressed in the mature pistil. Note that this coincides with the parts of the pistil through which the pollen tubes grow.
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throughout the year, such as the ornamental tobacco, Nicotiana alata. East and Mangelsdorf (1925) first demonstrated that self-incompatibility in N. alam is under the control of a single locus, but the products of this locus were unknown until Bredemeijer and Blaas (1981) found in the pistil different basic protein isoelectric forms (isoforms) that cosegregated with different S-alleles. Anderson et al. (1986) cloned a cDNA encoding the basic protein associated with the S2-allele of N. alata and, later, cDNAs for the $3- and S6-associated proteins of this species (Anderson et al., 1989). These proteins are all extracellular and are found only in the flower; their synthesis is developmentally regulated and correlates well with the onset of self-incompatibility (Cornish et al., 1987). Amino acid identity between the N. alata S-associated proteins is surprisingly low and only 56% of the residues are common to all three sequences. Of the conserved amino acids, nine are Cys residues and the others occur in short blocks of sequence that are interspersed between regions of more variable sequence. The most variable regions of the N. alata Sassociated proteins are dominated by hydrophilic amino acids, which suggests these regions are on the surface of the protein (Anderson et al., 1989). The S-associated proteins of N. alata are all glycosylated and the carbohydrate side chains must contribute significantly to the overall shape of the protein. There are four potential N-glycosylation sites present in the S2-associated protein and these are also present in the $3- and S6-associated proteins. Despite their sequence diversity, Anderson et al. (1989) concluded that the S-associated proteins of N. alata have similar structures composed of a framework of conserved regions held together by disulfide bonds between the conserved Cys residues, with N-glycosyl chains and variable regions exposed on the surface. Any interaction between the S-associated proteins and their pollen counterparts would presumably involve features on the surface of the protein. Subsequent to the cloning of cDNAs for the S-associated proteins from N. alata, sequence data have been published for several other selfincompatible species from the Solanaceae, including Petunia hybrida and Petunia inflata (Ai et al., 1990; Clark et al., 1990), potato (Solanum chacoense and Solanum tuberosum) (Kaufmann et al., 1991; Xu et al., 1990; Saba-EI-Leil et al., 1994), and wild tomato (Lycopersicon peruvianum) (Tsai et al., 1992; Royo et al., 1994a). Altogether, the sequences of over 30 S-associated proteins from four different genera in the family Solanaceae are known, usually from the sequence of the corresponding cDNA. Alignment of these sequences identifies five regions of highly
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conserved sequence and two regions of substantial sequence variation, with the amino acids in the conserved regions accounting for 40 out of the 202 amino acids found in an S-associated protein (Fig. 2). Two of the conserved regions (C2 and C3 in Fig. 2) are similar in sequence to the catalytic domain of an extracellular T: ribonuclease from the fungus Aspergillus oryzae (Kawata et al., 1988). In particular, both of the His residues required for catalytic activity of T2 ribonuclease are also present in the S-associated proteins (McClure et al., 1989). Indeed, the ribonuclease activity of the purified S-associated proteins from N. alata is comparable to that of T2 ribonuclease. This was later found to be true also for S-associated proteins from other self-incompatible plants such as P. hybrida (Broothaerts et al., 1991), P. inflata (Singh et al., 1991), L. peruvianum (Kowyama et al., 1994), and S. tuberosum (Kaufmann et al., 1991). Since this discovery, the solanaceous S-associated proteins are referred to as S-RNases (McClure et al., 1989). Until recently, the evidence linking S-RNases to self-incompatibility has been correlative: S-RNases are highly allelic, with each allele associated with a single allele of the S-locus. Furthermore, S-RNases are expressed in the reproductive tissues of the flower, along the path taken by pollen tubes as they grow toward the ovules and are synthesized at the same time the pistil becomes self-incompatible. Transgenic experiments have confirmed the involvement of S-RNases in self-incompatibility. Lee et al. (1994) transformed self-incompatible P. inflata plants (genotype $2S3) with a DNA construct containing the S3-RNase geneocoding region in the antisense orientation. Expression of this construct was controlled by the promoter of the S3-RNase gene. As expected, lower levels of S3RNase were found in the pistils of transformants expressing the antisense gene than were present in pistils of untransformed plants of the same S-genotype. Those transgenic plants with reduced amounts of S3-RNase were unable to reject $3 pollen; thus, S3-RNase expression was necessary for rejection of $3 pollen. Lee et al. (1994) extended these results by introducing a "sense" version of the S3-RNase coding region into P. inflata (genotype S~$2); again, expression of this construct was controlled by the S3-RNase promoter. Some of the transformed plants expressed S3-RNase in their pistil at levels similar to that found in untransformed plants bearing the S3-allele. As well as $1 and S: pollen, these transformed plants were now able to reject $3 pollen. This demonstrates that the S3RNase was sufficient for pollen rejection in these plants. Similar results to these were also obtained by expressing S-RNase constructs in a Nicotiana hybrid (Murfett et al., 1994).
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Fig. 2. Consensus of S-associated protein (S-RNase) amino acid sequences, showing those amino acids (in the one letter code) that are well conserved among 22 S-RNase sequences from four solanaceous genera. Uppercase letters indicate those residues that are almost invariant between S-RNases (the same residue is present in at least 20 of the 22 sequences), and lowercase letters indicate residues that are frequently conserved (the same residue is present in between 10 and 20 of the 22 sequences). Dashes represent positions where residues show little or no conservation (the same residue is present in fewer than 10 of the 22 sequences). The five highly conserved regions (C1-C5) and the two most variable regions (HVa and HVb) are boxed. The two putative active site His residues are shown (,) and the eight invariant Cys residues are circled.
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A. Role of Ribonucleases in Arrest of Pollen Tube Growth
The role of ribonuclease activity of S-RNases in self-incompatibility is not yet fully understood. Currently, it is thought that S-RNases act as cytotoxins by entering incompatible pollen tubes and degrading RNA, including ribosomal (r) RNA (McClure et al., 1989). The loss of rRNA and the consequent inability to synthesize protein would eventually lead to an arrest of pollen tube growth. From comparative ultrastructural analyses of compatible and incompatible pollen tubes of L. peruvianum, a model very similar to this was proposed earlier by de Nettancourt et al. (1974). These workers found that incompatible pollen tubes had major changes in the endoplasmic reticulum suggestive of a general cessation of protein synthesis. To test this model of self-incompatibility, McClure et al. (1990) made N. alata pollen radioactive by feeding plants with [32p]phosphate. The labeled pollen was used to pollinate either compatible or incompatible flowers. RNA was subsequently extracted from the pollinated pistils and analyzed by gel electrophoresis and radiography. As predicted by the model, less radioactive RNA was extracted from the pistils of flowers pollinated with incompatible pollen than from flowers pollinated with compatible pollen. Furthermore, the ribosomal RNA from incompatibly pollinated flowers was extensively degraded and that from the compatibly pollinated flowers was intact. This experiment does not unequivocally show that S-RNases are directly responsible for degradation of pollen tube RNA or even that degradation of RNA is the cause of the incompatibility response. Such RNA degradation could be an effect of the slowed growth characteristic of incompatible pollen tubes rather than its cause. If the ribonuclease activity of S-RNases is important in the selfincompatibility response, then expressing inactive forms of this protein should result in self-compatibility. Two sets of experiments--one using transgenic Petunia plants and the other using a naturally occurring selfcompatible variant of the normally self-incompatible plant, L. peruvia n u m - - h a v e shown this to be the case. As mentioned previously, SRNases contain sequences similar to those at the active site of T2 ribonuclease. In particular, these sequences contain the two His residues that are essential for catalysis of RNA by the fungal RNase. Within the S-RNases, these His residues are found at positions 34 and 95 relative to the mature N-terminal end of the protein (see Fig. 2). A version of the S3-RNase gene from Petunia was made in which a single base pair
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mutation converted the His-95 codon to an Asn codon (Huang et al., 1994). This D N A construct was introduced into Petunia and the pistils of some of the resulting transgenic plants accumulated the modified RNase at levels equivalent to those found in self-incompatible plants expressing the unmodified protein. As expected, the modified protein lacked ribonuclease activity and, consistent with the current model, these plants were unable to reject $3 pollen. The loss of ribonuclease activity is also associated with loss of selfincompatibility in wild populations of L. p e r u v i a n u m . Using fieldcollected material, two groups independently studied the genetic basis of self-compatibility in an accession of L. p e r u v i a n u m (Kowyama et al., 1994; Rivers and Bernatzky, 1994). By crossing the self-compatible accession with a closely related self-incompatible accession, a nonfunctional allele of the S-locus (Sc) was described that segregated with selfcompatibility. The pistils of progeny from these crosses were examined for ribonuclease activity and the presence of S-RNase-like proteins (Kowyama et al., 1994). Although a protein biochemically similar to SRNases was associated with the Sc allele, this protein lacked ribonuclease activity. A cDNA corresponding to the Sc protein was cloned and sequenced (Royo et al., 1994b). The sequence of the predicted protein resembles those of S-RNases except that His-34 was replaced in the sequence by an Asn residue; this could result from a single base pair change in the His-34 codon. Because the Sc protein appears normal in all other respects to functional S-RNases, it appears likely that the loss of ribonuclease activity caused a breakdown in self-incompatibility. However, the functional S-RNase gene that was the progenitor of the S~ allele is not available, although a candidate for this gene has recently been described in a self-incompatible accession of L. p e r u v i a n u m (Bernatzky et al., 1995). The available evidence points to S-RNases acting as cytotoxins within incompatible pollen tubes. However, the nature of the product of the S-locus in pollen is not known and consequently the interaction of S-locus products in pollen and pistil is not known. One possibility is that a single gene is expressed in both cell types. This implies that SRNases are the sole determinants of self-incompatibility. Indeed, low levels of S-RNases are found in developing pollen grains of several selfincompatible solanaceous species (Dodds et al., 1993; Clark and Sims, 1994; P. Dodds, C. Ferguson, and E. Newbigin, 1996, unpublished result). Other evidence points to the presence of two tightly linked genes at the
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S-locus, with one gene (now believed to be the S-RNase gene) expressed in the pistil and the other gene expressed in the pollen (Lewis, 1949). Research in many laboratories is now focused on identifying the product of the S-locus in pollen. Equally unknown is the site at which pollen and pistil products of the S-locus interact, although the current model predicts that S-RNases are specifically taken up by incompatible pollen tubes via the plasma membrane (Matton et al., 1994). An alternative hypothesis is based on nonspecific uptake of S-RNases into the pollen tube followed by specific inactivation or other modification (Thompson and Kirch, 1992).
B. Glycosylation of S-RNases The recognition event between the pollen product of the S-locus and its allelic S-RNase partner from the pistil presumably involves features on the surface of both molecules. All solanaceous S-RNases described so far are glycosylated, and carbohydrate side chains are a major surface characteristic of these molecules. There is considerable variation in both the number and structure of the side chains attached to S-RNases. For example in N. alata, the Sl-RNase has a single potential N-glycosylation site (Asn-X-Ser/Thr), whereas four sites exist in the $2-, $6o, and STRNases (Fig. 3A) (Woodward et al., 1989, 1992; Vissers et al., 1995). The S3-RNase from N. alata has the four potential sites present in the $2-, $6-, and ST-RNases plus one additional site (Woodward et al., 1989). One site is occupied in all cases, whereas the number of the other sites occupied varies with different alleles (Fig. 3A). Variation of S-RNases with respect to N-glycosylation is not restricted to the number and occupation of potential sites. Woodward et al. (1992) identified three N-linked glycan species on the Sl-RNase from N. alata. Because the S~-RNase has only one potential N-glycosylation site, there must be heterogeneity in the glycan chains at this site. Using a combination of high-field ~H NMR spectroscopy, fast atom bombardment mass spectrometry (FAB-MS), and matrix-assisted laser desorption-ionization mass spectrometry (MALDI-MS), Oxley and Bacic (1995) determined the site-specific heterogeneity and fine structure of the glycan chains at each of the four potential N-glycosylation sites of S2-RNase from N. alata (see Fig. 3A). Overall, the glycans of S2-RNase are similar to those of Sl-RNase and comprise three
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A Potential N-glycosylation site
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xylose-containing structures (glycans al, b, and c in Fig. 3B) and a small amount of Man3GlcNAc3 (type a2 in Fig. 3B). Site II of S2RNase [sites numbered according to Oxley and Bacic (1995)] contains only one type of glycan (type b in Fig. 3B) and site V is not glycosylated. The glycan chains at both sites I and IV are heterogeneous (types al, b, and c for site I, and types b and c for site IV in Fig. 3B), although the relative proportion of each structure varies between these sites. The variability in the carbohydrates attached to S-RNases and their presence on the surface of these proteins raises the possibility that the glycans are involved in recognition during self-incompatibility. More recent evidence, however, shows that this is unlikely. Karunanandaa et aL (1994) replaced the codon encoding Asn at position 29 of the petunia S3-RNase (position 30 in Fig. 2) with an Asp codon. Asn-30 is part of a potential glycosylation site (site I in Fig. 3A) present in almost all solanaceous S-RNases and is the only glycosylation site in the petunia S3RNase. The modified gene was introduced into P. inflata plants (genotype S1S2) and the presence of a nonglycosylated form of S3-RNase was detected by gel electrophoresis. Transformed plants expressing the unglycosylated S3-RNase at high levels rejected $3 pollen in the same manner seen in the S3-RNase "sense" plants described previously (Lee et al., 1994). Hence, the glycans at site I are not required for pollen rejection. A second conclusion from this experiment is that, because styles expressing the modified S3-RNase still rejected $3 pollen, modifying the protein sequence did not affect allelic specificity. Hence, S-RNases can tolerate some changes to their amino acid sequence and still retain biological activity.
Fig. 3. N-Linked glycans of the Nicotiana alata S-RNases. (A) A schematic representation of five S-RNases from N. alata. The S-RNases are all glycoproteins and the sites in the S-RNase sequence at which N-linked glycans can be attached are indicated by triangles. The number of sites in each molecule occupied by N-linked glycans is also indicated. The sites are numbered according to Oxley and Bacic (1995). (B) The structures of the Nlinked glycans from the N. alata S2-RNase. Three of the four potential sites on the S2RNase are glycosylated and more than one glycan chain is found at two of the sites (site I and site IV). See text for details. Figures reproduced and modified from Oxley and Bacic (1995) by permission of Oxford University Press.
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III. S-RNases F R O M THE R O S A C E A E
Many species in the family Rosaceae have a gametophytic selfincompatibility system similar to that found in the Solanaceae. Selfincompatible species in the Rosaceae include some that are horticulturally important, such as Malus (apples), Pyrus (pears), and Prunus (almonds and cherries). The earliest observations of self-incompatibility in this family were by Mau et al. (1982), who described a 37-kDa glycoprotein associated with self-incompatibility from the styles of cherry (Prunus avium). More recently, Sassa et aL (1992) detected several isoforms of a basic protein with RNase activity in the styles of Japanese pear (Pyrus serotina). By assaying styles from cultivars with different S-genotypes, these workers found that each RNase isoform was associated with a single S-allele. Significantly, the RNase activity in styles from a selfcompatible mutant of Japanese pear was much lower than that in the self-incompatible variety from which the mutant arose, suggesting the mutation causing self-compatibility also affects expression of the stylar RNase. Subsequent work by this group showed these RNases are similar in size and N-terminal sequence to the solanaceous S-RNases (Sassa et al., 1993), and cDNAs corresponding to these proteins are now available (Norioka et al., 1995). Results comparable to these were obtained from another rosaceous species, Malus x domestica (apple) (Sassa et al., 1994; Broothaerts et al., 1995), and cDNAs for these proteins are also available (Broothaerts et al., 1995). Comparison of the sequences of these cDNAs to the solanaceous S-RNases shows that the five short domains characteristic of the solanaceous S-RNases (C1-C5 in Fig. 2) are also found in the RNases from apple and Japanese pear (Broothaerts et al., 1995; Norioka et al., 1995), although C1, C4, and C5 are less easily recognized than C2 and C3. This finding suggests that the same self-incompatibility mechanism exists in these two distantly related taxonomic groups and, if this is the case, then the transgenic approaches used to overcome selfincompatibility in the Solanaceae could also be used to produce selfcompatible fruit tree cultivars. This would be of great commercial interest to horticulturists. However, because fruit trees take a long time to grow from seed and start flowering, the genetic analysis needed to show that the stylar RNases of apple and Japanese pear are indeed encoded by the S-locus is not straightforward. Hence, the link between these ribonucleases and self-incompatibility is not unequivocally established, although a recently described genetic map of Prunus should help to resolve this issue (Foolad et al., 1995).
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IV. E X T R A C E L L U L A R R I B O N U C L E A S E S A S S O C I A T E D WITH STRESS
A. Phosphate Starvation A major limiting factor to plant growth is the availability of inorganic phosphate. The amount of free soluble phosphate in the environment is very low and, accordingly, plants have evolved a number of mechanisms to scavenge this rare mineral nutrient. In higher plants, the existence of a pathway induced by phosphate starvation was proposed by Goldstein et al. (1989) based on studies of an acid phosphatase secreted by tomato plants and tomato cell cultures in response to phosphate deficiency (Goldstein et al., 1988a,b). Microorganisms such as Escherichia coli and yeast have well-characterized systems involving many enzymes acting together to release phosphate from organic sources and to transport the solubilized phosphate efficiently into the cell. Along with the acid phosphatase, suspension cultures of tomato cells produce an extracellular RNase termed RNase LE in response to phosphate starvation (Ntirnberger et al., 1990). When phosphate-starved cells are returned to medium containing phosphate, no further synthesis of RNase LE occurs. RNase LE is not accumulated by the cultures in response to nitrate starvation or following exposure to salt or osmotic stress, indicating that expression of RNase LE is not a general response of tomato cells to adverse growth conditions (Glund and Goldstein, 1993). RNase LE is thought to function as a phosphate-scavenging enzyme by hydrolyzing any RNA in the immediate vicinity of the plant. The sequence of the purified RNase LE protein is similar to the solanaceous S-RNases in that it contains the five regions C1-C5 and eight Cys residues present in S-RNases (Jost et al., 1991; see Fig. 2). In addition to RNase LE, cultured tomato cells express four other RNases in response to phosphate depletion. Three (RNases LV1-LV3), copurify with the vacuoles and a fourth (RNase LX) is intracellular but not in the vacuole (L6ffler et al., 1992). Interestingly, the sequences of RNase LE, RNase LX, and RNases LV1-LV3 are very similar (KOck et al., 1995). Indeed, the only difference between RNase LE and RNase LV3 is their cellular location and RNase LV1 and RNase LV2 may be derived by posttranslational processing from RNase LX. RNases similar to those from tomato are also found in A r a b i d o p s i s thaliana, a member of the mustard family (Brassicaceae). Using primers
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based on the sequences of C2 and C3 from the solanaceous S-RNases, Taylor and Green (1991) amplified D N A fragments for three RNases (RNS1-RNS3) from a cDNA library of Arabidopsis stems and leaves. RNA gel blot analyses showed that these genes are most abundantly expressed in floral tissues and that RNS2 and RNS3 transcripts are also found in several other organs. In contrast, expression of RNS1 is barely detectable in roots, inflorescence stems, and leaves (Taylor et al., 1993; Bariola et al., 1994). Members of the RNS gene family are also differentially expressed in response to phosphate starvation. RNS1 transcripts accumulated to a high level in plants grown under these conditions whereas only a slight increase was noticed in the level of RNS2 mRNA, and the level of RNS3 mRNA remained largely unchanged (Bariola et al., 1994). Arabidopsis seedlings grown without other macronutrients such as nitrogen or potassium show no comparable changes in the level of RNS mRNAs, indicating that the increased expression of RNS1 and RNS2 is specific to phosphate starvation (Bariola et al., 1994). Although the cellular localization of the Arabidopsis RNases is unknown, it is thought that RNS1 and RNS3 are secreted proteins whereas RNS2 is in the vacuole (Taylor et al., 1993).
B. Senescence
Senescence is a sequence of events occurring in tissues after their maturation. During senescence, macromolecules are degraded and important nutrients transported away from the dying tissues. Thus superficially, the biochemical events characteristic of senescence resemble those of phosphate starvation and thus it is not surprising that some hydrolytic enzymes are common to both events. Increases in RNase activity are frequently observed in senescing plant tissues (Farkas, 1982), and in Arabidopsis, levels of RNS1, RNS2, and RNS3 transcripts all increase in aging plants, albeit to different extents (Taylor et aL, 1993; Bariola et al., 1994). The effect of senescence on RNase gene expression is most apparent in the case of RNS2 transcripts, which markedly increase in abundance in aging petals and leaves (Taylor et al., 1993). The role of these Arabidopsis RNases in cellular R N A breakdown is unknown, although it is presumed to be similar to their role as phosphatescavenging enzymes (Green, 1994). Vacuoles lyse during senescence and,
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if RNS2 is in these compartments, it would be released into the cytoplasm and could degrade intracellular RNA. Blank and McKeon (1991a,b) identified three wheat leaf RNases that are induced by senescence or low levels of ambient light. All these RNases are expressed, but to different extents, by senescing flag leaves. Although their cellular location is not known, it is possible they degrade cellular RNA during leaf senescence in wheat (Blank and McKeon, 1991b).
V. S E E D RNases A N D O T H E R RNases OF U N K N O W N F U N C T I O N AND LOCATION
RNases have been purified from seeds of several species from the Cucurbitaceae (squashes and pumpkins). One of these, RNase MC from bitter gourd (Momordica charantia), has been completely sequenced and crystallized (Ide et aL, 1991; De and Funatsu, 1992) and partial protein sequences are available for two related proteins, cusativin and melonin, from cucumber (Cucumis sativis) (Ro~o et aL, 1994a) and melon (Cucumis melo) (Rojo et aL, 1994b), respectively. These proteins are similar in sequence to the solanaceous S-RNases and, in particular, contain the C2 and C3 peptides from the active site of these enzymes. Overall, RNase MC and the S-RNases are approximately 41% identical (Ide et aL, 1991). Although well studied as molecules, the biological role of the seed RNases is speculative and their cellular location is unknown. RNase activity is invariably higher in infected plants than in healthy plants although this increase may be more related to physical stresses and premature senescence than to expression of pathogen-specific RNases (Farkas, 1982; Green, 1994). For example, rust-infected wheat seedlings have considerably higher levels of RNase activity in their leaves than do uninfected plants (Rohringer et aL, 1961). This increase occurs in two phases; an early phase shortly after the initial infection, followed 3 to 5 days later by a second phase. Chakravorty et al. (1974) found that the second phase of RNase accumulation occurs only if the wheat plant is susceptible to the rust and not if the plant is resistant. The intercellular washing fluid of infected plants contains approximately six extracellular RNases, two of which are more abundant in susceptible than in resistant plants (Barna et al., 1989). These RNases have not been further characterized and their sequences are unknown.
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Changes in RNase activity are also observed during development of tomato fruit (McKeon et al., 1991). Activity of two of these RNases, Tfl and Tf2, increases to a maximum 5 days after anthesis, declines as the fruit matures, but increases again slightly during ripening. Tfl and Tf2 are glycosylated RNases with molecular masses of 59 and 29 kDa, respectively. Hence, these tomato fruit RNases are not identical to the previously mentioned tomato cell culture RNases, which are smaller (approximately 24 kDa) and not glycosylated. Tfl is composed of two subunits of 30 and 29 kDa, but only the 30-kDa subunit has RNase activity. Interestingly, the 30-kDa subunit of Tfl, and Tf2, appear to be related because an antibody raised against Tf2 only recognizes this subunit of Tfl. The biological roles of Tfl and Tf2 are unknown, although it is unlikely they act either as intracellular cytotoxins, like the S-RNases, or as general phosphate-scavenging enzymes, like the S-like RNases, because they are most abundant during early development of the fruit. A number of other RNases or RNase activities from plants have been identified. Some, such as the RNases produced by barley aleurones in response to the hormone gibberellin (Chrispeels and Varner, 1967), are probably able to degrade other nucleic acids as well and are, hence, not strictly RNases (Brown and Ho, 1986). Others, such as the pistil RNases from Petunia and Nicotiana, are very similar to the S-RNases but are not products of the S-locus and, therefore, not involved in the selfincompatibility response (Lee et al., 1992; Kuroda et al., 1994). The biological role of these extracellular proteins is unknown, although it is thought that the pistil RNases may be defense molecules that protect the pistil from pathogenic bacteria and fungi. Pistils are seldom attacked by pathogens despite being rich in nutrients such as proteins, free amino acids, lipids, and carbohydrates, and are known to produce several types of antimicrobial proteins (Atkinson et al., 1994).
VI. P L A N T E X T R A C E L L U L A R RNases: E N Z Y M E S IN S E A R C H OF A S U B S T R A T E
Extracellular RNases are widely distributed across all plant families. They are apparently involved in a number of physiological functions, although in many cases these functions have yet to be determined. Sometimes, these enzymes are cytotoxins. In other systems, cells that secrete
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cytotoxic ribonucleases also produce an intracellular inhibitor of this molecule (see Chapter 4 on a-sarcin and Chapter 2 on barnase/barstar, this volume); the inhibitor is probably required to overcome the unwanted effects of these toxic enzymes. No inhibitors of extracellular RNases have been found in plants although the search for such inhibitors has been neither systematic nor extensive, and we cannot rule out the possibility of their existence entirely. Although the enzymatic activity of plant extracellular RNases appears to be required for their biological functions, in no instance has the in vivo substrate been clearly identified. In other systems, such as the cleavage of the eukaryotic 28S ribosomal RNA by a-sarcin, the in vivo substrate is known and the site of cleavage is very specific (see Chapter 4, this volume). However, when ct-sarcin is assayed in vitro, it degrades protein-free RNA nonspecifically to low molecular weight oligoribonucleotides. Hence, the in vitro assay does not always reflect the biological activity of the enzyme. Plant extracellular RNases have a similar lack of specificity in vitro (for example, see McClure et al., 1990), 5ut this may not reflect the in vivo situation. More work on the biological substrates of plant ribonucleases is needed. Most of the extracellular RNases of plants for which sequence information is available have the short regions of conserved amino acid residues characteristic of the T2 RNases from fungi. Modifications to these sequences show, for example, that the RNase activity of the SoRNases is important for self-incompatibility. However, the biological mechanisms through which these molecules act are not known. Precisely how an extracellular ribonuclease can arrest the growth of a pollen tube or possibly contribute to fruit ripening frames important questions for future research.
ACKNOWLEDGMENTS
The authors appreciate the contributions of colleagues to this work and particularly thank Peter Dodds for help with the sequence alignments and David Oxley for comments on glycosylation. The Plant Cell Biology Research Centre is funded through a Special Research Centre grant from the Australian Research Council and SKP is the recipient of a Melbourne University faculty scholarship.
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Rohringer, R., Samborski, D. J., and Person, C. O. (1961). Ribonuclease activity in rusted wheat leaves. Can. J. Bot. 39, 775-784. Rojo, M. A., Arias, F. J., Iglesias, R., Ferreras, J. M., Mufioz, R., Escarmfs, C., Soriano, F., L6pez-Fando, J., M6ndez, E., and Girb6s, T. (1994a). Cusativin, a new cytidinespecific ribonuclease accumulated in seeds of Cucumis sativus L. Planta 194, 328-338. Rojo, M. A., Arias, F. J., Iglesias, R., Ferreras, J. M., Soriano, F., M6ndez, E., Escarmis, C., and Girb6s, T. (1994b). Enzymatic activity of melonin, a translational inhibitor present in dry seeds of Cucumis melo L. Plant Sci. 103, 127-134. Royo, J., Kowyama, Y., and Clarke, A. E. (1994a). Cloning and nucleotide sequence of two S-RNases from Lycopersicon peruvianum (L.) Mill. Plant Physiol. 105, 751-752. Royo, J., Kunz, C., Kowyama, Y., Anderson, M. A., Clarke, A. E., and Newbigin, E. (1994b). Loss of a histidine residue at the active site of S-locus ribonuclease is associated with self-compatibility in Lycopersicon peruvianum. Proc. Natl. Acad. Sci. U.S.A. 91, 6511-6514. Saba-EI-Leil, M. K., Rivard, S., Morse, D., and Cappadocia, M. (1994). The Sll and S13 self incompatibility alleles in Solanum chacoense Bitt. are remarkably similar. Plant Mol. Biol. 24, 571-583. Sassa, H., Hirano, H., and Ikehashi, H. (1992). Self-incompatibility-related RNases in styles of Japanese pear (Pyrus serotina Rehd.). Plant Cell Physiol. 33, 811-814. Sassa, H., Hirano, H., and Ikehashi, H. (1993). Identification and characterization of stylar glycoproteins associated with self-incompatibility genes of Japanese pear, Pyrus serotina Rehd. Mol. Gen. Genet. 241, 17-25. Sassa, H., Mase, N., Hirano, H., and Ikehashi, H. (1994). Identification of self-incompatibility-related glycoproteins in styles of apple (Malus • domestica). Theor. Appl. Genet. 89, 201-205. Singh, A., Ai, Y.. and Kao, T.-H. (1991). Characterization of ribonuclease activity of three S-allele-associated proteins of Petunia inflata. Plant Physiol. 96, 61-68. Taylor, C. B., and Green, P. J. (1991). Genes with homology to fungal and S-gene RNases are expressed in Arabidopsis thaliana. Plant Physiol. 96, 980-984. Taylor, C. B., Bariola, P. A., del Cardayr6, S. B., Raines, R. T., and Green, P. J. (1993). RNS2: A scnescence-associated RNase of Arabidopsis that diverged from the SRNases before speciation. Proc. Natl. Acad. Sci. U.S.A. 90, 5118-5122. Thompson, R. D., and Kirch, H.-H. (1992). The S locus of flowering plants: When selfrejection is self-interest. Trends Genet. 8, 381-387. Tsai, D.-S., Lee, H.-S., Post, L. C., Kreiling, K. M., and Kao, T.-H. (1992). Sequence of an S-protein of Lycopersicon peruvianum and comparison with other solanaceous Sproteins. Sex. Plant Reprod. 5, 256-263. Vissers, A., Dodds, P., Golz, J. F., and Clarke, A. E. (1995). Cloning and nucleotide sequence of the ST-RNase from Nicotiana alata Link and Otto. Plant Physiol. 108, 427-428. Woodward, J. R., Bacic, A., Jahnen, W., and Clarke, A. E. (1989). N-linked glycan chains on S-allele-associated glycoproteins from Nicotiana alata. Plant Cell 1, 511-514. Woodward, J. R., Craik, D., Dell, A., Khoo, K.-H., Munro, S. L. A., Clarke, A. E., and Bacic, A. (1992). Structural analysis of the N-linked glycan chains from a stylar glycoprotein associated with expression of self-incompatibility in Nicotiana alata. Glycobiology 2, 241-250. Xu, B., Mu, J., Nevins, D. L., Grun, P., and Kao, T.-H. (1990). Cloning and sequencing of cDNAs encoding two self-incompatibility associated proteins in Solanum chacoense. Mol. Gen. Genet. 224, 341-346.
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7 Evolutionary Reconstructions in the Ribonuclease Family S T E V E N A . B E N N E R , * ' t M A U R O I. C I G L I C , * M O N I K A H A U G G , * T H O M A S M. J E R M A N N , * J O C H E N G . O P I T Z , * SUN-AI RAILLARD-YOON,* JOSEF SOUCEK,~ JOSEPH .. STACKHOUSE,* N A T H A L I E T R A B E S I N G E R - R U F , * K A T R I N T R A U T W E I N , * A N D T O D D R. Z A N K E L w * Department of Chemistry ETH, CH-8092 ZUrich, Switzerland t Department of Chemistry University of Florida Gainesville, Florida 32611 $ Institute of Hematology and Blood Transfusion 128 20 Prague 2 Czech Republic wPGEC--Plant Gene Expression Center Albany, California 94710
I. Introduction II. Protein Engineering to Understand Evolution A. Ribonucleases as a System B. Questions in Molecular Evolution C. Experimental Paleomolecular Geobiology III. Reconstructing Evolution of Biomolecular Behavior in the RNase Superfamily A. Seminal RNase as a System for Studying Evolution of Behavior B. Collection of Additional Seminal RNase Sequences from Recently Diverging Artiodactyls C. Origin of Dimeric Structure in Seminal RNases D. Origin of Catalytic Activity against Duplex RNA E. Origin of Immunosuppressivity in Seminal RNase IV. Repair of Damaged Pseudogenes by Gene Conversion: A Mechanism for Obtaining Biomolecular Function in Proteins
213 RIBONUCLEASES: STRUCTURES AND FUNCTIONS
Copyright 9 1997by AcademicPress, Inc. All rightsof reproductionin any formreserved.
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V. PhysiologicalFunction of Seminal RNase VI. Conclusions References
I. INTRODUCTION Ribonucleases (RNases) of the bovine pancreatic superfamily have long played a central role in the development of ideas and technologies to study protein structure and catalysis (Richards and Wyckoff, 1971; Blackburn and Moore, 1982). In part, this is because RNase is small, present in large amounts in the pancreas of ruminants, stable under a wide range of conditions, and therefore amenable to full chemical analysis. It was among the first proteins to be sequenced, the first protein to be examined by NMR spectroscopy (Saunders et al., 1957), and the first to be unfolded and refolded in the laboratory (Anfinsen, 1973). RNase has also played an important role in the development of the protein engineering technology that emerged in the 1980s as a tool to study enzyme structure and mechanism. The first gene synthesized to encode an enzyme was for RNase (Nambiar et al., 1984). This was also the first gene designed to facilitate "cassette mutagenesis" (then called "modular mutagenesis") (Presnell and Benner, 1988). Expression of the recombinant RNase in bacterial cells followed quickly (Nambiar et al., 1987; McGeehan and Benner, 1989), and some 200 mutant forms of RNase A have been prepared in Zurich to date (Allemann et al., 1991; Trautwein-Fritz and Benner, 1991a; Trautwein-Fritz, 1991b; TrautweinFritz, 1991; Raillard, 1993; Boix et al., 1994, Jermann, 1995; Jermann et al., 1995; Opitz, 1995; Ciglic et al., 1996; Opitz et al., 1996; Haugg et al., 1995). Still more have now prepared by Miranda and Petsko (Miranda, 1990), Baldwin and co-workers (Schultz et al., 1992), Scheraga and coworkers (Laity et al., 1993; Dodge et al., 1994), Raines and co-workers (Delcardayre and Raines, 1994, 1995a,b; Raines et al., 1995; Kim et al., 1995a,b), and D'Alessio and co-workers (Di Donato et al., 1994, 1995; Cafaro et al., 1995). In part because the mechanism of RNase was so well studied using classical tools (Richards and Wyckoff, 1971; Blackburn and Moore, 1982), molecular biological methods have, at first glance, added little to
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the already elaborate picture of catalysis in RNase provided by classical enzymology. Mutagenesis studies with RNase A showed that His-12 and His-ll9 played critical roles in catalysis, whereas Lys-41 and Asp-121 played central, if less essential, roles (Trautwein and Benner, 1991a). Thr-45 was confirmed to play an important role in the binding of the nucleobase (Miranda, 1990), and Lys-7 and Arg-10 were shown to participate in the p2 phosphate binding site of RNase (Boix et al., 1994) by mutagenesis. Virtually all of the results were anticipated, however, by classical protein modification studies and crystallography (see, for example, Stern and Doscher, 1984). Folding in RNase proved to be more difficult to study by classical methods, providing an opportunity for protein engineering to make a contribution. For example, it was easy to show by mutagenesis, but not by classical studies, that introduction of single cysteines at positions 31 and 32 in the pancreatic RNase sequence yields dimeric RNases (Raillard, 1993; Jermann, 1995), whereas addition of Leu-28 and Pro-19 together with both cysteines yields a dimeric structure similar to that found in seminal RNase (Raillard, 1993; Di Donato et al., 1994, 1995). Mutagenesis also proved to be critical for defining the role of peptidylPro cis-trans isomerization in the folding of RNase (Schultz et al., 1992).
II. P R O T E I N E N G I N E E R I N G T O U N D E R S T A N D E V O L U T I O N
A. Ribonucleases as a System
Even in the early 1980s, when we began our work with this system (Nambiar et al., 1984), it was clear that molecular biological tools could provide far wider opportunities for exploring the structure, function, and evolution of proteins. In large part, the opportunity was defined by Jaap Beintema and co-workers in Groningen, who had undertaken the Herculean task of documenting the sequences of pancreatic RNases from many artiodactyls, the mammal order that includes the camel, giraffe, deer and moose, sheep and goat, the antelopes, and the ox (Beintema et al., 1986, 1988; Beintema, 1987). As early as 1979, it was possible to construct a reasonably complete evolutionary tree describing the interrelatedness of pancreatic RNases in artiodactyls, and reconstruct (at least on paper) the sequences that were the most likely intermediates
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in the evolution of this protein family (see review in Beintema et al., 1986, and chapter 8 this volume). In 1980, we set out to use these reconstructions to implement an idea, suggested two decades before by Linus Pauling and Emil Zuckerkandl (Pauling and Zuckerkandl, 1963), of preparing ancient sequences in the laboratory and studying the evolution of their behavior. As we outline in the following text, these reconstructions were to provide new insight into the structural origin of the quaternary structure in RNases, their catalytic activities, their immunosuppressivity and cytostatic activities, and ultimately their physiological function. Further, it was clear that pancreatic RNases were only part of a much larger protein family, and that their cousins were not "merely" digestive enzymes. Nearly two decades earlier, D'Alessio (1962) reported that an RNase was a major component of bovine seminal plasma, constituting some 2% of the total protein. Although the function of seminal RNase was unknown, it was clearly not a digestive protein in the same sense as pancreatic RNase (Barnard, 1969). In the early 1970s, Matousek and colleagues (Dostal and Matousek, 1973; Matousek, 1973) showed that seminal RNase blocked the growth of tumor cells. Soucek et al. (1983, 1986) showed that seminal RNase was also immunosuppressive in a mixed lymphocyte culture assay. The cytostatic activity of seminal RNase was shown to be remarkably selective, blocking the division of transformed cells with little effect on untransformed cells (Vescia et al., 1980; Vescia and Tramontano, 1981). In each case, these unexpected biological activities were largely lacking in pancreatic RNase A. These results suggested that an experimental analysis of the divergent evolution of RNases that included seminal RNase might also help us understand the nondigestive functions of members of the RNase superfamily, and perhaps the evolution of function in proteins more generally. These results also led us to speculate that the digestive family of RNases belonged to a still larger superfamily of homologous proteins performing nondigestive roles throughout higher eukaryotes (Benner, 1988). This speculation has proved to be far more accurate than we could possibly have imagined. Vallee and co-workers isolated angiogenin from tumor-conditioned cell medium while seeking substances important in the vascularization of solid tumors (Strydom et al., 1985); they showed this protein to be a member of the RNase superfamily of proteins. Similar discoveries have placed the eosinophile cationic protein and the eosinophile-derived neurotoxin, which causes neuronal degeneration (Rosenberg et al., 1989), the P-30 protein from amphibian eggs, which
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has entered clinical trials as a human antitumor agent (Ardelt et al., 1991), and a sialic acid-binding lectin from frog (Okabe et al., 1991) within the RNase superfamily. These properties as well are interesting targets for an evolutionary analysis.
B. Questions in Molecular Evolution
At this point, it is necessary to pause the narrative to introduce some general themes drawn from the field of molecular evolution. Under the Darwinian model as amplified to incorporate what we know about biomolecular structure, proteins divergently evolve from common ancestors by a process of random variation followed by natural selection. Variation that helps an organism survive, select a mate, and reproduce ("adaptive variation") is rapidly fixed in a population. Variation that confers no particular selective advantage ("neutral variation") can also be fixed in a population (Kimura, 1982; King and Jukes, 1969), although more slowly, leading to the gradual change in the structure of biomolecules ("neutral drift") at a rate that is independent of population size, expected to scale with geological time once adjusted for minor factors (such as the generation time of the organisms holding the diverging proteins). Although those advocating "adaptionism" and "neutralism" have engaged in heated discussions over the years, it is now generally recognized that both processes occur. Further, it is uniformly agreed that some features of a protein sequence are critical to function in virtually all environments, are "functionally constrained" from drifting, and therefore remain constant in virtually all proteins in a family. The explosion in the size of protein sequence data bases and powerful new tools for organizing and analyzing these data bases (Gonnet and Benner, 1991; Gonnet et al., 1992) have allowed us to gain a solid empirical understanding of how amino acids are substituted in real proteins undergoing real evolution subject to functional constraints and adaptive pressures. We and others have documented the pattern of substitutions, insertions, and deletions in proteins diverging under functional constraints based on the entire data base (Benner et al., 1993b, 1994a,b). These have provided an empirical model describing how proteins in the aggregate behave during divergent evolution. With this model in hand, we can reconstruct the history of the evolution of a protein family and reconstruct the sequences of ancestral proteins, at
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Steven A. Benner et al.
least in a probabilistic form, given the sequences of descendent proteins. Easiest to understand are reconstructions made by applying a rule of "parsimony," although maximum likelihood methods are also available for the use of the public (Benner et al., 1995). Parsimony methods reconstruct the sequences of ancient proteins in an evolutionary tree so that the descendent proteins can be accounted for by the smallest number of independent evolutionary events (Pauling and Zuckerkandl, 1963; Fitch, 1971). Parsimony reconstructions have two requirements: (1) An alignment of the homologous sequences, where the sequences of the proteins are juxtaposed so that their similarities are most obvious. In general, "obviousness" is judged by a scoring function based on the model for the evolutionary processes that lead to these sequences. (2) A tree showing the evolutionary relationship between the sequences. An example of a maximum parsimony reconstruction, done for a segment of the RNase protein, recreated the sequence of the most recent common ancestor of the RNases in swamp buffalo, river buffalo, and ox (Fig. 1). For example, the most parsimonious reconstruction at position 39 is Leu in this organism; any other reconstruction increases the number of point mutations that must be incorporated into the tree (Stackhouse et al., 1990). In its technical implementation, parsimony analysis is more problematic than Fig. 1 reveals. In reality, the alignment, the tree, and the
37 38 39 40 41 42 43 Arg Asn Met Thr Lys Asp Arg
Pachyportax
Arg Asp Met Thr Lys Asp Arg
Eland
~ .~M39L ",~
/
Arg Asn Leu Thr Lys Asp Arg Arg Asn Met Thr Ser Asp Arg Arg Ser Met Thr Ser Asp Arg
37 38 39 40 41 42 43
River
Buffalo
~
~
"
K41S
Swamp
Buffalo N38S
Fig. 1. Reconstructing a segment of the sequence of the ribonuclease in the most recent common ancestor of ox, river buffalo, and swamp buffalo by parsimony analysis.
7
Evolutionary Reconstructions
219
reconstructed sequences must be generated together, as each depends on the other. Thus, the tree itself is chosen based on a parsimony criterion, where the "best" tree is presumed to be the one that relates the contemporary sequences with the smallest number of mutations. The optimal alignment in turn can also be judged by a parsimony criterion. It is mathematically provable that the discovery of an optimal tree and/ or multiple alignment is computationally difficult ("nP complete"). As the number of sequences become large, this process becomes computationally prohibitive. Therefore, most trees are constructed in practice using one of a variety of heuristics that approximate the correct tree and then adjust the tree to obtain a local optimum that (one hopes) is not very different from the global optimum. Ancestral reconstructions are then based on this tree. The technical aspects of tree construction and the reconstruction of ancestral sequences are beyond this discussion (Fitch and Margoliash, 1967). However, it is worth making three comments concerning the practical aspects of reconstructing sequences. First, in handling real revolutionary data, the variable positions, not the conserved positions, define the connectivity of the tree. These are often few in number. For example, in the gene sequences from the seminal RNases discussed below, only four dozen positions are variable. As with any set of real data, sequence data contain "noise" that reflects the random events that define biological evolution, together with a small (one hopes) number of sequence errors. This implies that alternative trees with different connectivities will differ in their parsimony score from the "best" tree by a small amount, less than the expected statistical fluctuations in the data that define the tree. This means that the connectivity of the tree is always subject to uncertainty that is often difficult to quantitate. Next, the greatest ambiguity in reconstructing ancient sequences arises in practice from ambiguity in the connectivity of the tree. A good illustration is the reconstruction of the residue at position 38 in the pancreatic RNase family, a reconstruction that has been the target of recent discussion (Schluter, 1995; Benner et al., 1995). Figure 2 shows several alternative trees, differing only slightly, in the placement of the point of divergence of the pig, hippopotamus, and seminal RNase family. Grouping the pig and the hippo together as a suborder within the artiodactyl order, as opposed to modeling the divergence of pig and hippo as separate branches from the main lineage, changes the number of ancestors, and possibly creates ambiguities in the reconstructions. In many cases, these
Tree B
Tree A
/~ D
3 substitutions
D . / a ~
.,b~ D
c
D G
e 9
swamp buffalo river buffalo ox -~ eland nilgai
D D D
,/~ 4 substitutions
D N
impala thompson's gazelle bridled gnu topi goat
S D D D D
pronghorn antelope giraffe
G D
roe deer reindeer red deer fallow deer
D D D D
D D / a ~ D /b-...~ c / " ~~ D /d~ D / ~ ~
swamp buffalo river buffalo ox eland nilgai
impala
D D D D
pronghorn antelope giraffe
G D
roe deer reindeer red deer fallow deer
D D D D
~
D D
e
G G GD D i
bovine seminal plasma G
bovine seminal plasma G
G
50
' ' ' l'0 40 30 20 million years before present (approximate)
'0
camel, acidic camel, basic
G G
hippopotamus pig
D G
S
thompson'sgazelle bridled gnu topi goat
horse G
' 50
40 ' ' 30 20 10 million years before present (approximate)
camel, acidic camel, basic
G G
hippopotamus pig
D G
0'
Fig. 2. P a r s i m o n i o u s r e c o n s t r u c t i o n s of residue 38 based on a l t e r n a t i v e tree connectivities. T h e filled circles (O) indicate points on the tree w h e r e the a n c i e n t r e c o n s t r u c t i o n s r e q u i r e an a m i n o acid substitution. N o t e that Gly is the p r e f e r r e d ( m o s t p a r s i m o n i o u s ) r e c o n s t r u c t i o n regardless of h o w pig a n d h i p p o p o t a m u s are placed on the tree.
Tree D
Tree C
/,~ 3 substitutions
D /b
D
~
D c
D G G /
e ~
G G j/ G
swamp buffalo river buffalo ox eland nilgai impala thompson's gazelle bridled gnu topi goat
D D D
pronghorn antelope giraffe
G D
moose
D
roe deer reindeer red deer fallow deer
D D D D
/~ 4 substitutions
c
D
g
/~D/ G DJ
swamp buffalo river buffalo ox eland nilgai impala thompson's gazelle bridled gnu topi goat
D D
pronghorn antelope giraffe
G D
moose roe deer reindeer red deer fallow deer
D D D D D
G
D
D N S D D D D
bovine seminal plasma G
camel, acidic camel, basic
G G
camel, acidic camel, basic
G G
hippopotamus pig
D G
hippopotamus pig
D G
f'
30 20 0 million years before present (approximate)
D / a ~ D /b-..~
D N S D D D D
bovine seminal plasma G
50
D
0
50
410
I
30
210
1'0
million years before present (approximate)
i
0
222
Steven A. Benner et
al.
ambiguities are inconsequential. In this example, at position 38, the Gly in the horse and whale make it virtually certain that a Gly will appear in position 38 in some ancestral sequence, regardless of the specifics of the tree. When ambiguity remains, alternative sequences can be reconstructed and studied independently (Jermann et al., 1995). Nevertheless, the point should be made that these reconstructions remain hypotheses, to be explored experimentally, not unambiguous statements about past events. Finally, both the tree and the multiple alignment are intended to be models for a biological reality. Artiodactyls actually did diverge from common ancestors in a specific order. A "correct" tree is not, therefore, defined as the tree that most parsimoniously interrelates a set of protein sequences, but rather as the tree that correctly represents this biological reality. A variety of independent information is available to infer this reality, including a cladistic analysis of the physiology of the organisms, the fossil record, and the analysis of the sequences of other protein families. These factors are appropriately used in the selection of a preferred tree, and a less parsimonious tree may be accepted as a better representation of biological reality than a more parsimonious tree for these reasons.
C. Experimental Paleomolecular Geobiology
The RNase family of proteins has also played the key role in the establishment of a new field involving molecular reconstructions, needing a name, but currently referred to as "experimental paleomolecular geobiology." In its broadest sense, molecular geobiology is concerned with unraveling and interconnecting the biological, chemical, and geological events that have shaped the earth. It draws on information contained within both the biosphere and geosphere at the atomic and molecular level, to understand better the evolution of the earth and the life that it carries. The experimental fusion of the earth and life sciences has allowed ancient proteins to be reconstructed in the laboratory and studied. This was done first independently in the laboratories of Wilson and coworkers at Berkeley with the digestive protein lysozyme (Malcolm et al., 1990), and in our laboratories in Zurich with RNase (Stackhouse et al., 1990). In the case of RNase, the sequence for the ancestral protein reconstructed for the most recent common ancestor of swamp buffalo, river buffalo, and ox (Fig. 1) approximates (but does not correspond
7
223
Evolutionary Reconstructions
exactly to) the fossil organism P a c h y p o r t a x , known in the fossil record from the Indian subcontinent. Having an ancient protein in the laboratory allows us to learn w h e t h e r parsimony analysis has yielded plausible protein sequences for these evolutionary intermediates in the R N a s e family, and possibly something about their physiological function. M o d e r n digestive RNases are k n o w n to be catalytically active against small R N A substrates and singlestranded R N A (Blackburn and Moore, 1982), and stable to proteolytic digestion (Lang and Schmid, 1986). These are also properties of the reconstructed P a c h y p o r t a x RNase. Its kcat/Km values with U p A as substrate do not differ substantially from those of contemporary bovine digestive RNase (Table I) (Ipata and Felicioli, 1969), nor does it catalytic activity with poly(U) as substrate (Table II). This suggests both that the reconstructed sequence is plausible and that the reconstructed protein was a digestive enzyme in the extinct ancestor, just as it is in its descendants (Barnard, 1969). Sequence data from Beintema (1987), supplemented by sequence data from Breukelman et al. (1993) and from our laboratories in Zurich (see later), allow us to go still further back in time. RNases from the more ancient ancestors, reconstructed at the branch points leading to various antelopes, the goat, the giraffe, and the deer, also behave as digestive enzymes by these criteria. They are stable to trypsin, act on small R N A substrates, and have low catalytic activity against double-stranded R N A (Table III). Similar behavior is not seen, however, for all reconstructed ancient RNases. For example, in the ancestors of the ox and the camel, and ox and
TABLE I Kinetics of RNase from
"Pachyportax" ( U p A ) ~
RNase source
kcat (sec -l)
gm (mM)
Bos taurus
1459 + 2055 + 1906 + 2003 + 1602 + 1944 +
211 250 229 208 169 203
A19S A19S L35M L35M Pachyportax h
50 495 301 107 102 136
+ + + + + +
20 41 38 18 18 27
a From Stackhouse et al. (1990). b Reconstructions of ancestral sequences using parsimony rarely correspond to particular fossil species. The fossil species Pachyportax is speculatively close to the ancestor of the ox and the buffalo, and is listed here to indicate this.
Steven A. Benner et aL
224 T A B L E II Kinetics of RNases with U p A as Substrate a
Enzyme source Bos taurus Pachyportax Archaeomeryx Diacodexis
Km (mM)
kcat (sec-1)
165 190 420 177
1900 1944 468 253
kcat/Km
11.5 x 10.2 • 1.1 x 2.2 x
106 106 106 106
a The organisms listed from the fossil record correspond only approximately to the ancestral reconstructions obtained by parsimony. From Jermann et al. (1995).
pig/hippopotamus corresponding approximately to the fossil organism the behavior changes. The stability of the protein decreases, as measured by the assay of Lang and Schmid (1986). The ability of the reconstructed RNase to catalyze the hydrolysis of the duplex R N A p o l y ( A ) - poly(U) increases approximately fivefold (Table III). These are properties not obviously consistent with a digestive function. Therefore, it appears as if digestive behavior, at least in this branch of the RNase superfamily, arose ---40 million years before present. Is this plausible? To answer this question, we must turn to the geological sciences. The geological record shows that this period of time was interesting in its climate, which had substantial impact on the biosphere, which in turn influenced the food source for artiodactyls, which correlates, in turn, with changes in their digestive system. A r c h a e o m e r y x is
Archaeomeryx,
T A B L E !11 Kinetics of RNases with P o l y ( A ) - P o l y ( U )
Source~ Bos taurus "Pachyportax" "Archaeomeryx'" "Diacodexis"
Activity 1.0
1.2 5.2 4.6
"The reconstructed ancestral sequence does not normally correspond to any particular fossil species. Fossil species indicated here are only speculative guides.
7 Evolutionary Reconstructions
225
(approximately) the first ruminant artiodactyl, and lived ---40 million years ago, just at the time that the behavior of the reconstructed sequences undergoes the changes noted above. This hypothesis is consistent with a physiological hypothesis advanced some years ago by Barnard (1969). Ruminants feed grass to bacteria in a first stomach, and then eat fresh bacteria. Approximately 20% of the nitrogen intake of the ruminant comes as RNA. Digestive RNases are needed to break down this RNA. Accordingly, Barnard noted, ruminants have large amounts of RNase in the digestive tract. So do other herbivores with ruminant-like digestion. Nonruminants do not. For example, humans have ---0.1% of the catalytic activity of digestive RNase as seen in the ox, and this comes from an RNase that is distributed throughout the organism. The digestive RNase in ox is expressed nearly exclusively in the pancreas. Ruminant physiology appears to have substantial adaptive value in many herbivorous environments; it may have convergently evolved in marsupial kangaroos, the colobine monkey primates, and more than once within the artiodactyl lineage (Joll6s et al., 1989). Ruminants ferment cellulose with increased efficiency, and ruminant artiodactyls have been enormously successful in competition with the herbivorous perissodactyls (for example, horses, tapirs, and rhinoceroses, which maintain fermentation in digestion in the cecum following the small intestine. Some 170 species of ruminant herbivores have emerged as descendants of a single founder artiodactyl. These have displaced some 250 species of nonruminant herbivores (perissodactyls) from the Miocene; of these 250 species groups, 247 (99%) have become extinct. Only three species groups of perissodactyls have survived (horses, rhinoceroses, and tapirs). Simply in terms of percentages, this mass extinction in this mammal order surpasses the one that occurred at the Permian-Triassic boundary, and approaches the one that befell the dinosaurs at the Cretaceous-Tertiary boundary (where we assume that the dinosaur orders became 100% extinct).
III. R E C O N S T R U C T I N G E V O L U T I O N O F B I O M O L E C U L A R B E H A V I O R IN T H E RNase S U P E R F A M I L Y
A. Seminal RNase as a System for Studying Evolution of Behavior
This example shows how the paleomolecular biochemist can learn about the origin of new physiological function through changes in protein
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Steven A. Benner et al.
sequence and behavior using reconstructed sequences. Given the power of contemporary protein engineering, however, it is no longer sufficient simply to state that a behavior arose in the divergent evolution of a protein family. We must also ask how, at a molecular level, the behavior arose. From the work of Matousek, D'Alessio, and colleagues, it was clear already in 1980 that the RNase family was going to provide a rich system for studying the evolution of many physical, catalytic, and biological properties, and that these studies might be generalizable. The existence of RNase from bovine seminal plasma, a close homolog of pancreatic RNase, differing by 23 amino acid substitutions, with many interesting biological properties (Benner, 1988; Benner and Allemann, 1989; D'Alessio et al., 1991), offered us this opportunity. Simple sequence analysis suggests that seminal RNase diverged from the pancreatic lineage shortly after the origin of ruminant digestion (see later). Some 2% of the protein in bovine seminal plasma is this RNase variant. It has an unknown function, but many interesting properties. First, bovine seminal RNase has 25-fold enhanced catalytic activity against duplex RNA (under approximately physiological assay conditions) when compared to its pancreatic cousin. As noted previously, increased catalytic activity against duplex RNA is a primitive (in the evolutionary sense of this term) trait in the artiodactyl RNase lineage, being present in the common ancestor of bovine seminal and bovine pancreatic RNase. Nevertheless, seminal RNase has still higher catalytic activity against duplex RNA, elevated a further 5-fold over this activity in the ancestor. Seminal RNase has several remarkable molecular properties. It is a dimer, unlike the digestive RNase from ox, which is a monomer. It is joined by two intersubunit disulfide bonds, connecting Cys-31 on one subunit with Cys-32 on the other. In the dimeric structure, residues 1-20 from one subunit (the "S-peptide") swap with residues 1-20 from the other. The S-peptide swap creates a composite active site where His-12 comes from one subunit and His-119 comes from the other. Each of these properties is also not present in the ancestral protein, which is a monomer and shows little propensity to swap S-peptides. Thus, the dimeric structure is said to be "derived" in the seminal RNase lineage. As noted above, seminal RNase also has remarkable biological properties. The protein is immunosuppressive (in, for example, a mixed lymphocyte culture assay) and cytostatic. Further, seminal RNase has evolved a new binding affinity. It binds to seminolipid, a 1-alkyl-2-
7
Evolutionary Reconstructions
227
acylglycerogalactosyl-3-sulfate (Ishizuka et al., 1973; Lingwood et al.; 1981). Seminolipid is fusogenic, and its timely migration within the spermatozoa is believed to play an important role in the maturation of the spermatozoa (Vos et al., 1994). These properties are also derived. The ancestral protein neither binds seminolipid nor displays these biological activities. Bovine seminal RNase therefore offers the experimental paleomolecular geobiologist a remarkable opportunity to study recruitment in protein evolution, wherein structure, behavior, and (presumably) function have all undergone change. Further, the change has been quite rapid. Since the gene duplication, between 12 and 14 amino acids (depending on the precise ancestral reconstruction) have been replaced to yield modern bovine seminal RNase. This represents some 10% of the protein overall. Considering that the duplication occurred only ~-35 million years before present, this divergence was extremely rapid.
B. Collection of Additional Seminal RNase Sequences from Recently Diverging Artiodactyls The literature existing when this work began contained only a single seminal RNase sequence. This was, of course, insufficient to permit reliable reconstructions within the seminal RNase branch of the RNase family of proteins. Therefore, in a collaboration with Dr. Barbara Durrant at the Center for Reproduction of Endangered Species (San Diego Zoological Society) and the Zurich zoo, biological samples were recovered for a variety of artiodactyls, including for peccary (Tayassu pecari), Eld's deer (Cervus eldi), domestic sheep (Ovis aries), oryx (Oryx leucoryx), saiga (Saiga tatarica), yellow-backed duiker (Cephalophus sylvicultor), lesser kudu (Tragelaphus imberbis), Cape buffalo (Syncerus caffer caller), forest buffalo, sitatunga (Tragelaphus spekei), nyala (Tragelaphus angasi), eland (Tragelaphus oryx), Maxwell's duiker (Cephalophus monticola maxwelli), suni (Neotragus moschatus), sable antelope (Hippotragus niger), and impala (Aepyceros melampus). The genes for pancreatic and seminal RNases were isolated and sequenced from many of these; several new sequences are presented in Fig. 3. These supplement the sequences for giraffe RNases reported by Breukelman et al. (1993), which are also given in Fig. 3.
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15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 S S T S A A S S S N Y C N Q M M K S R N L T K D R C K P V N T g V H TCCAGCACTTCCGcTGC•AGCAC•cTCCAA•TACTGTAAC•AGATGATGAAGAGC•GGAACCTGAC•AAAGATCGATGCAAC•cCAGTGAA•A••TTTGTGCA•
ox pancreatic
A S T S N Y C N Q M M K S R N L T Q N R C K P V N T F V GCCAGCACGTCCAACTACTGCAACCAGATGATGAAGAGCCGGAACCTGACCCAAAATCGATGCAAGCCAGTGAACACCTTTGTC•cAC
H
S S T S S V S S S N Y C N E M M T S R N L T Q D R C K P V N T F V TCCAGCACCTCCTCCGTCAGCAGCTCCAACTACTGCAACGAGATGATGACAAGCCGGAATCTGACCCAAGATCGATGCAAGCCGGTGAACACCTTTGTGCAT
H
pancreatic
S G S S P S TCTGGCAGCTCCCCCAGCA
N L M M F C R K M T Q G K C K P V N T F G ................ CAACCTGATGATGTTCTGCCGGAAGATGACCCAGGGGAAATGCAAGCCAGTGAACACTTTTGGTCAC
H
S G S S P S S N S N Y C N L M M F C Q K L T E G K G K P V N T F V TCTGGCAGCTCCCCCAGTAGCAACTCCAACTACTGCAACCTGATGATGTTCTGCCAGAAGTTGACCGAGGGGAAAGGCAAGCCAGTGAACACCTTTGTGCAT
H
G
S
S
P
S
S
N
S
N
Y
C
N
L
M
M
F
C
R
K
M
T
Q
G
K
C
K
P
V
N
T
F
V
H
S
G
N
S
P
S
S
S
S
N
Y
C
N
L
M
M
C
C
R
K
M
T
Q
G
K
C
K
P
V
N
T
F
V
H
49 E
50 51 52 53 54 55 56 57 58 S L A D V Q A V C
59 60 61 62 63 64 65 S Q K N V A C
66 67 68 69 70 71 72 K N G Q T N C
73 74 Y Q
75 76 77 78 79 80 S Y S T M S
81 82 I T
GAGTCCCTC,GCTGATGT••AGGC•GTGTGCTCC•AGAAAAATGTTGCCTGCAAGAATGGGCAGACCAATTGCTACCAGAGCTACT•CA•CATGAGCATCACCGAC E S L A D V Q A V C S Q K N V A C K N G Q T N C Y Q S Y S T M S I T GAGTCCCTGGCCGATGTCCAGGCCGTGTGCTCCCAGAAAAATGTTGCCTGCAAGAATGGGCAAACCAATTGCTACCAGAGCTACTCCACCATGAGCATCACAGAC E
S
L
A
D
V
Q
A
V
C
S
Q
K
N
V
A
C
K
N
G
Q
T
N
C
Y
Q
S
N
S
A
M
S
I
giraffe seminal saiga seminal duiker seminal kudu seminal buffalo seminal
H
S
TCTGGCAGCTc••CCAGCAGCAACT•CAACTACTGCAACCTGATGATGTTCTGCCGGAAGATGACCCAGGGGAAATGCAAGCCAGTGAACACCTTTGTG•AT TCTGGCAACTCCCCCAC•CAGCAGCTCCAACTACTGCAACCTGATGATGTGCTGCCGGAAGATGACCCAGGGGAAATGCAAGCCAGTGAACACCTTTGTGCAT
giraffe
H
S G S S P S S N S N Y C N V M M F C R K M T Q G K C K P V N T F A TCTGGCAGCTCC•CCAGCAGCAACTCCAACTACTGCAACGTGATGATGTTCTGCCGGAAGATGACTCAGGGGAAATGCAAGCCAGTGAACACCTTTGCGCAT
S G S S P S S N S N Y C N L M M F C R K M T Q G K C K L V N T F V TCTGGCAGCTCCCCCAGCAGCAACTCCAACTACTGCAACCTGATGATGTTCTGCCGGAAGATGACTCAGGGGAAATGCAAGCTGGTGAACACCTTTGTGCAT
oryx pancreatic
ox
seminal
83 D D
T
D
E S L A N V Q A V C S Q K K V I C K N G L S N C Y Q S N S A I H Y T GAGTCCCTGGCCAATGTCCAGGCTGTGTGCTCCCAGAAGAAAGTCATCTGCAAGAATGGGCTGTCCAACTGCTACCAGAGCAACTCCGCCATTCATTATACAGAT
D
GAGT~CCTGGCTGATGT~AGGCCGTGTGCTC~CAGAAAAATGTTG~TC~CAAGAAT~CAGACTAACTG~TA~cAGAGCAA~T~TGC~ATGAG~ATCACAGA~
E F L A D V Q A V C S Q K K V T C K N G Q T N C Y Q S N S A M S I T GAGTTCCTGGCTGATGTCCAGGCTGTGTGCTCCCAGAAGAAAGTCACCTGCAAGAATGGGCAGACCAACTGCTACCAGAGCAACTCCGCCATGAGCATCACAGAC
D
E S L A D V K A V C S R K K V T C K N G Q T N C Y Q S N S A M R I T GAGTCCCTC~7~GATGTTAAGGCTGTGTGTTCCCGGAAGAAAGTCACTTC,CAAGAATGGC,CAGACCAACTGCTA~CAGAGCAA~TCCGCCATG~GCATCACAAAG
K
E
D
E S L A D V K A V C S Q K K V A C K N G Q T N C Y Q S N S A M R I T GAGTCCCTGC•CCGATGTCAAGGCCGTGTGCTCCCAGAAGAAAGTCGCCTGCAAGAATGGACAGACCAACTGCTACCAGAGCAACTCCGCCATGCGCATCACAGAC
E S L A D GAGTCGCTGGCCGATG S
L
A
D
V
DEL C S Q K K V T C K N G Q T N C Y Q S K S T M R I T ........... TGCTCCCAGAAGAAAGTCACTTGCAAGAATGGGCAGACCAACTGCTACCAGAGCAAATCCACCATGCGCATCACAGAC K
A
V
C
S
Q
K
K
V
T
C
K
N
G
Q
T
N
C
Y
Q
S
K
S
T
M
R
I
T
GAGTC•CTGGCCGATGTTAAGG••GTGTGCTCC•AGAAGAAAGTTA•TTGCAAGAATGGGCAGACCAA•TGCTA••AGAC•CAAAT••A•CATG•G•ATCACAGA•
D
D
ox
pancreatc oryx pancreatic
giraffe
pancreatic
giraffe seminal saiga seminal duiker seminal kudu seminal buffalo seminal ox
seminal
84 85 86 87 88 89 90 91 92 93 94 95 96 97 98 99 00 01 02 03 04 05 06 07 08 09 i0 ii 12 13 14 15 16 17 18 C R E T G S S K Y P N C A Y K T T Q A E K H I I V A C E G N P Y V P V TGCCGTGAGA•CGG•AG•T•CAAGTACCC•AACTGTGCCTACAAGA••ACC•AGGCAGAGAAACA•AT•ATTGTGGCTTGTGAGGGAAACCCGTA•GTGCCAGTC
ox pancreatic
C R E T G S S K Y P N TGCCGTGAGACCGGCAGCTCCAAGTACCCCAAC
oryx pancreatic
C R E T G N S K Y P N C A Y Q T T Q A E K H I I V A C E G N P Y V P TGCCGCGAGACCC~GCAA~TCCAAGTACCCCAACTGTGCCTACCAGACCACCCAGGCAGAGAAACACATCATTGTGGCTTGCGAGGGAAACCCATACGTGCCAGTT
V
giraffe
pancreatic
giraffe seminal saiga seminal C R Q T G S S K Y P N C T C K T T R A E K H I I V A C E G K ? F M P duiker TC~C~GC~AGACTC,C~AGCTCCAAGTAC~C~AACTGCA~CTC'CAAGACCACCCGGG~GGAGAAACACAT~ATAGTGG~TTGTGAGGGA~AAC~GTTCATGCCA seminal C R E T G S S K Y P N C A Y K T T Q V E K R I I V A C E G K DEL V kudu '~CCGCGAGACTC.~`'CAC~CTCCAAGTACCCC/~.CTG~-'CCTACAAGACCACCCAC.~GTGGAGAAACGCATCATAGTC.~`~aC'`I~rGTGAGGGT~%.A-CGCACATC'CGC~TC seminal C R E T G S S K Y P N C A Y K T T H V E K R I I V A C A G K P Y V P X buffalo TGCCGCGAGACTGGCAGCTCCAAGTACCCCAACTC•CGCCTACAAGACCACCCAAGTGGAGAAACGCATCATAGTGGCTTGTGCAGGTAAACCGTACGTGCCAGTX seminal C
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TGC•GCAAGACTGGCAGCTCAAA•TACCCCAACTGTGC•TACAAGACCACCCGGGCAGAGAAACGCATCATAGTGGCTTGTGAGGGAAACCTGTAGGTGCCAGTC
C R Q T G S S K Y P N C A Y K T T Q A Q K H I I V A C E G N P Y V P TGCCGC•AGACTGC•CAGCTCCAAGTACCCCAACTGCGCCTACAAGACCACC•AGG•GCAAAAACACATCATAGTGGCq-fGTGAGGGAAACCCGTATGTGCCAGTC
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Fig. 3. Sequences of the genes and the encoded polypeptide chain (using the oneletter code) of members of the pancreatic and seminal RNase families. A single base in codon 114 is deleted in the gene from lesser kudu, creating a frame shift covering residues 114-117. Published RNase sequences from giraffe (Breukelman et al., 1993), ox pancreas (Carsana et al., 1988), and ox seminal plasma (Preuss et al., 1992) are shown.
C. Origin of Dimeric Structure in Seminal RNase With these tools in hand, we were prepared to learn how each of the unique catalytic, structural, and biological properties in seminal RNase came into being, what amino acid changes were responsible for them, and when in geological history they arose. We first addressed the origin of dimeric structure in seminal RNase (Raillard, 1993; Jermann, 1995;
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Opitz, 1995; Ciglic et al., 1995). The problem has attracted the research interest of other laboratories as well (see, for example, the elegant work of Di Donato et al., 1994, 1995), with generally congruent conclusions. Evolutionary reconstructions suggest that Cys-32 was introduced early in divergent evolution, whereas Cys-31 was introduced much later, after the divergence of kudu and ox, less than 10 million years ago (Trabesinger-Ruef et aL, 1996). Jermann (1995) showed that introduction of single cysteines at position 32 in the pancreatic RNase yields a dimeric RNase covalently joined by a disulfide bond. This disulfide bond is, of course, not the natural one found in seminal RNase (where a Cys must be present at position 31 to make the 31-32' intersubunit disulfide bond). Accordingly, this dimer does not swap its S-peptide chains (Ciglic et al., 1995). Nevertheless, these studies show that every RNase gene in the seminal lineage could, if expressed as a protein, form a covalent dimer. A single Cys at position 31 also permits dimer formation, again without S-peptide swap (Jermann, 1996; Ciglic et al., 1996). Not surprisingly, introduction of two Cys residues, at positions 31 and 32, also allows dimer formation (Raillard, 1993), whereas addition of Leu-28 and Pro19 leads to a dimeric structure with S-peptide swap similar to that found in seminal RNase (Raillard, 1993; Di Donato et al., 1994, 1995). Thus, the determinants of quaternary structure, including the S-peptide swap, can be found at positions 19, 28, 31, and 32. Interestingly, however, the natural intersubunit disulfide connectivity can be formed by interactions elsewhere in the protein (Ciglic et al., 1995). For example, substitutions at positions 31 and 32 alone do not seem to generate the natural intersubunit disulfide connectivity. Adding to these substitutions at positions 38 and 111 does, however (see below).
D. Origin of Catalytic Activity against Duplex RNA Bovine seminal RNase catalyzes the hydrolysis of double-stranded RNA significantly better under physiological conditions than does its pancreatic homolog RNase A. The evolutionary reconstructions (see previous discussion) showed that at least part of the catalytic activity against duplex nucleic acids is primitive. Nevertheless, a further fivefold increase in catalytic activity gained after the divergence of seminal RNase must still be accounted for. Here, the results were surprising. Simply creating a dimeric RNase, for example, by introducing a single Cys
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residue at position 31 or 32, or by introducing Cys residues at both positions 31 and 32, increased catalytic activity against duplex R N A by only a factor of 1 (that is, no increase at all) to 3. To gain a full increase against double-stranded nucleic acid, two glycine residues found in seminal RNase at positions 38 and 111 were required. As noted previously, these substitutions guaranteed that the two subunits were joined predominantly by the natural disulfide bonds, between Cys-31 on one subunit and Cys-32 on the other (Ciglic et al., 1995). The formation of the natural intersubunit disulfide connectivity is believed to correlate well with the ability of the RNase to hydrolyze duplex RNA. Whereas Gly-38 was present in the ancestral RNase, and Cys-32 arose shortly after the gene duplication event that created the seminal RNase lineage, codons encoding Cys-31 and G l y - l l l are very late additions to the seminal RNase gene family, arising after the divergence of kudu and ox (Fig. 3; see later). This implies that a seminal RNase with full catalytic activity against duplex RNA could have been found only very recently. Another benefit of these evolutionary reconstructions was the discovery of a close correlation between the ability of an ancestral RNase to hydrolyze duplex RNA and its ability to bind and melt duplex DNA (Opitz, 1995; Opitz et al., 1996). This is consistent with what is known about the physical organic chemistry of phosphate ester hydrolysis. In the RNase A mechanism, the 2'-hydroxyl group attacks the 3'-phosphate in an "in-line" mechanism (Westheimer, 1992). In this mechanism, the 5'oxygen of the following nucleoside must lie 180~ away from the attacking oxygen (Campbell and Petsko, 1987). In a standard A-type double helix adopted by RNA, this geometry is not possible. The 2'-hydroxyl group is not within bonding distance of the 3'-phosphate, and the 5'-oxygen of the following nucleoside is not placed 180~ away from the 2'-hydroxyl group. Thus, local melting of the helix of duplex RNA is generally presumed to precede hydrolysis, and these studies strengthen this presumption.
E. Origin of Immunosuppressivity in Seminal RNase These results permitted us to turn to examine the evolutionary origin of the unusual biological activities (Benner, 1988; Benner and Allemann, 1989; D'Alessio et al., 1991) displayed by seminal RNase. By far the most obvious structural feature evident from the crystal structure of
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seminal RNase is the dimer, joined by two intersubunit disulfide bonds, with the S-peptide units (residues 1-20) from one subunit exchanged between the two subunits. Accordingly, both we (Trautwein-Fritz, 1991; Raillard, 1993; Jermann, 1995; Opitz, 1995) and others (Kim et al., 1995a,b; Di Donato et al., 1995) have focused on dimeric structure and the S-peptide swap has key elements in determining the biological activities of seminal RNase (including immunosuppressivity and cytostatic activity). This focus has not been entirely incorrect. In collaboration with Josef Soucek and colleagues in Prague, we showed some time ago that immunosuppressivity and cytostatic activity correlated with dimeric structure, created by the introduction of Cys residues at positions 31 and 32 (Raillard, 1993). These results have been confirmed and amplified both by Raines and co-workers (Kim et al., 1995a,b) and by D'Alessio and coworkers (Di Donato et aL, 1995). Careful studies of the evolution of the seminal RNase family have shown that this model is far from complete, however. Although introducing amino acids from seminal RNase into an RNase A background to create the RNase A variant (Q28L K31C $32C) confers both dimeric structure and substantial immunosuppressivity (Raillard, 1993), equal immunosuppressivity can be obtained in the variant (Q55K N62K A64T E l l 1G Nll3K), which is not a dimer and displays no S-peptide swap (Haugg et al., 1996). Combining the two sets of mutants to yield the variant of RNase A (Q28L K31C $32C Q55K N62K A64T E l l l G Nll3K) yields a protein that is as immunosuppressive as seminal RNase. In contrast, introducing residues 76 and 80 from seminal RNase into an RNase A background actually diminishes the already weak immunosuppressivity of RNase A in a mixed lymphocyte culture assay. The implications of these results for the mechanism for the biological activity are beyond the scope of this review. However, the mechanism is not as simple as we (and others) had originally hoped. Of the eight determinants of immunosuppressivity (Q28L, K31C, $32C, Q55K, N62K, A64T, E l l l G , and Nll3K), none is found in the most recent common ancestor of seminal and pancreatic RNase. Cys32 and Lys-62 were introduced early in the divergent evolution of the seminal RNase gene (Fig. 3). Leu-28 (first seen in sheep), Lys-55 (first seen in sheep), Thr-64 (first seen in saiga), and Lys-ll3 (first seen in duiker) arose during the divergence of the seminal RNase family. However, Cys-31 and G l y - l l l were placed in seminal RNase only after the divergence of kudu and bovine seminal RNase. As we will discuss in
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Section IV, the time of this divergence is important in our understanding of seminal RNase and its evolution.
IV. R E P A I R OF D A M A G E D P S E U D O G E N E S BY G E N E C O N V E R S I O N : A M E C H A N I S M FOR O B T A I N I N G B I O M O L E C U L A R F U N C T I O N IN P R O T E I N S
The discussion to this point has used evolutionary reconstructions to develop an understanding of the sequence determinants for the various catalytic, physical, and biological features that are found in contemporary bovine seminal RNase. Dimeric structure is minimally determined by a Cys at position 31 or 32, the S-peptide swap is controlled by additional residues at positions 28 and 19, and the correct disulfide connectivity requires both cysteines and at least one additional determinant elsewhere in the sequence (Ciglic et al., 1996). Catalytic activity against duplex RNA, together with the ability to bind and melt duplex DNA, is determined by substitutions at positions 31,32, 38, and 111 (Opitz, 1995; Opitz et al., 1996). Immunosuppressivity, as measured by a mixed lymphocyte culture assay, is determined by substitutions at positions 28, 31, 32, 55, 62, 64, 111, and 113 (Haugg et al., 1996). These results, of course, say only what might have happened in the evolution of RNase to yield these properties. We have not, however, made any statement about what actually happened in artiodactyls over the past 40 million years. In the simplest model, one might view the gradual accumulation of point mutations in the seminal RNase lineage to have gradually enhanced one or more of the features of the protein that are important for survival and reproduction in the host organism. This, a classical Darwinian model, would propose that these properties would then have been gradually selected for over time. The reconstructive work based on the newly determined genes for seminal RNase (Fig. 3) (Breukelman et al., 1993) has shown that this was not, in fact, what happened. Seminal RNase genes are distinguished from their pancreatic cousins by several "marker" substitutions introduced early after the gene duplication, including Pro-19, Cys-32, and Lys-62. By this standard, as well as by parsimony analysis, the genes from saiga, sheep, duiker, kudu, and the buffaloes (Fig. 3) all are members of the seminal RNase family. No evidence for a seminal-like gene could be found in peccary (T. R. Zankel,
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1991, unpublished). Thus, these data are consistent with a parsimony analysis of previously published genes that places the gene duplication separating pancreatic and seminal RNases ~35 million years before present (Beintema et al., 1986), with the divergence of giraffe preceding the divergence of sheep, saiga, duiker, kudu, Cape buffalo, and ox, in this order, consistent with the ordering suggested by mitochondrial sequence data (Allard et al., 1992). However, many of the seminal RNase genes could not have encoded an active protein. Particularly remarkable in this respect is the seminal RNase gene from kudu. A single base deletion disrupts codon 114, creating a flame shift that destroys the information in the subsequent codons. This protein cannot fold. Five amino acid substitutions in a critical folding element of the protein also exist, and Cys-40 is replaced by Gly. Likewise, the seminal gene from duiker encodes a substitution of Pro-42, likely to render the protein catalytically inactive. These defects in the gene force the conclusion that during much of its history, seminal RNase was an unexpressible pseudogene. Breukelmann et al. (1993) noted that the seminal RNase gene in giraffe might also be a pseudogene, based on the rate of sequence divergence. Lesions analogous to those in kudu have evidently been found in this gene and in a gene for hog deer (J. J. Beintema, 1995, personal communication). To show that these seminal genes were indeed not expressed in semen, seminal plasmas from 15 artiodactyls (ox, forest buffalo, Cape buffalo, kudu, sitatunga, nyala, eland, Maxwell's duiker, yellow-backed duiker, suni, sable antelope, impala, saiga, sheep, and Elds deer) were examined directly. Catalytically active RNase was not detected in the seminal plasma in significant amounts in any artiodactyl genus diverging before the Cape buffalo, except in Ovis. By Western blotting only traces of a monomeric, presumably pancreatic RNase, were detected in these seminal plasmas. In contrast, the seminal plasmas of forest buffalo, Cape buffalo and ox all contained substantial amounts of Western blotactive RNase. The seminal plasma from the Ovis genus (sheep and goat) was a notable exception. It contained significant amounts of both RNase activity and protein detectable by blotting. To learn whether RNases in the Ovis seminal plasma were derived from a seminal RNase gene, the RNase from goat seminal plasma was isolated, purified, and sequenced by tryptic cleavage and Edman degradation. Both Edman degradation (covering 80% of the sequence) and matrix-assisted laser desorptionionization (MALDI) mass spectroscopy showed that the sequence of
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the RNase isolated from goat seminal plasma is identical to the sequence of its pancreatic RNase (Jermann, 1995). This shows that the RNase in Ovis seminal plasma is not expressed from a seminal RNase gene, but rather from the O v is pancreatic gene. To confirm this conclusion, a fragment of the seminal RNase gene from sheep was sequenced and was shown to be different in structure from the pancreatic gene (Fig. 3). These results are not consistent with a model that the seminal RNase gene family gradually developed a new "seminal" function by stepwise point mutation and continuous selection under functional constraints in the seminal plasma following gene duplication. Rather, the duplicate RNase gene that founded the seminal lineage seems initially to have served no function at all. It therefore suffered damage, only to be repaired much later in evolution, after the divergence of kudu, but before the divergence of Cape buffalo, from the lineage leading to ox. Still more remarkable is the speed with which the seminal RNase gene was repaired. Clades (subbranches in a family tree) containing the saiga, duiker, and sheep are known in the early Miocene (23.8-16.4 million years before present), whereas clades containing the kudu and Cape buffalo are known in the late Miocene (11.2-5.3 million years before present). Despite the incompleteness of the fossil record, we can conclude that the damaged pseudogene was repaired extremely rapidly in only a few million years, some 25-30 million years after the gene duplication occurred and long after the presumed maximum time for the rehabilitation of a pseudogene (Jukes and Kimura, 1984; Marshall et al., 1994). Still more remarkable is the mechanism that these data suggest for the repair of the pseudogene. The data are not yet sufficient to establish this mechanism precisely. However, if the damage present in the kudu sequence was also present in the common ancestor, the repair would have required insertion of an additional nucleotide to repair a frame shift, and substitution at perhaps a half dozen positions elsewhere in the gene. The probability of this occurring by random insertion and point mutation (the classical Darwinian model) in just a few million years is vanishingly small. If the damage seen in the kudu pseudogene is indicative of the damage in the ancestor, a new evolutionary mechanism must be invoked to explain its extremely rapid repair. We have suggested (Trabesinger-Ruef et al., 1996) that the information required for this repair came from the sister gene for pancreatic RNase
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through a gene conversion event. This hypothesis suggests that information from the pancreatic RNase gene was used to repair the flame shift mutation in codon 114 as well as adjoining mutations. Consistent with this hypothesis, the expressed seminal RNase genes in this region are quite similar to the pancreatic gene, and the similarity extends some 66 base pairs into the 3' untranslated region, where it then ends abruptly. A recent search of the literature identifies some 2000 papers discussing gene conversion, containing considerable divergence of opinion concerning its existence, its mechanism, and its function. Gene conversion is well known in somatic mutation events, for example, in the process for generating diversity in antibodies (Wysocki and Gefter, 1989). The mechanism by which gene conversion takes place is not clear, however (Holliday, 1974). Indeed, it is not widely accepted that gene conversion is a mechanism used for good purpose during the evolution of higher organisms, nor is it clear what that purpose might be. One reason for the difficulties in obtaining a convincing argument for gene conversion in divergent evolution (that is, one that convinces the skeptics rather than one that persuades the believers) is that analyses are most often based on descendent structures in contemporary organisms, often distant from the conversion itself. Many doubt the importance of gene conversion as an evolutionarily important phenomenon because so many of the examples adduced to support it are minor changes in sequence with no obvious change in function. For example, data are available to clearly suggest that pseudogene and paralogous functioning genes most probably "talk" to each other (Eikenboom et al., 1994). However, the changes are few and have no obvious consequence on the functioning of the protein. Should the hypothesis of gene conversion as a mechanism for creating new biomolecular function (Trabesinger-Ruef et al., 1996) be confirmed, it offers a clear evolutionary role for gene conversion in higher organisms. New biomolecular function is believed to arise, at least in recent times, largely through recruitment of established proteins to play new roles following gene duplication. Under one model, one copy of a gene continues to evolve divergently under constraints dictated by the ancestral function. The duplicate, meanwhile, is unencumbered by a functional role, and is free to search protein "structure space." It may eventually come to encode new behaviors required for a new physiological function, and thereby confer selective advantage.
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This model contains a well-recognized paradox. Because duplicate genes are not under selective pressure, they should also accumulate mutations that render them incapable of encoding a protein useful for any function. Most duplicates therefore should become pseudogenes, inexpressible genetic information ("junk D N A " ) in just a few million years (Jukes and Kimura, 1984; Marshall et al., 1994). This limits the evolutionary value of a functionally unconstrained gene duplicate as a tool for exploring protein "structure space" in the search of new behaviors that might confer selectable physiological function. If gene conversion provides a mechanism for repairing badly damaged pseudogenes (Trabesinger-Ruef et al., 1996), it offers a mechanism for recovering at least some of the structural variation accumulated by pseudogenes as they drift under no selective pressure. This repair need not be "targeted" (that is, directed against only those defects that do damage to the functioning of the gene) to be useful. Rather, by effectively providing a mechanism for recombination between two types of genes, one constrained by function, the other free to search sequence space, the development of new function is facilitated. In this light, one final set of observations is important. In unexpressed seminal RNase sequences, the average ratio of nonsilent to silent substitutions is 2:1. This is close to that expected for random substitution in a gene serving no selected function (that is, a pseudogene). In a typical codon, changing the first or second nucleotide alters the meaning, whereas changing the third does not (this calculation must, of course, then be corrected for the details of the code and the distribution of amino acids within the typical protein). This ratio is consistent with the evidence previously presented that these seminal RNase genes are pseudogenes. In contrast, the nonsilent:silent substitution ratio within the pancreatic RNase family is less than 1:1. This is consistent with the ratio expected for a working gene wherein amino acid replacements are constrained by function. Most remarkable, however, is the ratio of nonsilent to silent substitutions (---10 : 1) in the repaired, expressed seminal RNases. Although the sample is small, this is consistent with a gene rapidly evolving to perform a new function, with amino acid substitutions rapidly introduced to provide new selected properties. This is as expected for a gene that recently emerged from a search of structure space as a pseudogene following repair using the gene conversion hypothesis (Trabesinger-Ruef et al., 1996).
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V. P H Y S I O L O G I C A L F U N C T I O N OF S E M I N A L RNase
The repair of a damaged RNase pseudogene involving a novel mechanism, followed by extremely rapid adaptive evolution for seminal RNase in a new environment to serve a new function remains hypothetical, and does not define, of course, the physiological function of seminal RNase. Such events, however, allow us to formulate some hypotheses as to what this function might be. First, the extremely rapid sequence divergence once the seminal RNase gene was repaired suggests that the protein has a function, and that the function is new in the seminal plasma. Function cannot be presumed for every protein expressed in large amounts in any particular tissue, making this confirmatory evidence valuable. Further, examination of both the proteins in the seminal plasma and the reproductive physiology in general suggests that the placental reproductive system has been doing considerable experimentation, because placental mammals became dominant following the extinction of the dinosaurs. Presumably, this experimentation continues today. The placental reproductive system has obvious advantages over alternative reproductive systems. These advantages are balanced, however, by specific disadvantages. For example, both the spermatozoa and the fetus are antigenic foreign entities, and the reproductive tract of mammals is under immune surveillance. Given this background, we can formulate several specific hypotheses that form the basis for future work in this area. (1) The immunosuppressivity of seminal RNase might be directly related to its physiological function, relating to the fact that the placental reproductive tract is under immune surveillance. An immunosuppressive RNase in the seminal plasma may assist in successful fertilization by antigenic spermatozoa. Cys-31 and Gly-111, introduced only after the damaged pseudogene was repaired, are important for immunosuppressivity. (2) The ability of seminal RNase to bind to seminolipid might be directly related to its physiological funtion. Seminolipid is a fusogenic lipid found in spermatozoa. Binding to seminal RNase is specific, both with respect to the protein (neither pancreatic RNase A nor the common ancestor bind) and the ligand (the anion, sugar, and lipid portions of the molecule all appear to be essential for binding) (T. R. Zankel, 1991, unpublished). Seminal RNase might help regulate the role of seminolipid in reproductive physiology. (3) The antispermatogenic activity long known in seminal RNase
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may be directly associated with its physiological function. Seminal RNase might help regulate the sperm count of bovids. (4) Seminal RNase might act directly against an extracellular RNA, including both retroviral R N A or extracellular RNA involved in cell-cell communication (Benner, 1988).
VI. C O N C L U S I O N S
The research outlined in this work is, of course, important to our understanding of ribonucleases. The strategy outlined in this work has wider implications for the study of structure, behavior, and physiological function of biological macromolecules in other systems. Again, it is noteworthy that the ribonuclease system is leading the way. Already, paleomolecular reconstructions are predicting models of the three-dimensional structure of proteins (Benner, 1989; Benner et aL, 1994b,c). Paleomolecular reconstructions are also serving to rationalize much of what is known about metabolism and biochemical physiology in organisms (Benner et al., 1989, 1993a). We believe that paleomolecular reconstructions will also be a tool in the future to connect biomolecular structure with themes interesting to the cellular, developmental, and organismal biologist. An example has already emerged from Hutchison and co-workers, who have recently resurrected an extinct ancestral promoter from mouse (Adey et al., 1994). We have recently studied the alcohol dehydrogenases from yeast expressed ---100 million years ago at a time when the first fleshy fruits allowed fermentation (D. H. Nakano, 1996, unpublished). As genome projects are completed, it will be possible to venture farther back in time. For example, at the start of the Cambrian, an explosion of multicellular phyla appeared within a very small space of time. As relatively complete genomes become available for several of the descendent phyla, it should be possible to reconstruct must of the genome of the most recent common ancestor of multicellular organisms. This should provide information concerning what was necessary to create multicellularity at its origin, in particular, what biomolecular innovation was associated with the origin of multicellularity. Likewise, some 1.5-2 billion years ago, the three "primary kingdoms" of contemporary life, the archaebacteria, eubacteria, and eukaryotes,
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diverged in uncertain order (Benner et al., 1989). Already, substantial parts of the genome of the common ancestor have been reconstructed (Benner et al., 1993a). With the full genome sequence of archaebacteria soon to be available, it should be possible in the near future using paleomolecular reconstructions to develop hypotheses concerning the molecular innovation made by the ancestor that allowed it to displace earlier forms of life.
ACKNOWLEDGMENTS
We are indebted to Dr. Alan Gentry, of the British Museum of Natural History, and Dr. Malcolm C. McKenna, Frick Curator in the Department of Vertebrate Palaeontology at the American Museum of Natural History, for helpful comments concerning artiodactyl evolution, and Jaap Beintema, for fascinating discussions and his many contributions to this field.
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Delcardayre, S. B., and Raines, R. T. (1995a). A residue to residue hydrogen-bond mediates the nucleotide specificity of ribonuclease A. J. Mol. Biol. 252, 328-336. Di Donato, A., Cafaro, V., and D'Alessio, G. (1994). Ribonuclease A can be transformed into a dimeric ribonuclease with antitumor-activity. J. Biol. Chem. 269, 17394-17396. Di Donato, A., Cafaro, V., Romeo, I., and D'Alessio, G. (1995). Hints on the evolutionary design of a dimeric RNase with special bioactions. Protein Sci. 4, 1470-1477. Dodge, R. W., Laity, J. H., Rothwarf, D. M., Shimotakahara, S., and Scheraga, H. A. (1994). Folding pathway of guanidine-denatured disulfide-intact wild-type and mutant bovine pancreatic ribonuclease A. J. Protein Chem. 13(4), 409-421. Dostal, J., and Matousek, J. (1973). Isolation and some chemical properties of aspermatogenic substance from bull seminal vesicle fluid. J. Reprod. Fertil. 33, 263-274. Eikenboom, J. C. J., Vink, T., Briet, E., Sixma, J. J., and Reitsma, P. H. (1994). Multiple substitutions in the von Willebrand factor gene that mimic the pseudogene sequence. Proc. Natl. Acad. Sci. U.S.A. 91, 2221-2224. Fitch, W. M. (1971). Toward defining the course of evolutionary minimum change from a specific tree topology. Syst. Zool. 20, 406-416. Fitch, W., and Margoliash, E. (1967). Construction of phylogenetic trees. Science 155, 279-284. Gonnet, G. H., and Benner, S. A. (1991). Computational Biochemistry Research at ETH. Technical Report 154, Departement Informatik, March (1991). Gonnet, G. H., Cohen, M. A., and Benner, S. A. (1992). Exhaustive matching of the entire protein sequence database. Science 256, 1443-1445. Haugg, M., Opitz, J. G., Raillard, S. A., Jermann, T. M., Soucek, J., Hiestand, P., and Benner, S. A. (1996). The structural origin of immunosuppressivity in seminal ribonuclcasc. Submitted. Holliday, R. (1974). Molecular aspects of genetic exchange and genc conversion. Genetics 78, 273-287. Ipata, P. L., and Felicioli, R. A. (1969). A convenient spcctrophotomctric assay for phosphodiesterases using dinucleoside monophosphates. FEBS Len. 1, 29-31. Ishizuka, I., Suzuki, M., and Yamakawa, T. (1973). Isolation and characterization of a novel sulfoglycolipid "seminolipid" from boar testis and spermatozoa. J. Biochem. (Tokyo) 73, 77-87. Jermann, T. M. (1996). "Der Ursprung und die Evolution der Ribonuklease aus dem Pankread und aus der Samenfltlssigkeit der Huftiere." ETH Dissertation No. 11059, Ztirich, Switzerland. Jermann, T. M., Opitz, J. G., Stackhouse, J., and Benner, S. A. (1995). Reconstructing the evolutionary history of the artiodactyl ribonuclease superfamily. Nature (London) 374, 57-59. Jolles, J., Jolles, P., Bowman, P. H., Prager, E. M., Stewart, C.-B., and Wilson, A. C. (1989). J. Mol. Evol. 28, 528-533. Jukes, T. H., and Kimura, M. (1984). Evolutionary constraints and the neutral theory. J. Mol. Evol. 21, 90-92. Kim, J. S., Soucek, J., Matousek, J., and Raines, R. T. (1995a). Catalytic activity of bovine seminal ribonuclease is essential for its immunosuppressive and other biological activities. Biochem. J. 308(Part 2), 547-550. Kim, J. S., Soucek, J., Matousek, J., and Raines, R. T. (1995b). Structural basis for the biological activities of bovine seminal ribonuclease. J. Biol. Chem. 270, 10525-10530. Kimura, M. (1982). The neutral theory as a basis for understanding the mechanism of evolution and variation at the molecular level. In "Molecular Evolution: Protein
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Raines, R. T., Toscano, M. P., Nierengarten, D. M., Ha, J. H., and Auerbach, R. (1995). Replacing a surface loop endows ribonuclease A with angiogenic activity. J. Biol. Chem. 270, 17180-17184. Richards, F. M., and Wyckoff, H. W. (1971). Bovine pancreatic ribonuclease. In "The Enzymes" (P. D. Boyer, ed.), 3rd Ed., pp. 4 and 647-806. Academic Press, New York. Rosenberg, H. F., Tenen, D. G., and Ackerman, S. J. (1989). Molecular cloning of the human eosinophil-derived neurotoxin: A member of the ribonuclease gene family. Proc. Natl. Acad. Sci. U.S.A. 86, 4460-4464. Saunders, M., Wishnia, A., and Kirkwood, J. G. (1957). The nuclear magnetic resonance spectrum of RNase. J. Am. Chem. Soc. 79, 3289-3290. Schluter, D. (1995). Uncertainty in ancient phylogenies. Nature (London) 377, 108-109. Schultz, D. A., Schmid, F. X., and Baldwin, R. L. (1992). Cis proline mutants of ribonuclease A. Elimination of the slow-folding forms by mutation. Protein Sci. 7, 917-924. Soucek, J., Hrub~i, A., Paluska, E., Chudomel, V., Dost~il, J., and Matousek, J. (1983). Immunosuppressive effects of bovine seminal fluid fractions with ribonuclease activity. Folia Biol. (Prague) 29, 250-261. Soucek, J., Chudomel, V., Potmesilova, I., and Novak, J. T. (1986). Effect of ribonucleases on cell-mediated lympholysis reaction and on GM-CFC colonies in bone marrow culture. Nat. Immun. Cell Growth Regul. 5, 250-258. Stackhouse, J., Presnell, S. R., McGeehan, G. M., Nambiar, K. P., and Benner, S. A. (1990). The ribonuclease from an extinct bovid. FEBS Lett. 262, 104-106. Stern, M. S., and Doscher, M. S. (1984). Aspartic acid-121 functions at the active-site of bovine pancreatic ribonuclease. FEBS Lett. 171, 253-256. Strydom, D. J., Fett, J. W., Lobb, R. R., Alderman, E. M., Bethune, J. L., Riordan, J. F., and Vallee, B. L. (1985). Amino acid sequence of human tumor derived angiogenic factor. Biochemistry 24, 5486-5494. Trabesinger-Ruef, N., Jermann, T., Zankel, T. R., Durrant, B., Frank, G., and Benner, S. A. (1996). Pseudogenes in ribonuclease evolution: A source of new biomacromolecular function? FEBS Lett. in press. 382, 319-322 Trautwein-Fritz, K. (1991). "Construction of an Improved Expression System for Bovine Pancreatic Ribonuclease A and Construction and Characterization of RNase A Mutants." ETH Dissertation No. 9613, Ztirich, Switzerland. Trautwein-Fritz, K., and Benner, S. A. (1991 a). Site-directed mutagenesis of bovine pancreatic ribonuclease: Lysine-41 and aspartate-121. FEBS Lett. 281, 275-277. Trautwein-Fritz, K., and Benner, S. A. (1991b). High level expression of bovine pancreatic RNase A." Proceedings of the International Symposium on Site Directed Mutagenesis and Protein Engineering" (M. R. EI-Gewely, ed.), pp. 53-61. Elsevier, New York. Vescia, S., and Tramontano, D. (1981). Antitumoral action of bovine seminal ribonuclease. Mol. Cell. Biochem. 36, 125. Vescia, S., Tramontano, D., Augusti-Tocco, G., and D'Alessio, G. (1980). In vitro studies on selective inhibition of tumor cell growth by seminal ribonuclease. Cancer Res. 40, 3740. Vos, J. P., Lopes-Cardozo, M., and Gadella, B. M. (1994). Metabolic and functional aspects of sulfogalactolipids. Biochim. Biophys. Acta Lipid 1211, 125-149. Westheimer, F. H. (1992). The role of phosphorus in chemistry and biochemistry. ACS Symp. Ser. 486, 1-17. Wysocki, L., and Gefter, M. L. (1989). Gene conversion and the generation of antibody diversity. Annu. Rev. Biochem. 58, 509-531.
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8 Evolution of Vertebrate Ribonucleases" Ribonuclease A Superfamily J A A P J. B E I N T E M A , * H E L E E N J. B R E U K E L M A N , * ANTONELLA CARSANA,t AND ADRIANA FURIAt * Department of Biochemistry University of Groningen 9747 AG Groningen, The Netherlands t Department of Organic and Biological Chemistry University Federico II of Naples 80134 Naples, Italy
I. The Ribonuclease Superfamily A. Classification B. Structure, Evolution, and Enzymatic Properties C. Mammalian Ribonucleases 2: Nonsecretory or Neurotoxin-type II. Mammalian Ribonucleases 1: Pancreatic-type A. Most Parsimonious Tree B. Primate Ribonucleases C. Rodent Ribonucleases D. Artiodactyi Ribonucleases References
I. THE RIBONUCLEASE SUPERFAMILY A. Classification The pyrimidine-specific ribonuclease A superfamily constitutes a group of homologous proteins with well-characterized and sequenced
RIBONUCLEASES: STRUCTURESAND FUNCTIONS
245 Copyright 9 1997by AcademicPress, Inc. All rights of reproduction in any form reserved.
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members isolated from many mammalian, avian, reptilian, and amphibian sources. No representatives have yet been found in fishes or nonvertebrate taxa. Possibly, a protein that has some sequence similarity with members of the ribonuclease A superfamily or has a similar fold of the polypeptide chain without demonstrable ribonuclease activity exists in these more distantly related taxa, but this has not yet been found. Distantly related members of the ribonuclease A superfamily differ at more than 50% of the amino acid positions. The nomenclature of members of the ribonuclease A superfamily is rather chaotic, especially that of paralogous ribonucleases identified in the same species. Zhou and Strydom (1993) proposed a very useful designation using numbers 1-4 for human ribonucleases. This nomenclature is extended here to other mammalian species, and a subclassification (not shown) may be proposed for paralogous ribonucleases belonging to the same family. Thus, we classify the primate eosinophil cationic proteins as ribonucleases 2, and do not separate them into a third group, done by Zhou and Strydom (1993). However, as discussed later, it is not yet possible to define orthologous relationships between mammalian ribonuclease families and those identified in other vertebrate classes. Amino acid sequences of bovine pancreatic ribonuclease and members of the superfamily not belonging to the mammalian (pancreatic-type) ribonuclease 1 family are presented in Fig. 1, together with literature sources of the sequences. Nine separate families with sequences that are more than 50% identical can be distinguished within the superfamily: 1. Mammalian (pancreatic-type) ribonucleases 1. These, often designated as "secretory" ribonucleases, will be discussed in Section II of this chapter. 2. Mammalian (nonsecretory or neurotoxin-type) ribonucleases 2. This family includes bovine kidney ribonuclease K2; porcine ribonucleases PL1 from liver and PK3 from kidney; human liver, spleen, and urine U~ ribonucleases, which are identical to human eosinophil-derived neurotoxin (EDN); and human eosinophil cationic protein (ECP). 3. Mammalian ribonucleases 4. Human plasma ribonuclease HT-29 and ribonucleases from bovine (BL4), porcine (PL3), and rat (RL3) liver belong to this family. 4. Mammalian angiogenins. 5. Chicken liver ribonuclease. 6. Chicken myelomonocytic cell and bone marrow cell ribonucleases. These amino acid sequences are 85% identical, with even more sequence identity at noncoding positions in the nucleotide sequences.
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7. Turtle pancreatic ribonuclease. 8. Iguana pancreatic ribonuclease. So far, snapping turtle (Chelydra serpentina) and iguana (Iguana iguana) are the only investigated nonmammalian species with sufficient amounts of ribonuclease in their pancreas to permit amino acid sequence studies. Although Barnard (1969) reported a ribonuclease content of 53 ~g/g tissue in the pancreas of the crocodile species Cayman crocodilus, we found only 0.2 /~g/g in the pancreas of Crocodilus niloticus (Zhao, 1996, unpublished). 9. Frog ribonucleases.
B. Structure, Evolution, and Enzymatic Properties Three-dimensional structures have been published for bovine pancreatic and seminal ribonucleases, and also for human angiogenin (Acharya et al., 1994) and the ribonuclease from Rana pipiens oocytes (onconase; Mosimann et al., 1994). The molecular structures of these enzymes are very similar, and the active site structures and hydrophobic clusters in the interior of the proteins (Kolbanovskaya et al., 1993) have been conserved. The alignment of the sequences in Fig. 1 is based predominantly on conservation of the active site structure and elements of secondary structure. However, external loops show more variation and an unambiguous alignment is often not possible. In several cases, one has to conclude that an evolutionary alignment based on sequence similarity is not necessarily the one that can be derived from the superposition of three-dimensional structures (Mosimann et al., 1994). We have not yet performed an extensive evolutionary analysis of all ribonuclease sequences. However, evolutionary trees of limited numbers of sequences have been published by Iwama et al. (1993) (Fig. 2) and Zhou and Strydom (1993). The four frog sequences have not been included in these trees, but it is rather sure that they group together and may be considered as the first diverging branch in the ribonuclease tree. However, the branching order of the remaining eight reptile, bird, and mammalian families is difficult to derive, although we have a slight preference to join the reptilian sequences with the angiogenins. A striking observation can be made from Fig. 2: there are large differences in evolutionary rates between three mammalian ribonuclease families, with the ribonucleases 4 having the slowest rates and the nonsecretory neurotoxin-type ribonucleases 2 having the highest rates. The pancreatic-type
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....
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Ribonuclease A Superfamily Evolution
249
ribonucleases 1 evolved at an intermediate rate. Different evolutionary rates between protein families within one superfamily have also been observed in alcohol dehydrogenases, with the class I enzymes evolving much faster than the class III enzymes (Danielsson et aL, 1994). This has been explained by the class III alcohol dehydrogenases being metabolically fixed enzymes, whereas the class I enzymes have been emerging in function and show more adaptations to the environment. A similar explanation may be proposed for the differences in evolutionary rates between the ribonuclease families. There are also differences in substrate specificity and other enzymatic properties between ribonucleases belonging to different families (Sorrentino and Libonati, 1994), but within a family there is less variation (Beintema, 1987). A characteristic difference between ribonucleases belonging to different families is their relative preference for cleavage of
Fig. 1. Alignment of the amino acid sequences of bovine pancreatic ribonuclease and other members of the ribonuclcasc supcrfamily [except those of other mammalian pancreatic-type ribonuclcascs 1, which arc presented in Bcintcma et al. (1988c) or in Fig. 4]. Deletions in the sequences arc indicated by dashes. Residues conserved in all sequences arc given on the bottom line. Ribonuclcasc 2 family includes ribonuclcasc U~ or human (h)EDN (Bcintcma et al., 1988b: Hamann et al., 1989: Roscnbcrg et aL, 1989): human (h)ECP (Barker et al., 1989); othcr primate EDNs and ECPs (Roscnbcrg et ak, 1995) arc as follows: ch, chimpanscc (Pan troglodytes); g, gorilla (Gorilla gorilla); o, orangutan (Pongo pygmaeus): me, crab-eating macaque (Macaca fascicularis); mm, cotton-top tamarin or marmoset (Saguinus or Leontocebus oedipus); Roscnbcrg and Dyer (1995) present an F instead of R at position 106 (hEDN and hECP numbering) in mmEDN; ox kidney ribonuclease K2 (Iric et al., 1988); and pig kidney ribonuclease PK3 (lwama et al., 1993). Ribonuclease 4 family includes ox liver, BL4 (Hosoya et al., 1990); pig liver, PL3 (Hofsteenge et al., 1989); human plasma (Zhou and Strydom, 1993) and rat liver (RL3) ribonucleases (S. Jarai and J. Hofsteenge, 1995, personal communication); angiogenins, from humans (Strydom et al., 1985), ox (Macs et aL, 1988), pig and rabbit (Sus scrofa and Oryctolagus cuniculus) (Bond et al., 1993), and mouse (Bond and Vallee, 1990); chicken liver ribonucleasc (Hayano et al., 1993); chicken myelomonocytic cell ribonuclease (Graf and Nakano, 1992); chicken bone marrow cell ribonucleasc (Klenova et al., 1992); turtle pancreatic ribonuclease (Beintema et al., 1988c): iguana pancreatic ribonuclease (Zhao et al., 1994); frog ribonucleases from Rana catesbeiana liver (Nitta et al., 1989) and Rana pipiens oocytes (onconase) (Ardelt et al., 1991 ); lectin from Rana japonica eggs (fr. jap. lc.) (Kamiya et al., 1990); and lectin from Rana catesbeiana eggs (fr. cat. lc.) (Titani et al., 1987). [Note: The C-terminal regions of ch, g, o and mc EDN and ECP presented by Rosenberg et al. (1995) have been derived from primer sequences of human EDN and ECP, respectively, used for PCR amplification (H. F. Rosenberg, personal communication, 1996)].
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250
Jaap J. Beintema et al.
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the polymeric substrates poly(C) or poly(U) (Sorrentino and Libonati, 1994). Several members of the superfamily show very low activity in the standard assay using yeast RNA. These are the angiogenins and ECP. They lack a basic amino acid residue at either position 66 or 122 (bovine pancreatic ribonuclease numbering), which may explain their low activity (Beintema, 1989). However, these proteins have descended from ribonucleases with higher enzymatic activity. In this respect they resemble zymogens, which have descended from active proteases. It is possible that the less active ribonucleases require another factor to become more active. Indications in this direction have been obtained for human angiogenin (Acharya et al., 1994; Riordan, Chapter 14, this volume). Mammalian (pancreatic-type) ribonucleases 1 catalyze two reaction steps: a transphosphorylation with formation of 2',3' cyclic nucleotide phosphates, followed by hydrolysis of these cyclic derivatives. The two active site histidine residues participate in a symmetrical way as reciprocal hydrogen donor and acceptor. The hydrolysis reaction is much slower
8
Ribonuclease A Superfamily Evolution
251
than the transphosphorylation, and members of several other families exhibit very low or even undetectable hydrolysis activity (Sorrentino and Libonati, 1994; Beintema, 1991).
C. Mammalian Ribonucleases 2: Nonsecretory or Neurotoxin-type
The chemically determined amino acid sequence of human nonsecretory (Beintema et al., 1988b) or liver ribonuclease (Sorrentino et al., 1988), or eosinophil-derived neurotoxin, is identical to that deduced from the DNA sequence (Hamann et al., 1989; Rosenberg et al., 1989) except for the identity of the residue at position 7. Whereas a tryptophan residue was predicted from the DNA sequence, no amino acid residue could be identified by chemical means. Hofsteenge et al. (1994) have demonstrated that an aldohexopyranosyl residue is linked via a C - C bond to the indole ring of this tryptophan. This is a new type of linkage between a carbohydrate and a protein, and has not been found in other glycoproteins. The C-terminal parts of the signal peptide sequences preceding the mature sequences of EDN and ECP are -Gly-Ser-Leu-His-Val and -Gly-Ser-Leu-His-Ala, respectively. The C-terminal valine in the EDN sequence does not conform to the consensus sequence for cleavage of signal peptides (Von Heijne, 1984; Hamann et al., 1989). In addition, Sakakibara et al. (1992) have isolated from the urine of pregnant women a variant of EDN that has the N-terminal sequence Ser-Leu-His-ValLys-Pro-, etc., which is identical to the EDN sequence if cleaved after a glycine residue in the signal peptide sequence, thus conforming to a consensus cleavage site. Therefore, not only EDN, but also ECP, is probably formed by signal peptidase cleavage of the Gly-Ser sequence, followed by further removal of the four N-terminal residues in both proteins. The proline residue at position 2 in all members of the family (Fig. 1) may prevent further processing of the mature protein (J. Hofsteenge, 1995, personal communication). Rosenberg et al. (1995) have sequenced homologs of EDN and ECP in five nonhuman primate species including three apes, an Old World monkey, and a New World monkey. They found that these proteins exhibit a higher evolutionary rate than the pancreatic-type ribonucleases 1, which is in agreement with Fig. 2 and trees derived from sequences of ribonucleases 1 from different mammalian orders. The difference between
252
Jaap J. Beintema et al.
EDN and ECP, respectively, of an Old World monkey and humans is about 15%, whereas that between pancreatic-type ribonucleases of both taxa is 11% (Beintema, 1990). Only one ribonuclease 2 homolog was identified in the genome of cotton-top tamarin, a New World monkey, with sequence characteristics intermediate between EDN and ECP. This observation suggests that EDN and ECP are the products of a gene duplication that occurred after the divergence of the Old World monkeys from the New World monkeys, and is in agreement with an earlier prediction made by Beintema et al. (1988c). As in ECP, the tamarin sequence has no lysine or arginine at position 122 (bovine pancreatic ribonuclease numbering), and, therefore, this protein shows a low enzymatic activity, as expected (Rosenberg and Dyer, 1995). This would mean that no active ribonuclease 2 may occur in this species. However, as suggested before, interaction with an additional factor may be required for normal activity.
II. M A M M A L I A N
RIBONUCLEASES
1: P A N C R E A T I C - T Y P E
A. Most Parsimonious Tree
The latest extensive evolutionary analysis of amino acid sequences of mammalian pancreatic-type ribonucleases was performed by Beintema et al. (1986) and resulted in a most parsimonious tree of 39 complete sequences (see also Beintema et al., 1988c). In a later analysis (Fitch and Beintema, 1990) the sequences of Spalax ehrenbergi, a rodent species (Schtiller et al., 1989), and of bovine brain ribonuclease (Watanabe et al., 1988) were added to the tree (Fig. 3). The Spalax sequence became
Fig. 3. Evolutionary tree of mammalian pancreatic-type ribonucleases. The most parsimonious tree observed is shown. Horizontal distances are proportional to the corrected number of nucleotide substitutions shown, rounded to the nearest integer, in the form a + b, where a is the number of uncorrected substitutions demanded by parsimony, and b is the number of additional substitutions required by the correction procedure. No b is shown if no correction is required. Vertical distances are solely to separate lineages (Fitch and Beintema, 1990). The diamonds ( 9 ) represent three demonstrated gene duplications. The total number of substitutions in the tree is 554. [For scientific names of species, see Beintema et al. (1988c).]
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the sister taxon of the other myomorph rodents, which is conventional. The new paralogous bovine brain sequence joined to the bovine seminal sequence at a point indicating that the ribonuclease gene duplicated at least twice at approximately the origin of the true ruminants. However, the reliability of these trees has not yet been tested by resampling methods ("bootstrap," "jackknife"), which are currently important procedures in molecular evolutionary studies (Swofford and Olsen, 1990). Jermann et al. (1995) have reconstructed 13 ancient ribonuclease sequences as ancestors in the evolution of the sequenced taxa in the subfamily of the Bovinae back to the artiodactyl ancestor, using the most parsimonious tree. These sequences have been prepared, and several of their properties have been investigated. Stable and enzymatically active proteins were obtained. Going backward in time, a fivefold increase in activity against double-stranded RNA was found. This feature can be related to a simultaneous increase in net positive charge and a less predominant role as a digestive enzyme (Beintema et al., 1988c). Jermann et al. (1995) suggest that the replacement of Gly at position 38 by aspartic acid may have a specific role in species with foregut digestion, whereas our interpretation of the effect of net charge on the activity on doublestranded RNA is more general (Beintema et al., 1988c). However, an excess of positive charge is not sufficient for destabilizing nucleic acid double-helical structures, as eosinophil cationic protein, a very basic member of the ribonuclease superfamily, lacks such an effect (Sorrentino and Glitz, 1991). Jermann et al. (1995) also found that, going backward in time, the ancestral ribonucleases become slightly less stable. However, they did not take into account possible posttranslational modifications. The less stable ancestral ribonucleases have two additional recognition sites for carbohydrate attachment at positions 62 and 76. Because the proteins were expressed in a prokaryotic system, they were synthesized without carbohydrate. However, if these ancestral ribonucleases were as extensively glycosylated as the porcine enzyme, the carbohydrate moieties may be expected to have exercised a protective effect (Wang and Hirs, 1977).
B. Primate Ribonucleases
Structural studies on the ribonucleases from human pancreas (Beintema et al., 1984) and urine (Beintema et al., 1988a) showed that they
8
Ribonuclease A Superfamily Evolution
255
are identical except for the presence of an additional threonine residue at the C terminus and complete glycosylation of the three Asn-X-Ser/ Thr sequences in the urine enzyme, whereas the pancreatic enzyme has carbohydrate attached to only one of these sequences in part of the molecules. Probably the enzyme preparation used for the structural studies of the pancreatic enzyme by Beintema et al. (1984) was of poor quality (Weickman et al., 1981), because Rib6 et al. (1991) isolated, from fresh human pancreas, ribonuclease preparations containing much more carbohydrate, attached to Asn-34 in all, to Asn-76 in about half, and to Asn-88 in a small proportion of the molecules (Rib6 et al., 1994). Another primate pancreatic ribonuclease with known amino acid sequence is that of the langur monkey (Beintema, 1990), a species with ruminant-like digestion. The amino acid sequences of the two primate ribonucleases (Fig. 4A) differ at 14 (11%) of the positions. This leaves no doubt that they are located on the same branch of a most parsimonious tree. A comparison of these two ribonucleases is interesting because we have here an example of two species within one mammalian order, one of which has a ribonuclease that shows adaptations to ruminant-like digestion. In previous reports (Beintema and Lenstra, 1982; Beintema et al., 1976, 1988c) we advanced the hypothesis that species with ruminant-like digestion not only have a higher ribonuclease content in their pancreas (Barnard, 1969), but also have smaller excess of positive charge and less attached carbohydrate, which is in agreement with data presented in Table I.
C. Rodent Ribonucleases
Several rodent ribonuclease sequences have been determined since our last review (Beintema et al., 1988c), in which we presented 12 sequences from 11 rodent species, including the cDNA sequences of mouse and rat (Schtiller et al., 1990). It has already been mentioned that the mole rat sequence (Spalax ehrenberg# SchUller et al., 1989) is correctly positioned in the most parsimonious tree as a sister taxon of the other myomorph rodents. This sequence turned out to be one amino acid longer in the hypervariable region (15-24) than those of other mammals, due to a leucine insertion between residues 21 and 22 (Fig. 4B). No differences in amino acid sequence were found between the ribonucleases from four chromosomal species of S. ehrenbergi ( J e k e l et al.,
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257
Ribonuclease A Superfamily Evolution
TABLE I Several Characteristics of Human and Langur Pancreatic Ribonucleases
Characteristic Fraction in tissue Net charge Number of possible glycosylation sites (Asn-X-Ser/Thr sequences) Extent of glycosylation
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1990). The ribonuclease from another mole rat species, Spalax leucodon, differs from that of S. ehrenbergi in having alanine instead of proline at position 42 (Zhao et al., 1993). DNA sequences of the complete mouse
Fig. 4. Amino acid sequences of (pancrcatic-typc) ribonuclcases 1. Complctc scqucnccs are given in the top line of each section ( A - G ) , and the differences are given in the lines below. Deletions in the sequences are indicated by dashes and unidentified residues are indicated by x. (A) Human and langur monkey (Presbytis entellus) ribonucleases. (B) Golden hamster (Mesocricetus auratus), Chinese hamster (Cricetulus longicaudatus), muskrat (Ondatra zibethica), and mole rat (Spalax ehrenbergi) (Schiiller et al., 1989) ribonucleases. The sequence of Spalax leucodon ribonuclease differs from that of S. ehrenbergi by having alanine instead of proline at position 42 (Zhao et al., 1993). (C) Porcupine (Hystrix cristata and Artherurus) and gundi (Ctenodactylus gundi) ribonucleases. (D) Ruminant pancreatic ribonucleases of ox, goat, giraffe, hog deer (Axis porcinus) and roe deer; *, A/V polymorphism at position 64. (E) Ruminant brain ribonucleases of ox, sheep, giraffe, hog and roe deer, and mouse deer (Tragulus javanicus). (F) Ruminant seminal ribonucleases of ox, sheep, giraffe, and hog and roe deer; 6, one nucleotide in codon deleted; +6, two additional nucleotides; A, stop codon. (G) Ribonucleases from mouse deer and bactrian camel; **, uncertain transition between signal and mature sequence. In the camel sequence, the sequence His-Asp was found at positions 11 and 12 (indicated as "hd") instead of the active site sequence Gin-His. This sequence was also observed in several pancreatic sequences, and may be an artifact because of the close position of the bands of a consecutive C-G pair on the sequence gel.
258
Jaap J. Beintema et
al.
ribonuclease gene (Samuelson et al., 1991), and almost complete coding sequences of Chinese hamster (Haugg and Schein, 1992), gundi (a rodent species with uncertain phylogenetic position), and artherurus (an Old World porcupine species) ribonucleases (J. A. Kolkman, 1995, unpublished), have been determined with the polymerase chain reaction (PCR). In Fig. 4, B and C, the latter three sequences are compared with those of related rodents. The Chinese hamster sequence differs from that of the golden hamster at four positions, three of which are the same in golden hamster and muskrat, another cricetid rodent species. As a rule a single band is detected by Southern blot analysis of the genomic DNA of nonruminant mammalian species (Breukelman et al., 1993). The human gene product is not only detected in the pancreas but in several other organs and body fluids as well (Beintema et aL, 1988a). Likewise, transcripts of the mouse ribonuclease gene have been detected not only in the pancreas but also in the parotid gland (Samuelson et al., 1991). Grassi Zucconi et al. (1992) have investigated the presence of rat ribonuclease in different cerebral regions by RNA blot analysis, in situ hybridization, and immunocytochemical analysis, and have demonstrated that it is synthesized in the neurons of rat brain.
D. Artiodactyl Ribonucleases
Carsana et al. (1988) have determined the structure of the gene of bovine pancreatic ribonuclease. The coding region is devoid of introns, whereas the 5' untranslated region contains an intron. This structural feature is typical of all members of the ribonuclease superfamily that have been sequenced at the gene level. The intervening sequence is endowed with signals that might act as regulatory elements, which suggests that bovine pancreatic ribonuclease might be expressed under the control of two different promoters. However, in later studies (Carsana et al., 1992) only sequences in the 5' flanking region of the gene were demonstrated to bind one or more factors present in nuclear extracts prepared from bovine pancreas. Nuclear extracts from the pancreas, but not from HeLa or bovine aorta endothelium cells, interact with these elements. Ribonucleases very similar to the pancreatic one have been demonstrated in a number of bovine tissues and body fluids (Carsana et al., 1988). However, two structurally different secretory ribonucleases occur
8
Ribonuclease A Superfamily Evolution
259
in addition to the pancreatic enzymes, namely, the bovine seminal and brain ribonucleases. The seminal enzyme is a dimer with subunits covalently linked by two disulfide bridges between half-cystine residues occurring at positions 31 and 32 (D'Alessio et al., 1991). An interesting feature is that its quaternary structure occurs in two forms, the major one of which has the N-terminal segments of the two subunits (including the active site region near His-12) exchanged, whereas there has been no exchange in the minor form (Piccoli et al., 1992). The complete cDNA encoding the seminal enzyme has been published by Preuss et al. (1990). The brain enzyme has been sequenced by Watanabe et al. (1988). Sasso et al. (1991) have determined its gene structure, which has the same structural features as pancreatic ribonuclease. The polypeptide chain of the brain enzyme is endowed with peculiar structures, such as a 17-residue carboxy-terminal extension that is highly hydrophobic and characterized by a high proline content and two O-linked oligosaccharide chains. The DNA sequence showed that one of the O-linked serine residues in the sequence of Watanabe et al. (1988) should be a threonine, and that the enzyme contains one more leucine at the C terminus. The pancreatic enzyme is expressed in many tissues, but expression of the seminal enzyme has been observed only in seminal vesicles, whereas expression of the brain enzyme has not only been observed in all regions of the brain examined (Sasso et al., 1991), but also in lactating mammary glands (Furia and Carsana, 1995, unpublished observations; Zhao, 1996). As already mentioned, evolutionary analysis of mammalian pancreatictype ribonucleases indicated that the presence of the three homologous bovine enzymes is due to gene duplications that occurred during the evolution of ancestral ruminants (Fitch and Beintema, 1990). Later on, this conclusion was confirmed by Southern blot analysis of the genomic DNA of mammalian species. If the sequences encoding bovine pancreatic or brain ribonuclease are used as a probe, a single band has generally been detected in DNA from nonruminant artiodactyls and other mammals, whereas a complex pattern of hybridization appears in genomic DNA of ruminant species (Breukelman et al., 1993). Lysozyme, another enzyme with an important function in ruminants, presents features very similar to those observed for ribonucleases (Joll6s et al., 1989). Genomic DNA coding for pancreatic-type ribonucleases from several ruminant species has been amplified by PCR, cloned, and sequenced, and several sequences were obtained (Confalone et al., 1995). As a rule, we could easily assign each of these sequences as pancreatic, seminal, or brain type, not only from the presence of several typical amino acids,
260
Jaap J. Beintema et aL
but also from a most parsimonious analysis, in which the three types form three different groups with 100% bootstrap values (Fig. 5). Figure 4D shows the amino acid sequences of the pancreatic ribonucleases from five species. The sheep and roe deer sequence are identi-
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8
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cal to the earlier determined ones. Only an Ala/Val heterogeneity was found at position 64 in sheep, at the same position where roe deer ribonuclease exhibits Ala/Ile polymorphism (Beintema et al., 1988c). Giraffe pancreatic ribonuclease has a Tyr/Asn heterogeneity at position 76, and position 28 is occupied by Glu instead of the previously reported Gln (Beintema et al., 1988c). Hog deer pancreatic ribonuclease differs from that of red deer (also belonging to the deer subfamily of the Cervinae) at positions 24, 91, and 122, where red deer pancreatic ribonuclease has the same residues as bovine pancreatic ribonuclease. Figure 4E shows complete derived amino acid sequences of brain ribonucleases from five ruminant species and the partial sequence of the brain enzyme from mouse deer. The latter is a species belonging to the Tragulidae, which is the earliest diverged taxon of the true ruminants. These sequences differ little from that of bovine brain ribonuclease, and exhibit a low proportion of nonsynonymous substitutions versus synonymous ones (Confalone et al., 1995). The strongly basic character is conserved in all of them. Only the partial mouse deer sequence deviates relatively strongly. It exhibits a replacement of Cys-72 by Ser, which may indicate that it represents a pseudogene (see later). Bovine brain ribonuclease has an Asn-Ile-Thr sequence at positions 62-64 with carbohydrate attached in all molecules (Watanabe et al., 1988). This site is conserved in the other sequences, and additional recognition sites for N-glycosylation occur in the other five sequences at positions 76-78 (Asn-Ser-Thr) and 80-82 (Asn-Ile-Thr). Like bovine brain ribonuclease, the sequences from sheep, giraffe, and deer have extensions at the carboxylic end. These extensions are located in a region of about 50 nucleotides, this sequence is repeated twice in the 3' noncoding regions of bovine pancreatic, seminal, and brain ribonucleases, with a region rich in C and A in between (Beintema et al., 1988c). These features may represent hot spots for gene rearrangement. The sequence of this part of the 3' noncoding sequence of bovine pancreatic ribonuclease is shown in Fig. 6. The 3' noncoding sequences of mouse and rat ribonuclease do not show significant sequence similarity with the bovine sequences (Schtiller et al., 1990). However, the cDNA sequence of human pancreatic ribonuclease (Seno et al., 1994) not only confirmed its previously determined amino acid sequence, but also shows sequence similarity with the repeated sequence in the bovine sequences (Fig. 6). In Figure 6 we stress the sequence similarity of the human sequence with the second repeat, with a short preceding sequence with some similarity to the first repeat, and long stretches of deleted sequence regions. However, this region of the human sequence may also be aligned with the first repeat.
HUMAN PANCREAS
TTTGATGCTTCAGTGTAGGT~TCTA~TAAGC~AGAGCAGCAAGATGCA~CA~TTCATCACAAAGGCACCTGCCT~TC~CCTC no r e c o g n i z a b l e h o m o l o g y in the TTTGATGCTaCtGTGTAGGgCTCcACaT-A~CAaAcCAGtgAGATGt( (rest of the 3 ' - n o n c o d i n g sequence TTTGATGCTTCt GTGgAGGaCTCTACCTAAGGt CAGAGCAGCgAGATaCcCCACcTc-TCtCAAcctCAt---CCTCTCC
BOVINE PANCREAS HUMAN PANCREAS
ATGTTTCCTTGT~TGG~3CAATAGCTCAAGTTAGTTACK~C~TCTTATCTCTGCGCA~CTTA~CAGAAA~ACACACA~ --AcAGCT-GcCTCTTccCTCT . . . . . . . . . .
BOVINE PANCREAS MOUSE P A N C R E A S
(Repeat : ) BOVINE PANCREAS HUMAN P A N C R E A S
BOVINE PAN(~EAS BOVINE BRAIN SHEEP BRAIN GIRAFFE BRAIN HOG DEER BRAIN SIZE
OF GAPS:
) )
( end of repeat ) AgGaTTCCcTGgCaTGaaaGCAATAaCTCAAGcTAG~AaGtCTtcTATCc - T c c T T C C c T G c t g T G a a a G a A g T A A C T a c A G T T A G .... ~ 3 C T C c T A T t T ( )
CACT TCATC2W:AAAGGCACCTGCCTCTCC CCT~CTTGTCCTOG000CAATAGCTCAR~TTAGTTAG~-42TCTT ~ T C A K G T GTAGGTCTCTACCTAA~GAGCAGCAAGATGCAC .... t T C T T .AgOcTTCt c ~._aCt T O a O O O C A A T A a C T C ~ A G g T ~TOgggO~TG . . . . . CTCTt aCx:t gcc a C A c c c g t a C c c t cact g C C A C c T C c aC2&C ~ c ~ _ a C t TGa(R~IC2&gTAaC T C A R ~ T T A G g T .... t T C T T TTTIIATOgTg(:gGTa . . . . . CTCTt aC~zt gcc a C A c c c C t a C c c t cact g C C A C c TCc aC2&C2%AgcGC ~ T O g ~ G . . . . . CTCTt aCct gcc aC c T C c aCAC2kAgc aC ~ c T G g C t c OaI3MI~.AAT~t T(:g l & l ~ A O a T .... t T C T c ~TOgTTC~OTG . . . . . tTCTt aCct gcc aCcc caCt a C c c t c act g C C A g c TCc a C A C A A g c aC AgOcTTCt c TGgCt T ~ T A a t TCAR~.._A(~gT .... t T C T T
I-5-- I
I. . . . . . . 20 . . . . . . . .
I
t-6--I
Fig. 6. Comparison of the nucleotide sequences of the C-terminal coding sequence (starting with the conserved codons for Phe-120 and Asp-121: T I T GAT) and part of the 3' noncoding region of bovine, mouse, and human pancreatic ribonucleases. Deletions in the sequences are indicated by dashes. The bovine sequence and identical nucleotides in the lines below are presented with capital letters. The third line of the bovine sequence represents a duplicated segment, and in this sequence fragment and in that of the human sequence, identical nucleotides with the first bovine repeat are presented with capital letters. Stop codons are underlined. Fig. 7. Comparison of the nucleotide sequences of the C-terminal coding sequence (starting with the conserved codons for Phe- 120 and Asp- 121: 7TT G AT) and part of the 3' noncoding region of bovine pancreas and ruminant brain-type ribonuclease sequences. Deletions in the sequences are indicated by dashes. The bovine pancreas sequence and identical nucleotides in the lines below are presented with capital letters. Identities in all five sequences are presented in boldface type. In-frame stop codons are underlined.
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Figure 7 shows the nucleotide sequences of the ruminant brain ribonucleases with the C-terminal extensions. These sequences are homologous with the pancreatic sequences if rather extensive deletions are introduced directly after the conserved Phe-Asp (120-121) sequence and more downstream in the 3' noncoding region. Bovine, sheep, and hog deer ribonucleases have their C-terminal residues in the abovementioned repeated structure. The deer sequences extend eight residues (Fig. 4E) more than that of sheep before being interrupted by an inphase stop codon. A deletion of 20 nucleotides in the sequence encoding the giraffe sequence (Fig. 7) results in a strongly deviating C-terminal amino acid sequence because of a different reading frame (Fig. 4E). Figure 4F shows derived amino acid sequences of sheep, giraffe, and roe and hog deer seminal ribonucleases. The deduced amino acid sequence of water buffalo (river type) seminal ribonuclease is identical to the bovine sequence, and only synonymous substitutions are observed. However, deletions occur in DNA fragments of three genomes (1 nucleotide in the codons at positions 38 in sheep and 20 in roe deer, and 16 nucleotides in the codons at positions 21-26 in giraffe) (Confalone et al., 1995), and an insertion of 5 nucleotides occurs near residue 10 in that of hog deer (Fig. 4F), causing out-of-frame joining of the downstream regions and introducing nearby located stop codons. In addition, many point mutations create a high proportion of nonsynonymous versus synonymous substituted codons in the reading frames corresponding to the one encoding the bovine enzyme. His-12, which is essential for catalytic activity of mammalian ribonucleases, is replaced by leucine in the giraffe, by serine in the sheep, and by tyrosine in the hog deer sequence, respectively. Position 115 in the giraffe sequence is encoded by a stop codon. In the deer sequences, the active site residue Lys-41 is replaced by arginine, and Cys-72 by serine in hog deer. These are all typical features of pseudogenes. The codons for the Cys-31 and Cys-32 residues, responsible for the covalent dimeric structure of bovine seminal ribonuclease, have become codons for Phe-31 and Cys-32, respectively, in the other three sequences (Fig. 4F). A single Cys residue is able to form a covalent dimer, but not the perfect symmetrical structure as observed in the bovine enzyme. Indirect evidence for the sheep seminal ribonuclease being a pseudogene was obtained from an analysis of the transcriptional expression of ribonuclease genes in sheep tissues. A transcript corresponding to the sheep seminal pseudogene has been detected in sheep seminal vesicles and brain, but neither the strict tissue specificity nor the high amount of the message, which has been observed in bovine seminal vesicles, is conserved in sheep tissues (Confalone et al., 1995).
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The pseudogene character of the seminal ribonuclease gene in most investigated ruminant species confronts us with a problem concerning its evolutionary history. Two scenarios are possible. One is that, after its origin by gene duplication, it was an active gene all the time up to the origin of the taxon with ox and water buffalo, which have active seminal enzymes, and that in the lines to deer, giraffe, and sheep the capacity to express active seminal ribonuclease was lost independently. The other possibility is that, after gene duplication in an ancestral ruminant, an inactive pseudogene was conserved and has been reactivated only in the ancestor of ox and water buffalo. Both scenarios are rather illogical. In our studies on nucleotide sequences encoding ribonucleases in the genome of mouse deer, we encountered a sequence that cannot yet be assigned to any of the three types of ruminant ribonucleases (Fig. 4G). The encoded sequence has all the features of an active ribonuclease, but the border region of the signal sequence and the mature sequence is Leu-Glu-Glu (with the second Glu being the N-terminal residue presented in Fig. 4G). This sequence does not conform to the consensus sequence for cleavage of signal peptides (Von Heijne, 1984), which may indicate that this gene as well is a pseudogene. In order to have an outgroup sequence for the evolutionary analysis of ruminant ribonucleases, we decided to sequence the gene of bactrian camel ribonuclease. However, to our surprise we did not find a gene encoding the pancreatic enzyme, but found one encoding a much more basic protein (Fig. 4G). This means that our original observation by Southern blotting--that only one pancreatic-type ribonuclease gene is present in camels (Breukelman et al., 1993)mis not correct and that a second ribonuclease may be present in tissues other than the pancreas. A preliminary evolutionary analysis does not group this sequence with any of the three ruminant types, which may indicate that its origin is a separate gene duplication in an ancestor of the tylopods (camels and relatives).
ACKNOWLEDGMENTS
We thank Reinhard Kleineidam, Joost Kolkman, Robert te Poele, Lourens van der Vliet, and Hayo Warmels for contributions to the described investigations, and Dr. R. N. Campagne for critically reviewing the manuscript.
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9 Pancreatic Ribonucleases CLAUDI M. CUCHILLO,*'t '1 MARIA VILANOVA,~ AND M. VICT()RIA NOGUi~S* *Departament de Bioquimica i Biologia Molecular, Facultat de Ci6ncies Universitat Autbnoma de Barcelona 08193-Bellaterra, Spain tlnstitut de Biologia Fonamental "V. Villar-Palasi" Universitat Autbnoma de Barcelona 08193-Bellaterra, Spain ~:Unitat de Bioqufmica i Biologia Molecular Departament de Biologia, Facultat de Ci~ncies Universitat de Girona 17071-Girona, Spain.
I. Introduction II. Reaction Catalyzed by Pancreatic Ribonucleases III. Specificity of Reaction and Catalytic Mechanism A. Primary Specificity B. Catalytic Mechanism C. Transition State Analogs D. Role of Different Amino Acids Involved in Catalysis IV. Substrate-Binding Subsites: Structure and Function A. Preference for Purines, B2R2 Binding Subsite B. Noncatalytic Phosphate-Binding Subsites V. Carbohydrate Moiety: Structure and Function VI. Folding/Unfolding of Ribonuclease A A. Folding/Unfolding Studies of Disulfide-Intact RNase A: Slow-Folding Reactions B. Folding/Unfolding Studies of Reduced/Native RNase A: Disulfide Bond Formation References I To whom correspondence should be addressed.
RIBONUCLEASES: STRUCTURES AND FUNCTIONS
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I. INTRODUCTION Comprehensive reviews on ribonucleases appear periodically. In the case of pancreatic ribonucleases the best known reviews are those of Richards and Wyckoff (1971), Blackburn and Moore (1982), and Eftink and Biltonen (1987), which, with some exceptions, deal mainly with ribonuclease A (RNase A), the major component found in the bovine pancreas. In addition to RNase A there is information about other pancreatic RNases, such as bovine pancreatic RNase B, human and porcine pancreatic RNases, and a number of RNases sequenced by the group of Beintema in Groningen, all of which constitute the basis of evolutionary studies (see Chapter 8 in this volume). In this chapter we shall review recent data on several aspects of the catalytic mechanism of pancreatic RNases as well as some molecular properties such as carbohydrate content and folding and unfolding pathways. The very important topic of RNase structure is covered in Chapters 10 and 11 of this volume.
II. REACTION CATALYZED BY PANCREATIC RIBONUCLEASES The depolymerization of RNA by RNase is usually described as taking place in two steps (Scheme I): in the first step there is a transphosphorylation reaction from the 5' position of one nucleotide to the 2' position of the adjacent nucleotide with the formation of a 2',3'-cyclic phosphodiester. In the second step the 2',3'-cyclic phosphodiester is hydrolyzed to a 3' nucleotide (Richards and Wyckoff, 1971; Eftink and Biltonen, 1987). + H20
RNA ~
2',3'-cyclic phosphodiesters + R-OH --'-) 3'-phosphomonoesters SCHEME I
The description of this reaction has been a source of ambiguity with respect to the mechanism, especially as to the role of the 2',3'-cyclic phosphodiesters. The classical description of the reaction as it appears in Scheme I may be interpreted as a sequential mechanism in which R - O H is the first product and the 3'-phosphomonoester is the second product of a double-displacement mechanism (Ping-Pong) similar to that of chymotrypsin, and in which the 2',3'-phosphodiester intermediate
9
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Pancreatic Ribonucleases
would be an enzyme-bound intermediate similar to the acyl-enzyme. This sequential interpretation of the RNase reaction is found, for example, in the mechanisms proposed by Hammes (1968), Roberts et al. (1969), and Rabin et al. (1970), in which not only is dissociation of the cyclic phosphodiester not considered, but the second step is necessary to recover the original state of ionization of the active site histidines to initiate a new catalytic cycle. In many articles and reviews the cyclic intermediates are considered, implicitly or explicitly, sometimes as enzyme-bound intermediates and sometimes as free intermediates, depending on the context. Cuchillo et al. (1993) showed that the 2',3'-cyclic phosphodiesters are not enzyme-bound intermediates but are true reaction products that are released into the solution. The hydrolysis of this product takes place only in aqueous solution once practically all of the poly- or oligonucleotide substrates have been used. Thus, the two steps of the reaction can be reduced to only one, as represented in Fig. 1, where R, in the forward direction, can be from a methyl group to a polynucleotide chain. In the reverse reaction, formally an alcoholysis, R can also be a hydrogen atom, and in this special case the reaction is a hydrolysis. The differences in rate between the forward transphosphorylation and hydrolysis can be explained in terms of the position of the chemical equilibrium, the con-
OR (H)
O
O"
O
O ,,
O~P---O" O ~"
H2
"
O
0 ~--- - - 0 -
I
OR'
OH
Pyr
R-OH
(H) R-OH
H2
O ~
Pyr
O ~ P--'O-
I
OR'
Fig. 1. Schematic representation of the reaction catalyzed by RNase A. Reverse transphosphorylation and hydrolysis reactions are essentially equivalent. The only difference lies in the R group of the substrates, which in the hydrolytic reaction is just H.
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Claudi M. Cuchillo et al.
centration of the different chemical species present, and, most importantly, the precise interactions of the different R - O H groups involved in the back reaction with specific enzyme subsites. The true product nature of the 2',3'-cyclic phosphodiesters formed during the reaction has also been demonstrated by the group of Raines (Thompson et aL, 1994) for a number of RNases (RNase A, a dimer and a trimer of RNase A, bovine seminal RNase, RNase T~, barnase, and RNase I). In a trapping experiment to assess the throughput of the reaction catalyzed by RNase A, they found that only 0.1% of the R N A substrate was both transphosphorylated and hydrolyzed without dissociating from the enzyme. All these results suggest that ribonucleases have evolved primarily to catalyze RNA transphosphorylation and not RNA hydrolysis. RNases are considered as endoribonucleases (EC 3.1.27.5), but they can also act on phosphodiester bonds close to the end of the substrate when this is a small molecule such as a di- or a trinucleotide. However, experiments carried out with polycytidylic acid [poly(C)] and RNase A show that there is a preference for phosphodiester linkages that are some six or seven units apart from at least one of the ends of the substrate molecule (Moussaoui et aL, 1996). This endoribonucleolytic character of the enzyme is most likely due to the special configuration of subsites that facilitate the correct and efficient binding of the substrate, as described in Section IV,B. Interestingly, deletion of the P2 subsite changes the cleavage properties of the catalyst, transforming it into an exoribonucleolytic enzyme (Moussaoui et al., 1996) (Fig. 2). Enzymes acting on polymeric biomolecules can be classified as distributive or processive. Distributive enzymes bind a polymeric substrate, catalyze a chemical reaction, and release to the solvent a polymeric product, whereas processive enzymes bind a polymeric substrate and catalyze a series of identical chemical reactions along that polymer before releasing it to the solvent (delCardayr6 and Raines, 1994). RNase A is a nonprocessive endoribonuclease that binds the bases of adjacent RNA residues in several enzymatic subsites (see Section IV,B). However, mutants of RNase A have been obtained in which Thr-45, an amino acid involved in the binding of the pyrimidine base in B~, has been transformed in either Ala or Gly. In contrast with the native enzyme the Ala-45 and Gly-45 enzymes not only cleave poly(C) and polyuridylic acid [poly(U)], but also cleave polyadenylic acid [poly(A)] very effi-
9
275
Pancreatic Ribonucleases RNAase A-DII 15
10
g RNAase A-K7QR10Q
r
o L -
30-
Q.
< 20
10
!
!
!
!
!
!
|
|
2
3
4
5
6
7
8
9
Oligonucleotide Size Fig. 2. Patterns of (Cp),C>p (n from 0 to 8) oligonucleotide formation from poly(C) cleavage by RNase A (I-l), derivative II ([]), and K7Q plus R10Q RNase A mutant ([~). In each case the peak area was determined from product separation by reversed-phase HPLC. Due to the different enzyme activities, comparisons have been established using as reference the same undigested poly(C) fraction (30%) (Moussaoui et al., 1996).
ciently. B o t h m u t a n t e n z y m e s cleave p o l y ( A ) processively w h e r e a s cleavage of poly(C) and p o l y ( U ) r e m a i n s distributive ( d e l C a r d a y r 6 and R a i n e s , 1994).
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III. S P E C I F I C I T Y O F R E A C T I O N
AND CATALYTIC MECHANISM
A. Primary Specificity As shown in Fig. 3, the base on the nucleotide at the 3' side of the phosphodiester bond broken by RNase A must be a pyrimidine. This primary specificity of the enzyme was found as early as in 1938 by Schmidt and Levene (cited in Rushizky et al., 1961), although more careful determinations demonstrated that RNase A is also capable of a transphosphorylating activity toward poly(A) (Beers, 1960; Imura et aL, 1965) and polyformycin, an unusual purine (Ward et aL, 1969). In this latter case the activity of the enzyme is much higher than it is toward
B,
\2
-P~O~CH
",'%
Thr-45 Phe. 120 Ser-123
~176
B2 Asn-71 ..~-,~_. Glu-111
I p,.o,,,j
r1.-.yI ~pZ*
"~ HJ$-12
N ._,O_.=CHa
~
i
N -
"
B3
o OH
Lys- 1
.N-~N
)(/""-" 0 - ~ " ell20Lys. 7
Arg- 10
R:='~,,= 0 " "~'11
OH
X
Fig. 3. Schematic representation of the active center cleft in the RNase A-substrate complex. Base (B), ribose (R), and phosphate (p) binding subsites are indicated. The phosphate group of the phosphodiester bond hydrolyzed by the enzyme binds to p~; B~ is specific for pyrimidines and B: shows a preference for purines. Amino acids located near each subsite are indicated (Par6s et al., 1980).
9
Pancreatic Ribonucleases
277
the normal purines and is comparable to that toward the pyrimidines. However, the hydrolysis of the corresponding 2',3'-cyclic nucleotides (of adenine and formycin) is extremely slow. In the case of polyformycin it was shown that the nucleosides in the polymer have the unusual syn conformation, a fact that was critical for an understanding of the data. The nucleoside can "fit" the pyrimidine-binding site only when in the syn conformation. Parallel to this primary specificity, which depends on the interaction between the pyrimidine base and the subsite B1 (see later), the base composition of adjacent nucleotides has a profound influence on the kcat/Km. In this way oligonucleotides containing one or more purines and the terminal pyrmidine are released earlier than the pyrimidine nucleotides (Rushizky et al., 1961). Similar results were found by Witzel and Barnard (1962) using dinucleoside monophosphates as substrates. The rate was very dependent on the base of the nucleoside at the 5' side of the phosphodiester bond broken, the order being A > G > C > U. This "secondary" specificity arises from the interaction of the substrate at subsites other than B1 and we shall deal with this in Section IV,B. Here we only analyze some of the characteristics of the B~ subsite. Structural and phylogenetic data suggest that the specificity of the B1 subsite is mediated by the side chains of Thro45 and Pheol20 (Wlodawer and Sj01in, 1983; Santoro et al., 1993; delCardayr6 and Raines, 1994). The hydroxyethyl side chain of Thro45 forms hydrogen bonds with a pyrimidine base (U or C) and excludes sterically the purine bases (A and G). The benzylic side chain of Phe-120 makes van der Waals contact with a pyrimidine base and with the side chain of Thro45. It is interesting to note that replacement of Phe-120 by a Tyr residue in a semisynthetic RNase results in an enzyme that is fully active with either RNA or cytidine 2',3'-cyclic phosphate (C>p) as substrate, but its activity toward uridine 2',3'ocyclic phosphate (U>p) is twice that of the complex in which the original Phe-120 is conserved. The activity toward U > p of giraffe RNase (which contains a tyrosine in position 120) is also increased (Hodges and Merrifield, 1974). The side chain of Ser-123 shows persistent hydrogen bonding interaction with the uridine base in active site dynamics studies of RNase A with the uridine-vanadate complex (Bringer et al., 1985; Seshadri et al., 1993), whereas no bond between Ser-123 and the uridine base is seen in the X-ray structure. In addition, the molecular dynamics simulation shows that hydrogen bonds between enzyme and bound water molecules in the free enzyme replace the corresponding hydrogen bonds between enzyme and substrate. These waters and those
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that interact with the pyrimidine-site residues Thr-45 and Ser-123 help to preserve the protein structure in the optimal arrangement for binding. The presence of such waters in the active site has the consequence that water extrusion and its inverse must be intimately correlated with substrate binding and product release. Computer modeling studies have shown that in the case of pyrimidine mononucleotides, Val-43, Asn-44, and Thr-45 may be important for the proper positioning of the base and that the segments vala3-Thr 45 and Hisll9-Asp 121 together form a groove in which the base can slide for proper positioning. Ribose and phosphate then adjus~ themselves with the side chains of Asn-44, His-ll9, and Phe120 in such a way that the phosphate moiety is placed in favorable stereochemical position for optimal binding of these ligands (Seshadri et aL, 1993). The specificity of the B1 subsite for pyrimidine nucleotides is not absolutely strict. Ards et al. (1981) showed that the pK values of His-12 and His-ll9 and the shift of an aromatic resonance of the native enzyme found on interaction with some purine nucleotides can be interpreted by postulating that the interaction of 5'-AMP, 5 '-GMP, and 5'-IMP takes place not only in the purine-binding site B2R2p~ but also in the primary pyrimidine-binding site B~R~p0. Later, Aguilar et al. (1991) corroborated this finding by means of X-ray crystallography (they called it "retrobinding"). Finally, delCardayr6 and Raines (1994), by mutating Thr-45 of the native enzyme to either Ala or Gly, found that the mutated enzymes cleave poly(A), poly(C), and poly(U) efficiently and with 103to 10S-fold increases in purine/pyrimidine specificity.
B. Catalytic Mechanism The first proposed mechanism for the reaction catalyzed by bovine pancreatic RNase A was formulated in 1961 by the group of Rabin (Findlay et al., 1961). This mechanism postulates that two histidines participate in the process (Fig. 4). In the forward transphosphorylation there is an intramolecular attack of the vicinal 2'-hydroxyl on the phosphorus atom facilitated by His-12 acting as a general base catalyst. His119 acts as a general acid catalyst and facilitates the cleavage of the phosphorus-5'-O bond. The attack by the 2 ' - 0 is "in-line," with the 2'-0 and 5 ' - 0 assuming apical positions in a pentacoordinate trigonal bipyramidal transition state (Usher et al., 1970). It is not yet clear whether
9
279
Pancreatic Ribonucleases
o o.
oi
f I
-
~
HO
OH
;
.
v
"~HO
(~)
OH ~
B~ #
P
~--..~.P.,
It + ) ,
....
119
HN
Fig. 4. Chemical mechanism for the RNase A catalytic reaction. His-12 and His-119 are involved in an acid-base bifunctional mechanism. Adapted from Roberts et al. (1969).
His-12 and His-119 act in a concerted fashion. For the back transphosphorylation, including hydrolysis, the role of the histidines is reversed. With some modifications, for example those of Roberts et al. (1969), this mechanism has stood well the vast amount of new data that have been produced since its original formulation. Of the other alternative mechanisms that have appeared, the most interesting ones are those of Witzel (1963) and of Breslow (Anslyn and Breslow, 1989). In the mechanism proposed by Witzel a key role is ascribed to the keto oxygen in the 2 position of the base; the two histidines are also necessary, but the only interaction with the substrate is through His-ll9, and His-12 is assumed to be hydrogen bonded to His-119. To reconcile their mechanism with the data of Usher et al. (1970), Witzel and associates (Rtibsamen et al., 1974) invoked a turnstile rotation to permit inversion of the absolue configuration as a consequence of an adjacent mechanism. In the Witzel mechanism the two active site histidines must lie close together; however, in the crystal structures of both
I
280
Claudi M. Cuchilio
et al.
the RNase A and the RNase S normally obtained in the presence of phosphate, sulfate, substrate analogs, or competitive inhibitors, the two histidines are not close enough for a hydrogen bond to exist between them. It could be argued that the inhibitors or analogs may interact in a manner different from the substrate and transition state, but Campbell and Petsko (1987) resolved the structure of RNase A freed from sulfate at 1.5 A and found that the two histidines of the active site had moved very little--the distances between the e-nitrogen of His-12 and the 6nitrogen of His-119 increased from 6.05 to 6.26 A m a n d thus the active site geometry is in disagreement with the mechanism of Witzel and coworkers. Similar results concerning the histidines in the active site were obtained by Wlodawer et al. (1988) with phosphate-flee RNase A refined at 1.26/k. They found only a small tilting of His-119, which seemed to favor a better geometry for hydrogen bonding to Asp-121. Molecular dynamics studies have also shown that the two histidines never get closer to each other than 5 A. (Brtinger et al., 1985). The group of Breslow (Anslyn and Breslow, 1989; Breslow and Xu, 1993) proposed a different mechanism in which attack by the 2'-hydroxyl group is facilitated by protonation of a nonbridging phosphoryl oxygen atom, instead of the leaving-group oxygen atom, to give a "triester-like" mechanism of catalysis with the formation of a phosphorane monoanion intermediate. This mechanism has been questioned (Menger, 1991; Haim, 1992) and theoretical calculations (Haydock et al., 1990; Lim and Tole, 1992) have shown that at least some of the details of the mechanism do not fit with the predicted models. Herschag (1994) found that thio effects (rate effects on substitution of nonbridging phosphoryl oxygen atoms by sulfur) are inconsistent with predictions based on the mechanism of Breslow but that they are in agreement with the classical mechanism of Rabin.
C. Transition State A n a l o g s
Information on the catalytic mechanism can be obtained by the use of transition state analogs. Both the classic acid-base and the triesterlike mechanisms involve a pentacoordinate phosphorus intermediate. Lindquist et al. (1973) proposed that complexes of uridine with oxovanadium(IV) ion and vanadium(V) ion could be transition state analogs in the RNase A reaction. At pH 7.0 and 25~ the dissociation constants
9
Pancreatic Ribonucleases
281
of the complexes were 1.0 • 10 -5 and 1.2 • 10 -5 M, respectively, a strong binding that could be explained by the resemblance of structure of the complexes to the structure of the substrate portion of the transition state for the RNase-catalyzed hydrolysis of U > p . The structure and properties of the complexes of RNase A with either vanadate or uridine vanadate adducts have been studied by a variety of techniques, for example, X-ray crystallography (Wlodawer et al., 1983), N M R and neutron diffraction (Borah et aL, 1985), N M R (Veenstra and Lee, 1994), molecular dynamics (Brtinger et aL, 1985), electron-transfer spectra (Ray and Post, 1990), or study of the electrostatic potential (Krauss and Basch, 1992). In this latter work some doubts about the suitability of vanadate as a transition state analog are raised because the five-coordinate vanadate inhibitor is found electronically to have both similarities to and differences from the phosphorane, and the observed Lys-41 and His-12 orientations are not found relevant to the RNase A mechanism. According to these authors, because the electrostatic potential at H-bonding distances of O - ( V ) for both monoanion and dianion vanadates are considerably smaller than expected for O - ( P ) in the transition state phosphorane, the flexible Lys-41 side chain can search for and, apparently, bind to the more electronegative axial O - ( H ) site. The position of this residue is, therefore, not probing the electronic properties of the transition state and is irrelevant to the mechanism, but is an indication of the flexibility of this side chain.
D. Role of Different Amino Acids Involved in Catalysis
Here we shall comment only on the residues of bovine pancreatic RNase A that are known to play a role in the catalytic mechanism. Comments on other residues that are important in the overall binding of the substrate molecule will be dealt with in Sections III,A and IV,B. 1. Histidine-12
As stated, His-12 acts as a base in the transphosphorylation reaction, abstracting a proton from the 2'-OH of the ribose, thus favoring the attack on the phosphorus atom to give the pentacoordinate transition state. However, in its absence, Asn-44 can fulfill this role (Deakyne and Allen, 1979). In the back transphosphorylation (hydrolysis) reaction His-
282
Ciaudi M. Cuchillo et al.
12 acts as an acid favoring the attack of water on phosphorus and, obviously, cannot be replaced by Asn-44.
2. Histidine-ll9 His-119 acts as an acid in the transphosphorylation, partially protonating the leaving group (5'-OR) weakening the phosphorus-5'-O bond and facilitating an in-line addition of 2 ' - 0 (Deakyne and Allen, 1979). In the mechanism of Anslyn and Breslow (1989) His-119 acts also as an acid in the transphosphorylation step, but in this case it protonates a nonbridging phosphoryl oxygen atom. It is known from the earlier studies on the X-ray structure of RNase A that His-119 can occupy several positions (Richards and Wykoff, 1971). Several crystallographic studies (see, for instance de Mel et al., 1994) have shown that there are two predominant conformations, called A and B [or "down" and "up" according to Kim et al. (1992)]. Modeling studies (de Mel et aL, 1992) have shown that His-ll9 is positioned equally well to participate in both transphosphorylation and hydrolytic reactions by the enzyme, whether it is in position A or B.
3. Aspartate-121 Asp-121 is another residue that has been implicated in the catalytic mechanism because only 0.5-4% activity toward various substrates is retained when the four terminal residues of the enzyme (121-124) are removed by proteolysis (Lin, 1970). Theoretical studies by Deakyne and Allen (1979) showed that although a hydrogen bond between His-ll9 and Asp-121 had been proposed, a strong interaction between these two residues was unlikely and Asp-121 had little importance in the transphosphorylation reaction other than electrostatic stabilization of the positive charge on His-ll9. Taken together with crystallographic data, their results suggested that a possible role for Asp-121 would be to position the adenine ring (in B2) and this ring in turn would align His-119 in site IV (Richards and Wyckoff, 1971) via base stacking. The NMR results of Arfis et al. (1982) are in agreement with this interpretation. The conservative mutation of Asp-121 to Glu-121 results in an enzyme species that retains 17% of the activity of the native enzyme (Trautwein et al., 1991), whereas substitution of Asp-121 by Asn-121 in a semisynthetic RNase produces an analog with only 2.7% activity (Cederholm et al., 1991).
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Pancreatic Ribonucleases
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4. L y s i n e - 4 1
The chemical modification of Lys-41 (Hirs, 1962) yields a derivative with less than 0.4% activity with respect to the native enzyme. Crystallographic studies have shown that when RNase A is bound to the vanadate complex of uridine, Lys-41 appears to interact with the negatively charged vanadate oxygen. The average temperature factor for Lys-41 is lowered from a quite high value (B = 20 A2) in the enzyme crystallized in the presence of phosphate to 8 ,~2 in the presence of the transition state analog (Wlodawer et al., 1983). The stabilization of Lys-41 in the transition state complex reinforces the assumption that this residue plays a direct role in the catalytic mechanism. It is believed that the rigid Lys41 stabilizes the transition state, whereas the flexible side chain allows its formation (as well as decomposition) (Wlodawer, 1985). Site-directed mutagenesis in which Lys-41 was replaced by Arg resuited in a mutant enzyme with a kcat/gm of only 1.9% that of the native enzyme using UpA as substrate (Trautwein et al., 1991). It is known that there is a large conformational change undergone by the loop containing Lys-41 when substrate binds (Douzou and Petsko, 1984), a conformational change that is believed to deliver the side chain of Lys-41, but not Arg-41, to the position where its protonated group can best interact with the negatively charged pentacoordinate phosphate group in the transition state (Trautwein et aL, 1991).
5. L ysine- 7, L ysine-66, and Water Networks Several positively charged groups are located near the active site of RNase A. Some of them may be involved in guiding and/or binding the substrate (Matthew and Richards, 1982). Molecular dynamics simulations (Brtinger et al., 1985) showed that these residues, including Lys-7, Lys-41, Lys-66, Arg-39, and the doubly protonated His-119, are stabilized in the absence of ligands by well-defined water networks. The bridging waters are able to stabilize the otherwise very unfavorable configuration of positive groups. It is seen that both Lys-7 and Lys-66 are correlated significantly with the uridine vanadate ligand, although the side chain nitrogens of the two lysines are more than 5 A removed in both cases. The dynamic coupling appears to be mediated by an intervening water network. Such correlated fluctuations between distant lysine residues (indeed, neither of these residues belong to the BIRlp~, i.e., the "catalytic site") and the phosphate group of the substrate may indicate a role for
284
Claudi M. Cuchillo et
al.
these residues in stabilizing the transition state. Support for this comes from experimental evidence using a mutated enzyme in which Lys-7, a residue of the P2 subsite, has been replaced by Gin. In this mutant enzyme there is a significant decrease in the rate of catalysis of C>p, a substrate that interacts only in the catalytic site, without apparent change in Km (Boix et aL, 1994).
IV. SUBSTRATE-BINDING SUBSITES: STRUCTURE AND FUNCTION A. Preference for Purines, B2R2 Binding Subsite
Kinetic studies using RNA as substrate indicated that, besides the primary specificity for pyrimidines, RNase A shows preference for phosphodiester bonds that have a purine nucleotide in the 5' position (Rushizky et al., 1961). The existence of a secondary binding site on the 5' side of the active center, which would prefer purines, was hypothesized by Deavin et al. (1968). Kinetic studies of C>p hydrolysis by RNase A in the presence of 5'-AMP indicated that the formation of a ternary complex produces an increase in the enzyme activity (Haffner and Wang, 1973). X-Ray crystallography of complexes between RNase A and dinucleotide substrate analogs such as UpcA [uridylyl-3',5'-(5'-deoxy-5'methylene)adenosine] (Richards and Wyckoff, 1973), 2',5'-CpA (cytidylyl-2',5'-adenosine) (Wodak et al., 1977), and 2'-FdUpA (2'd e o x y - 2 ' - f l u o r o u r i d y l y l - 3 ' , 5 ' - a d e n o s i n e ) (Pavlovsky et al., 1978) showed the relative positions of the pyrimidine nucleoside moiety (B~RI), the phosphate group (pl), and the purine nucleoside moiety (B2R2) (Fig. 3). Amino acid residues (Gin-69, Asn-71, G l u - l l l , and His-ll9 were reported to belong to the purine-binding region (B2R2). These interactions have been revised in complexes solved by X-ray crystallography at high resolution between RNase A and the ligands d(CpA) (Zegers et al., 1994) and the oligonucleotide d(ApTpApApG) (Fontecilla-Camps et al., 1994). In both cases the purine-binding site has multiple conformations, although there is a clear stacking interaction between His-119 and the adenine base in B2. The purine base also hydrogen bonds to Asn-71. A contact between Gln-69 and the adenine base in B3 was also observed but no contacts are seen with the carboxyl group of Glu-lll, although this residue is near this region.
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Pancreatic Ribonucleases
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Amino acid residues involved in the purine-binding site have also been analyzed by site-directed mutagenesis and kinetic studies: Asn-71 seems to be involved in productive interactions when CpA is used as substrate, but no effect due to either Gin-69 or Glu-111 is observed. However, Glu-111 appears to be involved in catalysis when CpG is used as substrate (Tarragona-Fiol et aL, 1993).
B. Noncatalytic Phosphate-Binding Subsites The existence of noncatalytic phosphate-binding subsites additional to the active site in the RNase A has been demonstrated by both kinetic and structural studies. Steady-state kinetic studies with U p A p A and U p A p G as substrates show an increase in the kcat values with respect to UpA (Irie et aL, 1984a). The breakdown of oligouridylic acids of increasing chain length indicates that the catalytic efficiency increases with the length of the substrate in the case of di- or trinucleotides, although no further increase is observed with both tetra- and pentanucleotides. These results suggest that the number of subsites important for catalysis corresponds to three nucleotides (Irie et al., 1984b). On the other hand, the Vmax value obtained with poly(U) as substrate is in all cases 3 to 20 times higher than the values obtained with the oligouridylic acids (Irie et aL, 1984b), and thus additional binding subsites must contribute to the catalysis of long polynucleotides. The products of poly(C) cleavage as a function of digestion time have been analyzed by HPLC. Under the initial conditions RNase A prefers the binding and cleavage of long substrates, but as the reaction proceeds an accumulation of oligonucleotides having six to seven residues takes place (Fig. 2) and, eventually, the formation of mononucleotide product is observed (Moussaoui et al., 1996). A model of a complex between the RNase A and the oligoribonucleotide p A p U p A p A p G was proposed from modeling studies (de Llorens et al., 1989) based on electrostatic interactions between phosphate groups of the ligand and basic amino acid residues located at the surface of the protein, and the crystal structure of the complex between RNase A and d(ApTpApApG) is also known (Fontecilla-Camps et al., 1994). In both cases, a good complementarity between negative phosphate groups of the substrate and positive amino acid residues of the enzyme was observed.
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Claudi M. Cuchillo et al.
A detailed description of phosphate-binding subsites adjacent to the active center has been published (Nogu6s et al., 1995). As seen in Fig. 3, P0 is the phosphate-binding subsite at the 5' side of the scissile phosphodiester bond in p~, whereas P2 is the corresponding phosphate subsite at the 3' side of p~. The involvement of the P0 subsite in the binding of RNA was deduced from studies on the binding of the nucleoside bisphosphate, uridine 2'(3'),5'-bisphosphate, which has an association constant with RNase A 14-times stronger than that of the nucleoside monophosphate 2'(3')-UMP (isomers mixture) (Sawada and Irie, 1969). On the other hand, a five-fold increase in the catalytic efficiency of RNase A in the hydrolysis of pU>p (uridine 5'-phosphate, 2',3'-cyclic phosphate) with respect to U>p was found (Li and Walz, 1974). X-Ray crystallography of RNase A - A p C complexes (Mitsui et al., 1978) and model building studies (de Llorens et al., 1989) indicated that Lys-66 was a candidate for P0. However, in the crystallographic studies of the RNase A-d(ApTpApApG) complex this interaction is not observed, although the authors point out that this region of the protein can adopt different orientations, and, in solution, a conformation in which Lys-66 interacts with the phosphate group is not excluded (Fontecilla-Camps et al., 1994). The phosphate-binding subsite P2 was postulated from the reaction between RNase A and the nucleotide 6-chloropurine riboside 5'-monophosphate, which yields a major derivative, named derivative II, in which the nucleotide is attached through the purine ring to the c~-amino group of Lys-1 (Par6s et al., 1980, 1991). The specificity of the reaction is a consequence of the binding of the phosphate group to this specific region of the enzyme. This fact was corroborated by NMR studies of derivative II, RNase A, and their complexes with nucleotides (Art, s et al., 1981, 1982; Alonso et al., 1988, 1989). Kinetic studies of C>p hydrolysis in the presence of pAp and 5'-AMP as inhibitors indicated a higher inhibitory effect of pAp with respect to 5'-AMP as a consequence of the additional binding of the 3'-phosphate group of pAp to the P2 subsite (Irie et al., 1984a). Chemical modification of RNase A and derivative II indicated that Lys-7 and Arg-10 are the amino acid residues constitutive of this phosphate-binding subsite (Richardson et al., 1990). Crystallographic studies confirmed the role of Lys-7, although no interaction with Arg10 was observed. The crystal structure of the RNase A - d ( A p T A p A p G ) complex unambiguously demonstrated the involvement of Lys-7 in this subsite (Fontecilla-Camps et al., 1994). The X-ray structure of derivative II demonstrated that the nucleotide label is located at the N-terminal region of the protein occupying the P2, B3, and R3 subsites (Fig. 3), with Lys-7 involved in electrostatic interaction with the phosphate group (Boqu6 et al., 1994).
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The contribution of Lys-7 and Arg-10 to the catalytic process was also analyzed by site-directed mutagenesis (Boix et al., 1994). Kinetic studies of K7Q, R10Q, and the double mutant K7Q plus R10Q confirmed the role of these amino acid residues in the establishment of electrostatic interactions with the substrate and confirmed that an operative P2 subsite was necessary for the synthesis of C p C > p using C > p as substrate. As pointed out in Section Ill,D, an indirect role in the catalytic mechanism has also been proposed. Deletion of this subsite also modifies the RNAse A endonuclease activity, which is apparently changed to an exonuclease activity (Fig. 2) (Moussaoui et aL, 1996). The present knowledge on the structure and location of the phosphatebinding subsites, together with the kinetic properties of the enzyme, demonstrate that the better catalytic efficiency seen with high molecular mass substrates is due to multiple cooperative binding of the substrate to the enzyme. From these results a model of the cleavage of an RNA chain by RNase A has been proposed (Fig. 5) (Par6s et al., 1991).
V. C A R B O H Y D R A T E MOIETY: S T R U C T U R E AND F U N C T I O N
Amino acid sequence analysis of many pancreatic RNases reveals the presence of carbohydrate attachment sites corresponding to the Asn-XThr/Ser sequence. The attachment of carbohydrate takes place through an N-acetylglucosamine residue in N-glycosidic linkage with an Asn residue. Although the carbohydrate attachment sites are located at exposed regions of the RNase molecules, not all potential glycosylation sites are coupled to carbohydrate and there are notable differences between species in the complexity of the carbohydrate chains. The carbohydrate moiety of pancreatic RNases is not required for the enzymatic activity. The function is unknown, although a resistance to absorption by the gut and to degradation by proteases has been proposed. Both processes increase the half-life of pancreatic RNases in the digestive tract and contribute to the RNA digestion, this latter fact being specially important in ruminants and other species having cecal digestion wherein the RNA of the microflora is an important source of nitrogen and phosphorus (Blackburn and Moore, 1982). It is interesting to note that the activity of pancreatic-type RNases on double-stranded RNA is directly dependent on the extent of glycosylation (Carsana et al., 1981). Bovine pancreatic RNase can be separated into several fractions that differ in the glycosylation pattern. The RNase A fraction consists of a
shorter fragments
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single polypeptide chain whereas the RNase B fraction has, in addition to the same polypeptide chain, an oligosaccharide chain bound to the Asn-34 residue. The specificity and kinetic properties of the reaction catalyzed by RNase B appear to be identical to those of RNase A. Early studies demonstrated that in addition to RNase B there are other glycosylated fractions (RNases C and D) that differ only in the oligosaccharide composition (Plummer, 1968). The carbohydrate chain of the major glycosylated fraction, RNase B, is composed of two N-acetylglucosamine residues and five to eight mannose groups with two 1,3 and 1,6 branch points. RNases C and D are less abundant forms and have a more complex oligosaccharide composition (Eftink and Biltonen, 1987). A sequence for the carbohydrate content of RNase B was proposed by Liang et al. 1980), but Fu et al. (1994) have reported a much more detailed structural characterization in which a high degree of heterogeneity is found with respect to the carbohydrate chains. Nine different oligosaccharide chains containing mannose and N-acetylglucosamine are described. The crystal structure of RNase B has been obtained at 2.5 A resolution by Williams et al. (1987). The oligosaccharide moiety does not seem to make any contact with the protein except at the covalent attachment point, appears to have no real influence on the conformation of the protein, extends into the solvent, and is for the most part disordered. The polypeptide chains of the RNase A and B forms have essentially identical structures. Beintema et al. (1988) have described differences in the glycosylation pattern of human pancreatic-type RNases even if the polypeptide primary structures of the enzyme from different tissues and urine are identi-
Fig. 5. Model of the cleavage of an RNA chain by RNase A that explains the preference of the enzyme for long polynucleotide substrates. This model is based on the cooperative binding between the multiple protein subsites and the phosphates of the RNA that contribute to the catalytic efficiency. The RNA substrate, a long chain, binds strongly to the subsites of the protein; when cleavage of the chain takes place in the transphosphorylation reaction, the cooperativity is partially lost in the resulting shorter fragments and a new intact chain will then displace the fragments. When the longer substrate molecules have already been cleaved, shorter fragments bind to the enzyme and oligonucleotides containing six to seven residues are accumulated according to the occupancy of the protein subsites. Oligonucleotides are eventually cleaved and then the hydrolytic step takes place when most of the 3',5'-phosphodiester bonds are already broken. Reproduced with permission from Nogu6s et al. (1995).
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cal. Breukelman et al. (1993) indicated that all human pancreatic-type RNases are products of the same gene and that the amino acid sequence derived from the DNA sequence is identical to the previously determined protein sequence of the pancreatic enzyme (Haugg and Schein, 1992). Human pancreatic-type RNase has three Asn-X-Thr/Ser sites (Asn-34, Asn-76, and Asn-88) that present different glycosylation patterns depending on the organ or fluid analyzed. Pancreatic-type RNase from urine has each of the three sites glycosylated with complex-type oligosaccharide chains (Beintema et al., 1988), whereas RNase purified from pancreas has carbohydrates attached to Asn-34 and is only partially glycosylated at the other two positions. A significant degree of microheterogeneity in the carbohydrate moiety is observed (Rib6 et al., 1994). Differences are also observed in RNases from pancreatic juice (Thomas et al., 1984), seminal plasma (De Prisco et al., 1984), and kidney (Mizuta et al., 1990). Neither the origin of the urine enzyme nor the role of the different glycosylation patterns is known.
VI. F O L D I N G / U N F O L D I N G OF R I B O N U C L E A S E A
RNase A has been a classic in the folding/unfolding studies of proteins (Blackburn and Moore, 1982; Kim and Baldwin, 1982, 1990; Matthews, 1993). This enzyme has some structural characteristics that make it a very interesting model for these kinds of studies. It is a small globular protein with a single domain that is stabilized by four disulfide bonds and has two X-Pro peptide bonds in the cis conformation. What is known about slow-folding reactions and about the mechanistic details of disulfide bond formation derives in great part from studies of this small protein, and, thus, these are the topics covered in this section.
A. Folding/Unfolding Studies of Disulfide-Intact RNase A: Slow-Folding Reactions
RNase A was the first protein found to have both slow-folding (Us) and fast-folding (UF) forms of the unfolded protein, showing multiphasic refolding kinetics, with fast and slow reactions arising from proline cistrans isomerization (Blackburn and Moore, 1982; Kim and Baldwin,
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1990; Nail, 1994). Unfolded RNase A refolds in three different kinetic phases: two major slow-refolding phases (60-70% and 10-20% of total refolding amplitude) (Schmid and Blaschek, 1981; Schmid, 1982, 1983) and a fast-folding reaction that makes up the remaining (18%) of the total amplitude for refolding. Under some conditions a third minor (7%) slow phase is also detected (Lin and Brandts, 1983b). These three major kinetic phases are believed to arise from at least three different unfolded species, named UF, US,I, and US,II , which make up ~20, ---15, and -~65% of the total unfolded species, respectively (Schmid, 1983). An additional minor Us species has also been reported (Lin and Brandts, 1987). Each one of these species is proposed to fold independently to the native conformation, which is considered to be a single well-defined species. The UF species has been generally regarded as an unfolded state in which all the prolines are in the native conformation. This species folds in a fast phase (Lin and Brandts, 1983b) in a reaction that does not involve imide isomerization, whereas US, I and US,II species are converted to intermediate species and eventually pass through an isomerization step. Very little is known about the nature and refolding pathway of the Usj species because of its very small amplitude and its very slow refolding rate. Mui et al. (1985) postulated the existence of an intermediate in the refolding pathway (designated I') on the basis of a kinetic analysis of the data present in the literature, although it has not been observed experimentally. Refolding of the Us,~l species has been characterized best. In its refolding pathway, two intermediates are found. An early hydrogen-bonded intermediate (I1) is formed rapidly (Udgaonkar and Baldwin, 1988, 1990) whereas a folded nativelike intermediate (IN) accumulates late. It seems that in this pathway, folding precedes isomerization (i.e., both intermediates have at least one Pro in nonnative conformation) (Cook et al., 1979; Schmid and Blaschek, 1981; Schmid, 1983), although this conclusion has been disputed by Lin and Brandts (1988). The nativelike intermediate IN has been well studied (Cook et al., 1979; Schmid and Blaschek, 1981; Schmid, 1983; Schmid et al., 1986) and shares most of the properties of the native protein. The I~ species has the optical properties of the unfolded protein but, as judged by amide hydrogendeuterium ( H - D ) exchange experiments, is highly stable and has much of the H-bonded structure of the native protein, especially in the/3-sheet region (Udgaonkar and Baldwin, 1990). In native RNase A, Pro-93 and Pro-ll4 are in the cis conformation (Richards and Wyckoff, 1971; Wlodaver and Sj61in, 1983) and thus they
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are the logical candidates for producing the slow-folding species. Cook et al. (1979) proposed that at least Pro-93 is in a nonnative trans confor-
mation in the Us,ii species. This is now generally accepted on the basis of the refolding studies by Schmid et al. (Krebs et al., 1983; Schmid et al., 1986; Lang et al., 1986; Lang and Schmid, 1990) on homologous RNases in which Pro-93 is conserved. Several studies have been carried out to determine whether isomerization of Pro-93 and/or Pro-114 accounts for the Us species of unfolded RNase A, but contradictory results have been obtained (Lin and Brandts 1983a, 1984; Adler and Scheraga, 1990; Biringer and Puntambekar, 1991). The more recent approach to solve which of the proline residues is responsible for the slow-folding species of RNase A uses recombinant RNase A proteins that have either one or both proline residues substituted, i.e., single and double mutants (Schultz and Baldwin, 1992; Schultz et al., 1992). The results with the double mutant (P93A, P114G) are clear and indicate that the replacement of both proline residues is sufficient to eliminate the major Us species (Usa~). Kinetic analysis of the folding/unfolding of two single mutant forms of RNase A with replacement of Pro-ll4 ( P l l 4 G and P l l 4 A ) shows that the slow-refolding species (Us) is generated in the unfolded state of these mutant proteins, but this species results in a maximum of only 50-60% of the total unfolded species, compared to the about 80% Us species found for native RNase A. This result is consistent with the idea that Pro-93 and Pro-114 both generate slowrefolding species and that removal of Pro-114 decreases the total amount of Us species, i.e., only the species generated by cis to transisomerization of Pro-93 remain. The folding/unfolding kinetics of Pro-93 single mutants of RNase A (P93A and P93S) are complex, with many aspects remaining unexplained. Thus, the main conclusion of this work is that the two proline residues are responsible for the major Us species of RNase A, but it is not yet known how they account for the individual species, Us,~ and Us.~. The generally accepted refolding model for RNase A is as follows: UF ~ N, 20% of the amplitude, fast Us.i~ ---> I~ ~ IN ~ N, 60-70% of the amplitude, slow Us.l ~ (I') ---> N, 10-20% of the amplitude, very slow
Intermediate I' has not been observed experimentally; the UF species is supposed to have both proline residues, 93 and 114, in the native cis
9
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conformation and Us,i, US,II, 11, and IN are supposed to have at least one proline residue (Pro-93) in nonnative conformation. On the other hand, the generally accepted unfolding model for RNase A is fast N
slow
> U F ~
US,I,II
where the fast phase is postulated to be a conformational unfolding step and the slow phase is proposed to be a proline isomerization step. Recent studies have complicated these models. Houry et al. (1994) propose for the RNase A unfolding pathway a new model that is based on the presence of two independent isomerization processes: one at Pro93 and the other at Pro-ll4. This model has been proposed to account for the existence of a new unfolded species (Uvf) that gives rise to a new very fast folding phase, not detected in earlier works. These refolding studies were carried out at very low pH and at various guanidinium chloride concentrations, conditions that slow down the refolding phase of the Us species. By analysis of the kinetic data based on this new model, the authors propose that both prolines are in their native conformation in the Uvf species, that Pro-114 is in the nonnative conformation in the UF species, and that at least Pro-93 is in the nonnative conformation in the slow-folding species, Us. which actually consist of (at least) two species: Us,s, with Pro-93 in a nonnative (trans) conformation, and Us.~, with both Pro-93 and Pro-ll4 in nonnative (trans) conformation. It is not clear whether these two slow-folding species correspond to the Us.~ and Us,ii species proposed in earlier works. Although both isomerizations about Pro-93 and Pro-ll4 are proposed to give rise to the unfolded species, only the cis-trans isomerization of Pro-93 is suggested to be rate limiting in the refolding pathways of RNase A. The study of the kinetics of the refolding of Uvf species (refolding of RNase A in the absence of complicating effects of cis/trans-proline isomerization) indicates that the pathway proceeds through the formation of a hydrophobically collapsed intermediate with properties similar to those of equilibrium molten globules (Houry et al., 1995). Furthermore, the authors suggest that the rate-limiting transition states on the unfolding and refolding pathways are substantially different, with the refolding transition state having nonnativelike properties. Several works suggested that nonrandom structures exist in the thermally denatured state of RNase A (Seshadri et al., 1994), although a
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stable hydrogen-bonded structure was not detected by amide proton protection experiments (Robertson and Baldwin, 1991). In contrast, Zhang et al. (1995) report that the pressure-denatured state of RNase A contains some secondary structure and displays the characteristics of a molten globule. Kiefhaber et al. (1995) have also detected a molten globule in the guanidinium chloride unfolding of RNase A. This species is proposed to be an intermediate in the unfolding process of RNase A that precedes the rate-limiting step. It seems also to be a "dry" molten globule (water has not yet penetrated the hydrophobic core of RNase A, although most side chains have become free to rotate) and not an equilibrium ("wet") molten globule species.
B. Folding/Unfolding Studies of Reduced/Native RNase A: Disulfide Bond Formation
The oxidative folding of RNase A is a very complex process with a large number of intermediates. The regeneration of RNase A by small molecular mass thiol redox couples has been studied in a number of laboratories (Creighton, 1977,1979; Konishi et al., 1981, 1982a-c; Wearne and Creighton, 1988), but the results obtained are still the subject of controversy and have led to the proposal of two different mechanisms for the regeneration of native RNase A from a fully reduced-unfolded species: (1) In the first mechanism multiple pathways are adopted, with each pathway having a different rate-determining step. The relative amount of native protein generated through each pathway depends on the solution conditions (Konishi et al., 1982a-c; Scheraga et al., 1987). (2) In the second mechanism there is a single folding pathway with the rate-determining step being the formation of the final disulfide bond (Creighton, 1977, 1979, 1988; Wearne and Creighton, 1988). One of the points that has generated controversy has been the interpretation of the experimental data obtained using two different redox pairs, oxidized glutathione-reduced glutathione (GSSG/GSH) and oxidized dithiothreitol-reduced dithiothreitol (DTTox/DTI'rcd). The two reagents react differently with thiol groups (Gilbert, 1994). In a series of recent works, Rothwarf and Scheraga (1993a-d) have studied the regeneration pathway of RNase A using both reagents and different temperatures. These authors conclude that the regeneration pathways depend on the nature of the redox agent as well as the redox potential at which they
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are used. The use of GSSG/GSH favors multiple regeneration pathways, whereas the use of DT'['ox/DT'['red favors the regeneration through fewer pathways. Also, they conclude that the regeneration processes with the two types of redox agents proceed through different pathways. Early attempts to get fully regenerated RNase A using the DTI'ox/DTI'red couple were unsuccessful (Wearne and Creighton, 1988; Creighton, 1988). The first successful regeneration of fully reduced RNase A with the DTT couple (under anaerobic conditions) was that reported by Rothwarf and Scheraga (1991). A major regeneration pathway was identified when the regeneration pathway of RNase A was followed using the pair DT'['ox/DTTred (Rothwarf and Scheraga, 1993a,b). This pathway proceeds through a ratedetermining step involving an intramolecular rearrangement of one or more three-disulfide species and is essentially identical to the model proposed by Creighton (1988). Thus, the experimental observations and conclusions of Creighton can be, in part, reconciled with the multiple pathway model of Scheraga. The major difference in the form of the model involves the proposal by Creighton that the pathways of formation and reduction of the native protein are the same. Although the regeneration of native RNase A using the glutathione redox couple has been attained successfully in many laboratories, the isolation of disulfide-bonded intermediates has been hampered by the complexity of the equilibrium mixture of redox isomers. The sole exception was a three-disulfide species, isolated in homogeneous form, lacking the 40-95 disulfide bond. This intermediate was shown to have a compact nativelike structure and enzymatic activity, but a less stable conformation that was disrupted by only half the concentration of urea required to unfold native RNase A (Creighton, 1980; Galat et al., 1981). In the reduction pathway of RNase A, specially using dithiothreitol (DTI') as reducing agent, it has been difficult not only to isolate but even to detect intermediate species with broken or rearranged disulfide bonds (Creighton, 1977, 1979, 1988). However, Rothwarf and Scheraga (1991), using DTI" as reducing agent and 2-aminoethylmethane thiosulfonate as blocking agent, reported the isolation of a nativelike three-disulfide species lacking the 65-72 disulfide. This intermediate, named des-[6572]-RNase A, has been characterized by examining its thermal transition curve by means of two- and three-dimensional homonuclear 1H NMR spectroscopy and by determining its activity (Talluri et al., 1994). The results obtained indicate that the thermal stability of des-[65-72]-RNase A is substantially lower than that of native RNase A. However, the des-
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[65-72]-RNase A presents 81% of the enzymatic activity of native RNase A in front of C>p, whereas the conformations found for this derivative exhibit a well-defined structure where the native protein has a regular backbone, the only major differences occurring in a loop comprising residues 60-72. The close structural similarity between des- [65-72] RNase A and the native enzyme suggested that no major conformational change was necessary for their interconversion. It is then concluded that in reduction pathways that include des-[65-72]-RNase A, the ratedetermining step corresponds to a partial unfolding event in one region of the protein and not to a global conformational unfolding process, as suggested by earlier works (Creighton, 1988). The results further suggest that in the regeneration pathways involving des-[65-72]-RNase A the loop region from 60 to 72 are the last to fold. Li et al. (1995), reexamining the reduction pathway of RNase A over a range of pH, temperature, and concentrations of DTT, reported the isolation and characterization of two nativelike three-disulfide intermediates, named It and 12. It was identified as des-[65-72]-RNase A, previously reported by Talluri et al. (1994), whereas I2, which lacks the 40-95 disulfide bond (des-[40-95]-RNase A), could be the intermediate reported by Creighton (1980) found in the refolding pathway of RNase A using oxidized glutathione. This last intermediate has only 5% of the activity of native RNase A in front of C>p, suggesting that Lys-41 is more perturbed in des-[40-95]-RNase A than in des-[65-72]-RNase A. Both intermediates lack the same disulfide bonds that are lacking in the two thermally stable RNase A mutants (C[65-72]S and C[40-95]S) (Laity et al., 1993). The reduction kinetics of both native RNase A and purified intermediates in unblocked form suggest that the reduction unfolding of RNase A with DTI" proceeds through parallel pathways in which each of the above intermediates is involved, and that two distinct local unfolding events rather than a global one are involved in the rate-limiting steps (Li et al., 1995). These results are contrary to the current view that protein unfolding generally follows an all-or-none mechanism and that the rate-limiting step is controlled by an extensive rearrangement of the native structure (Creighton, 1994).
ACKNOWLEDGMENTS
This work was supported by Grant PB93-0872 from the DGICYT of the Ministerio de Educaci6n y Ciencia (Spain) and Grant GRQ93-2093 from the CIRIT of the Generalitat
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de Catalunya (Spain). We also thank the Fundaci6 M. F. de Roviralta (Barcelona, Spain) for grants for the purchase of equipment.
REFERENCES
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10 Crystallographic Studies of Ribonuclease Complexes GARY
L. GILLILAND
The Center for Advanced Research in Biotechnology of the University of Maryland Biotechnology Institute and National Institute of Standards and Technology Rockville, Maryland 20850
I. II. III. IV.
V. VI.
VII. VIII.
IX.
Introduction Phosphate/Sulfate-Free RNase Phosphate/Sulfate Binding Site Substrate Analog-RNase Complexes A. U p c A - R N a s e B. C p A * - R N a s e S C. 2 ' - F - U p A - R N a s e S D. d ( C p A ) - R N a s e A E. d(pA)4-RNase A F. d ( A p T p A p A p G ) - R N a s e A Transition State Analog-RNase Complex Product-RNase Complexes A. 4 t U - R N a s e S and A p C - R N a s e S B. p T p - R N a s e S C. 2 ' - C M P - R N a s e A D. O3-2'-CMP-RNase A E. 3 ' - C M P - R N a s e A Nonproductive Ligand-RNase Complexes Other Ligand-RNase Complexes A. 41-DNP-RNase S B. 7,41-DNP-RNase A C. T-H12-RNase A and U-HI 19-RNase A D. Derivative II-RNase A E. Propidium Iodide-RNase A-d(pA)4 Semisynthetic RNases A. N-Terminus RNase Variants B. C-Terminus RNase Variants
RIBONUCLEASES: STRUCTURES AND FUNCTIONS
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X. Conclusions A. Substrate Binding and Specificity B. Catalysis References
I. INTRODUCTION Bovine pancreatic ribonuclease A ( E C 3.1.27.5; RNase A)* hydrolyzes single-stranded R N A . Ribonuclease S (RNase S), an active variant of RNase A, can be produced by cleavage of the peptide bond between residues 20 and 21 by subtilisin (Richards and Vithayathil, 1959). Early biochemical studies showed that RNase A prefers a pyrimidine base on the 3' side of the R N A cleavage site [for a review see, Richards and Wyckoff (1971)]. Crystallographic studies of RNase A and RNase S with substrate analogs, a transition state analog, products, and inhibitors have been carried out to provide a structural basis of substrate specificity, catalysis, and inhibition to c o m p l e m e n t the biochemical and biophysical investigations of this enzyme. These studies have provided many details of how nucleotides and other ligands interact with the enzyme. The initial crystallographic studies have been summarized in several reviews (e.g., Richards and Wyckoff, 1971; Blackburn and Moore, 1982; Wlodawer, 1985). * RNase A, Bovine pancreatic ribonuclease A (EC 3.1.27.5); RNase S, ribonuclease S; UpcA, uracil-3',5'-adenosine in which the 5'-oxygen atom of the ribose attached to the adenosine is replaced with a methylene group; CpA*, cytidylyl-2',5'-adenosine;2'-F-dUpA, 2'-deoxy-2'-fluorouridylyl-3',5'-adenosine; d(CpA), deoxycytidyl-3',5'-deoxyadenosine; UV, uridine vanadate; 4tU, 4-thiouridylic acid; ApC, adenylyl-3',5'-cytidine; pTp, thymidine 3',5'-diphosphate; 2'-CMP, cytidine 2'-phosphate; O3-2'-CMP, cytidine N(3)-oxide 2'-phosphate; 3'-CMP, cytidine 3'-monophosphate; 2',5'-CpG, cytidylyl-2',5'-guanosine; 3',5'-dCpdG, deoxycytidylyl-3',5'-guanosine; Os-2'-GMP, 8-oxoguanosine 2'-phosphate; 41-DNP-RNase S, a covalent complex formed by the reaction of dinitrofluorobenzene with Lys-7; 7,41-DNP-RNase S, a covalent complex formed by the interaction of 1,5difluoro-2,4-dinitrobenzene with Lys-7 and Lys-41; propidium iodide-d(pA)4-RNase A, propidium iodide noncovalently bound to the d(pA)4-RNase A complex; derivative II, a covalent complex formed by reacting 6-chloropurine 9-/3-D-ribofuranosyl 5'-monophosphate with the a-amino group of Lys-1; T-H12-RNase, [N~Z-[[[(3'-deoxy-3'thymidinyl)amino]carbonyl]methyl]histidine-12]-ribonuclease A; U-Hll9-RNase, [N8~[[[(3'-deoxy-3'-uridinyl)amino]carbonyl]methyl]histidine-119]-ribonuclease A.
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The original structure determinations of RNase A (Avey et al., 1967; Kartha et al., 1967) produced details of the overall fold of the polypeptide chain and the location of the active site (Fig. 1). Difference Fourier studies at 5.5/k resolution with the inhibitor 2'-cytidylic acid were used to identify the location of the active site in the cleft on the surface of the protein (Avey et al., 1967) and hence the locations of His-12, Lys-41, and His-ll9. The location of Lys-41 was further pinpointed by difference Fourier analysis of crystals containing mercury associated with a chemically modified Lys-41 converted to homocysteine. Indirect evidence was also obtained from difference Fourier studies of crystals grown in the presence of arsenate rather than phosphate (Kartha et al., 1967, 1968). These studies localized the active site to the cleft of the kidneyshaped molecule. The structure determination of RNase S (Wyckoff et al., 1967a,b, 1970) complemented the structure of RNase A and provided new details of the structure and its interactions with nucleotides. Early crystallographic studies of RNase S at 6.0 A resolution (Wyckoff et al., 1967a) used halogenated nucleotides for two purposes: (1) for identifying the active site and (2) for solving the crystallographic phase problem. The results of four separate studies with different iodonucleotides identified a single major site, again in the cleft of the molecule. When the structure was extended to 3.5 fi~ resolution the first details of nucleotides binding at the active site began to emerge (Wyckoff et al., 1967b). The proximity of His-12 and His-119 were confirmed, and the residues Val-43, Thr-45, Phe-120, and Ser-123 were found to form a nucleotide base binding; pocket. Extending the structure determination of RNase S to 2.0 A resolution confirmed the presence of a sulfate ion in the active site (Wyckoff et al., 1970). One nucleotide, 5-iodouridine 3'-monophosphate, used in the phase calculations, was also used as a derivative in the 6.0 structure determination of a monoclinic (space group C2) crystal form of RNase S (Mitsui and Wyckoff, 1975). As already mentioned, early studies provided the first structural details of how substrate analogs and inhibitors interacted with the enzyme. High-resolution structure determinations have extended this knowledge. Presented here is a summary of the results of crystallographic studies of the free enzyme compared with the enzyme in complex with phosphate or sulfate, substrate analogs, a transition state analog, and products. In addition, a brief discussion of crystallographic results from studies of semisynthetic forms of the enzyme is presented along with the crystallographic results of nonproductive complexes of nucleotides with the en-
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zyme and of covalently modified variants of the molecule. The results of these studies provide a series of snapshots along the catalytic pathway, show how sequence variants perturb the structure, and reveal structural details of the interactions of the enzyme with noncovalent and covalent ligands. These and further crystallographic studies, along with other biochemical and biophysical studies, are essential for a complete understanding of how the structure of this enzyme relates to its function. To simplify the discussion of the complexes, the following conventions for naming the nucleotide binding sites have been used. The R N A substrate binds to RNase A with a specific orientation with respect to the active site. The ribose, base, and phosphate groups of the nucleotide that binds on the 3' side of the cleavage site near His-12 and His-119 are associated with what are designated the R1, B1, and P1 subsites, respectively (Richards and Wyckoff, 1973). The phosphate associated with the 5'-hydroxyl of the RI ribose is designated as the P0 subsite. The ribose, base, and phosphate groups of the nucleotide that is on the other side of the cleavage site are associated with the R2, B2, and P2 subsites, respectively. The next base in the sequence is at the B3 subsite and so on. The details of the interactions at each of these subsites will be described below.
!1. P H O S P H A T E / S U L F A T E - F R E E
RNase
The first crystal forms, designated Y, Z, and W, of RNase S were all grown from solutions containing salt that included a high concentration of ammonium sulfate. Thus, it was not surprising to find sulfate bound at the active site. On the other hand, crystals of RNase A were grown from alcohol solutions with no salt added (Fankuchen, 1941). Nevertheless, phosphate or sulfate was observed bound at the active site. The presence of the anion at the active site made it difficult to interpret the changes in the structure induced by ligand binding. Some time after the structure determination of RNase A and RNase S, two structures of unligated RNase A were reported, one at 1.5 ,~ (Campbell and Petsko, 1987) and the other at 1.26 A (Svensson et al., 1986; Wlodawer et al., 1988). Crystals for the 1.5 ,~ structure determination were grown in the usual way, but before X-ray data collection the crystals were soaked in solutions at elevated pH (pH 9.0) to remove the sulfate before returning them to solutions at pH 6.8. The pH of the
10 Crystallography of RNase Complexes
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crystals was altered in steps of 1 pH unit each day, with soaking at elevated pH for 8 to 9 days. The crystals used in the 1.26 A structure were grown from a salt-free commercial preparation of the enzyme. The 1.5 A electron density map of the unligated RNase A indicated that the phosphate at the P1 subsite had been replaced by two water molecules (Campbell and Petsko, 1987). One water molecule occupies the position of one of the oxygen atoms of the phosphate group and forms a hydrogen bond with His-12 and the other water molecule. Only small changes in the conformation of residues in the region of the active site are observed. The largest movement in this region was for the side chain of Lys-41 (0.38 A). The two active site histidines maintain the positions found in the presence of sulfate. Gln-11 is the only other active site residue that adjusts its position. The side chain rotates so that the side chain nitrogen atom moves toward the former sulfate binding site. The largest changes observed at Asp-38, Arg-39, Leu-51, Thr-87, Asn103, and A s n - l l 3 are not directly associated with the active site. The 1.26 ,~ structure provided many additional details of unligated RNase A (Svensson et al., 1986; Wlodawer et al., 1988). The active site of this structure is shown in Fig. 2. The final structure from this study included 13 residues with two conformations. The proximity of three of these residues, Val-43, Asp-83, and Arg-85, led to the hypothesis that the conformations of these residues may be interdependent. Only one of these 13 residues, Gin-l l, is associated with the active site. It was suggested that the presence of the phosphate or sulfate in the active site stabilizes a single conformation of the residue. This structure also revealed the association of a single molecule of 2-methyl-2-propanol bound to the enzyme on the side opposite from the active site. This molecule binds in a small hydrophobic pocket formed by amino acid residues 15-16, 20-25, 82, and 99-101, a segment of one of the antiparallel/3-strands. Several other interesting features were described, including an intercalated water network found between two symmetry-related molecules in the crystal lattice and a substantial deviation in the conformation of Gin-101 compared to that found in other structures. The conformations of residues in the active site of the phosphate-free enzyme were very similar to that observed for the protein in the presence of the phosphate or sulfate anion. A few subtle changes were observed. A slight movement of His-ll9 resulting from a rotation of A'2 by - 1 2 ~ appears to allow His-ll9 to form a more favorable hydrogen bond to Asp-121. The side chain of Lys-7 has only an altered position, a 0.68 shift, and G l n - l l was observed as mentioned above, in two conforma-
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Gary L. Gilliland
tions. As in the 1.5 ,& structure described above, two new waters appear in the region of the active site replaced by the sulfate or phosphate. A water molecule (W203) occupies the 0-2 atom position of the phosphate. The second water molecule (W271) is close to the phosphate binding site, but it does not bind within hydrogen bonding distance to the first (W203) [a difference from that observed by Campbell and Petsko (1987)]. Additional water molecules and solvent reordering were observed.
III. P H O S P H A T E / S U L F A T E B I N D I N G SITE
The initial crystallographic structures of both RNase A at 2.0 A resolution (Kartha et al., 1967) and RNase S at 2.0 ,& resolution (Wyckoff et al., 1970) provided details of the overall fold of the polypeptide chain. In both structures phosphate or sulfate was found bound in the active site cleft (see Fig. 3). These structures, determined by the multiple isomorphous replacement techniques, have been extended to high resolution and refined by a variety of least-squares procedures. The structure of RNase A has been refined first to 2.0 A resolution using both X-ray (Wlodawer et al., 1982) and combined X-ray and neutron (Wlodawer and Sjolin, 1983) diffraction data. The neutron data provided the means to assign the orientation of the active site histidine side chains, His-12 and His-119 (Wlodawer and Sjolin, 1981), and established the location of Lys-41 (Wlodawer and Sjolin, 1983). The RNase A structure was further extended to 1.45 /k and then to 1.1 ,& (Svensson et al., 1991) resolution. The RNase S structure and its comparison with RNase A has been recently reported at 1.5 ,& resolution (Kim et al., 1992). The location of the divalent anion, the phosphate or sulfate, that is bound in the P1 subsite of RNase is consistent in all of the structures. The oxygen atoms of the divalent anion form hydrogen bonds with N e2 atom of His-12, N 81 atom of His-119, and N atom of Phe-120. The oxygen atoms also form a hydrogen bond with a water that in turn hydrogen bonds with the O e~ atom of Gln-ll. The high-resolution structure also showed that the side chain of Gln-ll exists in two different conformations. His-119 is also seen to occupy one of two alternate conformations, designated A and B (Wlodawer and Sjolin, 1981; Wlodawer et al., 1982; Svensson et al., 1991; Kim et al., 1992). In the 1.45 ,~ structure (Borkakoti et al., 1982) the residue is disordered and is present in both conformations.
10
Crystallography of RNase Complexes
311
Another notable difference in the region of the binding site is the position of Lys-41. This varies considerably in the enzyme structures (Wlodawer et al., 1986). The differences in the crystallization conditions for the various RNase A and RNase S crystals used in the structure determinations are likely to contribute to the differences observed for the location and conformation of the active site residues.
IV. SUBSTRATE A N A L O G - R N a s e C O M P L E X E S
Many investigations of substrate analog-RNase complexes have been carried out to provide details on substrate binding. These studies have taken advantage of the fact that RNase has a high specificity for a pyrimidine base at the B~ subsite and that catalysis requires the presence of a 3'-5' phosphodiester linkage between the riboses and the 2'hydroxyl of the ribose. The following discussions are summaries of the results from the structure determinations of complexes of RNase S with UpcA (a dinuc|eotide in which the 5'-oxygen atom of the ribose attached to the adenosine is replaced with a methylene group) (Richards and Wyckoff, 1971, 1973), with cytidylyl-2',5'-adenosine (CpA*) (Wodak et al., 1977), and with 2'-deoxy-2'-fluorouridylyl-3',5'-adenosine (2'-FUpA) (Pavlovsky et al., 1978), and of complexes of RNase A with deoxycytidyl-3',5'-deoxyadenosine [d(CpA)](Zegers et aL, 1994), with d(pA)4 (McPherson et al., 1986a-c), and with d(ApTpApApG) (FontecillaCamps et al., 1994). The space group, the unit cell parameters, and resolution for each of these studies are presented in Table I. The Protein Data Bank (PDB) identifier is also provided if coordinates have been deposited (Bernstein et al., 1977).
A. UpcA-RNase The three-dimensional structure of the UpcA-RNase S complex was essentially the first substrate analog-RNase complex to be described in detail (Richards and Wyckoff, 1971, 1973). The methylene group replacing the ester oxygen atom prevents the enzyme from cleaving the dinucleotide. Though the structure was not refined, many details of the interaction of the dinucleotide have been verified in other high-resolution,
TABLE I Ribonuclease A-Ligand Complexes Determined by X-Ray Crystallography Ligand" SO4 SO4 (pH 5.5) SO4 (pH 4.75) Substrate analogs UpcA CpA* 2'-F-dUpA d(CpA) d(pA)4 d(ApTpApApG) Transition state analog UV UV
Protein
Space group
Unit cell dimensions a, b, c (~,);/3 (~
Resolution PDB code:
RNase A RNase S RNase S
P3221 P3121 P3t21
64.8, 64.8, 65.2 44.7, 44.7, 97.0 44.5, 44.5, 97.6
2.2 1.6 1.6
RNase RNase RNase RNase RNase RNase
P3121 P3~21 P3~21 P2~ P212121 P2t2121
44.5, 44.5, 97.3 44.8, 44.8, 96.7
1.8 2.0 3-2.5 1.4 2.5 2.3
P2z P21
30.3, 38.4, 53.7:106.4 29.8, 38.2, 53.2:106.1
S S S A A A
RNase A RNase A
30.0, 38.3, 53.2 44.4, 75.3, 44.6 71.9, 43.2, 43.8
2.0 1.3
1RPH 1RNU 2RNS
Ref. Zegers et al., 1994 Kim et al., 1992 Kim et al., 1992
1RCN
Gilliland et aL, 1994 Wodak et al., 1977 Pavlovsky et al., 1978 Zegers et al., 1994 McPherson et al., 1986a Fontecilla-Camps et al., 1994
6RSA 1RUV
Borah et al., 1985 Ladner et al., 1996
1RPG
Products 4tU ApC pTp 2'-CMP O32'-CMP 3'-CMP Nonproductive ligands 2',5',CpG 3',5'-dCpdG O8-2'-GMP
C2 C2 C2 P2~ P2l P3221
101.5, 31.8, 69.6; 90.0 101.6, 32.1, 69.5; 89.6 -30.5, 38.7, 53.2; 106.5 30.5, 38.1, 53.2; 105.9 65.3, 65.3, 65.5
4.0 4.0 4.0 1.6 2.3 2.2
RNase A RNase A
P21 P21
RNase A
P2~
30.3, 30.3, 30.5, 30.5,
1.5 1.5 1.9 2.3
RNase RNase RNase RNase RNase RNase
S S S A A A
38.3, 38.3, 38.7, 38.2,
52.9; 105.9 52.9; 105.9 53.7; 106.3 53.5; 106.0
1ROB 1RPF 1RNC 1RND 1RCA
Torii et al., 1978 Mitsui et al., 1978 Iwahashi et al., 1981 Lisgarten et al., 1993 Borkakoti, 1983 Zegers et al., 1994 Aguilar et al., 1991 Aguilar et al., 1991 Lisgarten et al., 1995 Borkakoti, 1983; Aguilar et al., 1992
Abbreviations for ligands: UpcA, uracil-3',5'-adenosine in which the 5'-oxygen atom of the ribose attached to the adenosine is replaced with a methylene group; CpA*, cytidylyl-2',5'-adenosine; 2'-F-dUpA, 2'-deoxy-2'-fluorouridylyl-3',5'-adenosine; d(CpA), deoxycytidyl-3',5'-deoxyadenosine; UV, uridine vanadate; 4tU, 4-thiouridylic acid; ApC, adenylyl-3',5'-cytidine; pTp, thymidine 3',5'-diphosphate; 2'-CMP, cytidine 2'-phosphate; 032 'CMP, cytidine N(3)-oxide-2'-phosphate; 3'-CMP, cytidine 3'-monophosphate; 2',5'-CpG, cytidylyl-2',5'-guanosine; 3',5'-dCpdG, deoxycytidylyl-3' ,5 'guanosine; O8-2'-GMP, 8-oxoguanosine 2'-phosphate. b PDB, Protein Data Bank. a
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Gary L. Gilliland
highly refined complex structures. In this structure the uridine bound in the B1 subsite has the most extensive interactions with the enzyme. Thr45 forms two hydrogen bonds with the base. The 0 - 2 of the base forms a hydrogen bond with the backbone NH group of Thr-45, and the base N-3 atom interacts via a hydrogen bond with the hydroxyl group of Thr45. Richards and Wyckoff (1971) proposed that the Thr-45 hydroxyl group could play a role as hydrogen bond donor or acceptor depending on which base, uridine or cytosine, was present in the B1 subsite. It was suggested that Asp-83 has the potential to form a hydrogen bond with the Thr-45 hydroxyl group. An interaction of the 0 - 4 with the Ser-123 hydroxyl group was proposed. The interactions involving Asp-83 and Ser-123 were later shown to be mediated by water molecules (Giniland et al., 1994). The base aromatic ring is also close to the aromatic ring of the side chain of Phe-120. The ribose 2 ' - O H is near His-12. The position of His-ll9 was stabilized from its disordered state in RNase S, but the specific interaction with the ligand was not defined. The potential for a hydrogen bond between the side chains of His-ll9 and Asp-121 was described. At the B2 subsite the adenosine interacts with the side chains of Asn-71 and G l u - l l l and the base stacks against His-119. It should be mentioned that Lys-41 was not observed to interact with the substrate analog.
B. CpA*-RNase S The crystal structure of the complex of RNase S with bound CpA* was determined by difference Fourier methods at 2.0 A resolution followed by model building and energy minimization (Wodak et al., 1977). The cytosine base occupies the B1 subsite interacting with Thr45 and Phe-120. The cytosine N-3 atom forms a hydrogen bond with the O ~ atom of Thr-45. The second hydrogen bond observed in earlier studies between the base 0 - 2 atom and the N atom of Thr-45 is not observed (the distance between the atoms is 4.0 A). No hydrogen bonds between the ribose at the R1 subsite and the enzyme are found. The N ~ atom of Lys-41 can be positioned within 2.6 A from the O-1' atom of the ribose, allowing it to participate in catalysis. The ribose is in the C3'-endo conformation. The phosphate forming the 2',5' linkage is in nearly the same position as observed in other complexes forming hydrogen bonds with the side chains of Gln-11 and His-12, and the carbonyl oxygen atom of Phe-120. The difference electron density map revealed
10 Crystallographyof RNase Complexes
315
a negative peak at the location of the phosphate. Several explanations were suggested, but it undoubtedly indicates a shift in position of the phosphate between that observed in the native conformation and the CpA* complex. The ribose associated with the adenosine base at the R2 subsite is also in the C-3'-endo conformation, and one possible hydrogen bond between the ribose O-1' atom and the N 81 atom of His-ll9 is observed. The adenosine in the B2 subsite forms three potential hydrogen bonds (N-1 with O e of Glu-lll, N-6 with O e of Gin-69, and N-6 with 0 8 of Asn-71). Several enzyme residues appear to alter their positions when the nucleotide binds. These include Lys-41, Thr-45, Gin-69, Asn71, and His-ll9. No movement of His-12 is detected.
C. 2 ' - F - U p A - R N a s e S
The structure of the complex of RNase S with the inhibitor 2'-F-UpA has been determined by difference Fourier techniques at pH 5.5 and 7.2 at 2.5 and 3.0 A, respectively (Pavlovsky et al., 1978). This inhibitor was chosen because the presence of the substitution of the 2'-OH with 2'-F inhibits the cleavage of the 3'-ester linkage. The structure determined at pH 7.2, near the pH with maximum activity, was virtually identical with that at pH 5.5. The structure, for the most part, agrees with other complexes with the exception that the adenosine is in the syn conformation. This is contrary to other reports (e.g., Zegers et al., 1994) and is likely a result of the fitting procedure. The positions of the active site residues His-12, His-ll9, Gln-ll, Asn-44, Asp-121, Phe-120, and Thr-45 agree with other studies. It appears that Gln-ll, Phe-120, and Asp-121 show the most marked alterations in their positions, along with His-119, which appears to move closer to the phosphate in the P1 subsite. Observed remote from the active site were conformational changes in the loop, composed of amino acid residues 65-72. The interaction of the adenosine with the protein and changes in positions of Phe-120 and Asp121 may be contributing to this displacement.
D. d ( C p A ) - R N a s e A
The complex of d(CpA) with RNase A was determined at 1.4 ~, resolution (Zegers et al., 1994). The binding of d(CpA) in the active site
316
Gary L. Gilliland
of RNase A is illustrated in Fig. 4. At the B1 subsite the cytosine forms hydrogen bonds with Thr-45 and is in close contact with the aromatic ring of Phe-120, consistent with the complexes described above. Two water molecules were also found within hydrogen-bond distance to the N-4 atom of the base. Both of the ribose moieties are in the C - 2 ' - e x o conformation. The ribose does not form any hydrogen bonds directly to the protein, but the 0 - 4 ' and 0-5' oxygen atoms form hydrogen bonds with two water molecules. His-12 and Lys-41 interact with the 2'-OH. His-12 also hydrogen bonds with a P1 phosphate oxygen. One water molecule forms hydrogen bonds with two of the phosphate oxygen atoms. Lys-41 has a second conformation involved in a hydrogen bond with Asn-44. His-119 interacts with 0 - 4 ' and 0-5' oxygens and hydrogen bonds with 0 8of Asp-121. The deoxyribose at P2 does not form any direct interactions with the protein. The adenosine forms two hydrogen bonds with Asn-71 of the protein and two hydrogen bonds with water molecules. The adenosine base displaces two water molecules near the N-1 and N-6 positions of the base. No interactions with Gin-69 or Glu-111 are apparent.
E. d(pA)4-RNase A A complex of four oligomers of d(pA)4 bound to a single molecule of RNase A provides the first details of how a ribonuclease may interact with a long single strand of RNA (McPherson et al., 1986a-c). A "virtual" continuous RNA strand of 12 nucleotides is observed. Only two of the four nucleic acid strands, I and IV, are associated with a single ribonuclease in the crystal lattice. The other two strands bridge pairs of enzyme molecules. Strand II has three of the nucleotides, beginning with the 5' end directly associated with the enzyme. Strand III has only a single nucleotide at its 5' end associated with RNase A. The 12 phosphates of the virtual continuous strand of RNA, beginning from the 5' end, would interact with nine lysine and arginines, Lys-7, Lys-41, Lys-66, Arg-85, Arg-39, Lys-91, Lys-98, Arg-33, and, finally, Lys-31. The primary associations of the RNA with ribonuclease are electrostatic interactions between the phosphate moieties and the positively charged groups of the surface. The base closest to the B1 subsite does not bind in this pyrimidinespecific binding pocket. Specific interactions between the bases and the protein are apparent only at the B2 and B3 subsites. The other bases appear to project out into the solvent channels and are apparently stabilized by base-stacking interactions.
10 Crystallography of RNase Complexes F. d ( A p T p A p A p G ) - R N a s e
317
A
The crystal structure of d ( A p T p A p A p G ) - R N a s e A complex provides information on a substrate analog binding at four different nucleotide binding sites (Fontecilla-Camps et al., 1994). Adenosine is at the B0 subsite. Interestingly, the primary interactions of the base, including two hydrogen bonds and van der Waals interactions, are with a symmetryrelated protein molecule. Thus, the authors postulate that RNase has no specific B0 site. No direct hydrogen bond between RNase A and the ribose is observed at the R0 subsite, but a bridging water molecule between the N; atom of Lys-66 and the ribose 5'-oxygen atom is found. Lys-66 could form a direct interaction with the phosphate at the P0 subsite. However, in this crystal form it is involved in a salt bridge with Glu-49 of a symmetry-related molecule. The nucleotides with groups at the BlR~P1 and B2R2P2 sites are very well defined in the electron density map of the RNase A d(ApTpApApG) complex. The thymidine, ribose, and phosphate at the B1, Rl, and P1 subsites are consistent with those found in other RNase-nucleotide complexes. As observed in other studies, the base at the B~ subsite does not directly interact with the side chains of either Phe-120 or Ser-123. The adenosine at the B2 subsite hydrogen bonds with Gin-69 and Asn-71 and stacks with the imidazole of His-119. The ribose is observed to have only a single hydrogen bond between the 0-5' atom and the N ~ atom of His-119. The phosphate at the P2 subsite interacts with the side chain of Lys-7 (2.8 A). In contrast to the other three sites, the electron density at the B3R3P3 site is not well defined, and only interpretable density for the base at the presumed B3 subsite is observed. The position and orientation of the ribose and phosphate could not be detected. The adenosine, occupying the B3 subsite, stacks with the base at the B2 subsite. No interpretable density for the final nucleotide of the DNA oligomer is present in the electron density map.
V. TRANSITION STATE A N A L O G - R N a s e COMPLEX
Lindquist and co-workers (1973) proposed that a complex of uridine with certain oxyvanadium compounds that can achieve a pentacoordinate
318
Gary L. Gilliland
arrangement might be useful as a stable analog for the pentacoordinate trigonal bipyramid (TBP) phosphorane. The TBP phosphorane is postulated to be an important structure along the reaction pathway, consistent with the "in-line" inversion displacement mechanism (Usher et al., 1970, 1972). Three different crystal structures of complexes of uridine vanadate (UV) inhibitor with RNase A have been reported (Alber et al., 1983; Wlodawer et al., 1983; Ladner et aL, 1996), and one of these has been compared with the results of high-field NMR studies of the complex (Borah et al., 1985). The primary result of the first X-ray crystal structure analysis (Alber et al., 1983) was the determination of how a "transition state"-like analog influences the mobility of the important interacting active site residues. It was concluded that Lys-41, in particular, becomes highly immobilized in the presence of UV and that the side chain of Lys-41 interacts directly with one of the vanadate oxygens. The second structure of UV-RNase A, determined using combined 2.0 A X-ray and 2.2 A neutron data (Wlodawer and Sjolin, 1983), and its comparison with the NMR results (Borah et al., 1985), revealed a number of important features of the active site. Similar to the earlier results, Lys-41, His12, and His-119 side chains were in direct contact with UV; however, the interactions were inconsistent with expectations based on the proposed mechanism. His-12 was found positioned to interact with an equatorial vanadate oxygen whereas the side chain of Lys-41 was closest to an axial 0-2' oxygen. Recently, UV-RNase A has been investigated using oneand two-dimensional NMR techniques (Veenstra and Lee, 1994). Based on the appearance or absence of certain cross-peaks in the NMR spectrum, it was postulated that the position of the side chain of His-ll9 may be different for the complex in solution than in the crystal structure. The three-dimensional structure of UV-RNase A complex has been extended to 1.3 A resolution (Ladner et al., 1996). The refinement process and the interpretation of the final structure were aided by high-level ab initio quantum mechanical results on model oxyvanadates. One of the primary goals of this study was to determine whether the UV exists as the mono- or dianion by a direct comparison of the high-resolution structure of UV-RNase A with those determined computationally, and to determine what significance, if any, the protonation state may have on the true phosphate-ester hydrolysis mechanism. Using the theoretically determined vanadate structures as initial starting points, the effect of the starting structure on the outcome of the X-ray data refinement process was also evaluated, leading to an unambiguous final structure of this important enzyme-inhibitor complex.
10 Crystallography of RNase Complexes
319
As seen in Fig. 5, the P1 subsite of RNase A contains the amino acid residues Gln-ll, His-12, His-ll9, Lys-41, Phe-120, and water molecules, W229, W411, and W425, each of which is in close contact with the TBP vanadate structure. The apical vanadate oxygens, 0-2' and O-3V, form interactions with a number of different P1 components. Specifically, 0-2' is within hydrogen bonding distance to Lys-41 N c (2.7/k) and His12 N e2 to a lesser extent (3.4 A), whereas O-3V interacts primarily with His-ll9 N 81 (2.6 .&), W425 (2.2 A), and W411 (2.7 ~,). The position of the essential residues at the P1 subsite, His-12, Lys-41, and His-ll9, relative to the UV inhibitor is illustrated in Fig. 5. The primary interaction between equatorial vanadate oxygens and components in the P1 site include O-3', which is within hydrogen bonding distance to W411 (2.6 A). The 0-3' oxygen atom also interacts with His-ll9 N 81 (3.2 A). The O-1V atom interacts with His-12 (N e2 (2.7 A), Phe-120 N (2.9 A), and W229 (2.7 A). Finally, O-2V is within hydrogen bonding distance to Gln-11 (N ~2 (2.9 A). Although Lys-7 has been postulated as a key active site residue in catalysis, it is not part of the P1 subsite and may not be directly involved in binding of the UV. The Rl ribose-binding subsite includes residues His-12, Lys-41, Val43, His-119, Phe-120, and water molecules W358, W411, and W414. Many components formally considered part of the Rl subsite are, however, also part of the P1 subsite and interact primarily with the TBP vanadate atoms. For example, W411 is only 3.5 A from C-3' and 3.6 A from C-4' of the ribose, but its primary interactions are with 0-3' of the ribose and O-3V of the TBP vanadate as previously described. In fact, the only significant interactions in the R~ subsite are those associated with Val43 and W358. The ribose ring oxygen O-1' hydrogen bonds to the Val43 O (3.4 A). W358 is within hydrogen bonding distance to a number of the Rl components, including W414 (3.1 A), Val-43 backbone (3.0 to O and 3.2 A to N), the ribose oxygen O-1' (3.1 A), and W365 (2.8 A), found outside the R~ subsite. From its many close contacts, W358 may be an essential structural component in this region. The B1 pyrimidine-binding subsite formed by the six amino acid residues, His-12, Val-43, Asn-44, Asn-45, Arg-83, Phe-120, and two water molecules, W310 and W373, has been described in detail by Gilliland and co-workers (1994). An additional water, W401 could be considered part of the B~ subsite (3.9 A from C-6) but is more properly considered part of the P0 subsite because it is 3.1 A from 0-5' of the ribose. As seen in Fig. 5, two important hydrogen bond interactions are formed directly between protein atoms and the base. The peptide N of Thr-45
320
Gary L. Gilliland
is 2.9 /k from the uridine 0-2, whereas 0 71 of Thr-45 is 2.7 ,~ from N-3 of the uridine. The two conserved water molecules, W310 and W373, mediate the interaction of the nucleotide base with the protein and allow for the accommodation of any pyrimidine base in the B1 pocket.
VI. PRODUCT-RNase COMPLEXES
Crystallographic investigations of a number of RNase-product complexes have been carried out to provide information concerning specificity and catalysis. The first such structure determinations involved complexes of RNase S with 3'-CMP, 3'-UMP, 2'-CMP, dTpT, 4-thiouridine, 3' :5'-A>p, 3'-AMP, 5'-AMP, and ATP with RNase S (Richards and Wyckoff, 1971). These studies established that the B1 subsite was occupied in complexes of 3'-CMP, 3'-UMP, 2'-CMP, dTpT, and 4-thiouridine with RNase S. The B2 subsite was occupied in the 3' :5'-A>p, 3'-AMP, 5'-AMP, and ATP complexes with RNase S. These studies were followed by more detailed reports of other low-resolution difference Fourier studies of 4-thiouridylic acid (4tU) (Torii et al., 1978), adenylyl-3',5'-cytidine (ApC) (Mitsui et al., 1978), and thymidine 3',5'-diphosphate (pTp) (lwahashi et al., 1981). Further high-resolution refined structures of cytidine N(3)-oxide-2'-phosphate (O3-2'-CMP) (Borkakoti, 1983), cytidine-2'phosphate (2'-CMP) (Borkakoti, 1983; Howlin et al., 1987), and cytidine 3'-monophosphate (3'-CMP) were reported. As with the other complexes previously described, the space group and unit cell parameters and resolution for each of these studies are presented in Table I. For the refined structures deposited in the Protein Data Bank (Bernstein et al., 1977) the PDB identifier is included.
A. 4 t U - R N a s e S and A p C - R N a s e S
The three-dimensional structure of the product complex of 4tU with RNase S was carried out with the C2 crystal form of RNase S (see Table I) by difference Fourier analysis by Torii and co-workers (1978). A preliminary study of a different crystal form of this complex was reported
10 Crystallographyof RNase Complexes
321
earlier by Richards and Wyckoff (1971). The base and ribose of the inhibitor were observed in the B1 and R1 subsites, respectively. The phosphate was also proposed to bind at the P~ subsite. The anti conformational relationship between the base and the ribose was proposed, as observed in other nucleotide ribonuclease complexes. A second 4.0 ,~ study using the same crystal form examined the binding of adenylyl3',5'-cytidine (ApC) to RNase S (Mitsui et aL, 1978). The difference Fourier analysis showed that the cytidine binds in the B~ and R1 subsites and that the adenosine protrudes into the solvent channel. It was postulated that the phosphate group is associated with the P0 subsite interacting with Lys-66, but no difference density was present in the electron density map to support this (Iwahashi et al., 1981).
B. p T p - R N a s e S
The three-dimensional structure determination of pTp was undertaken specifically to investigate the interactions between the enzyme and the phosphate in the P0 subsite (Iwahashi et al., 1981). This difference Fourier study at 4.0 ~ resolution with the C2 crystal form clearly showed the base and ribose bound to the B1 and R1 subsites, respectively. The conformation of the nucleotide was described as most likely gauche § for the C4'-C5' torsion angle 0, the base is in the anti conformation with respect to the ribose, and the ribose pucker is either C - 2 ' - e n d o or C-3'-endo. The P0 subsite may be occupied by the 5' phosphate and one of the phosphate oxygen atoms is presumably bound to the N c of Lys-66.
C. 2 ' - C M P - R N a s e A
The crystal structure of the complex of 2'-CMP-RNase A was initially determined at 2.3 ~ resolution (Borkakoti, 1983; Howlin et al., 1987) then extended to 1.6 ,/k resolution (Lisgarten et al., 1993). The highresolution structure (Lisgarten et al., 1993) confirmed the conformational features found at lower resolution. The cytidine is in the B1 pyrimidine binding site next to Thr-45. The 0-2, N-3, and N-4 atoms form hydrogen
322
Gary L. Gilliland
bonds with the N, O v~, and O 3,1 atoms of Thr-45, respectively. Only one of the ribose oxygen atoms forms a hydrogen bond with the protein. The 0 - 2 ' is a hydrogen bond donor to the backbone O atom of Val-43. Three water molecules hydrogen bond to oxygen atoms associated with the ribose. The ribose conformation was very close to the preferred region for C3'-endo. The inhibitor phosphate moiety forms the most complex interactions with the protein. The O-1 atom forms hydrogen bonds with a water molecule, the amide N atom of Phe-120, and the N ~2 atom of His-12. The 0-2 atom is within hydrogen bond distance of the N .2 atom of Gin-l l, and the 0-3 atom forms a hydrogen bond with the N 82 atom of His-ll9. The solvent analysis of this structure and its comparison with the unligated RNase A (Wlodawer et al., 1988) indicate that the inhibitor displaces six water molecules. A crystallographic investigation of the binding of 2'-CMP as a function of temperature was carried out by Rasmussen et aL (1992). This difference Fourier study showed that 2'-CMP would not bind to the protein at 212 K, but binding was evident at 228 K. If the temperature is lowered to 212 K the inhibitor cannot be flushed from the active site. The investigators propose that this data supports a biphasic behavior of the enzyme in which above a transition temperature (220 K) substrate binding requires large-scale anharmonic motion of protein atoms. Below the transition point the motions of individual atoms are dominated by harmonic vibrations that are not sufficient to allow binding on the time scale of the experiment (days) or to allow release of the inhibitor if it were bound at a higher temperature.
D. O3-2'-CMP-RNase A The three-dimensional structure of O3-2'-CMP was determined at 2.3 A resolution (Borkakoti, 1983). The inhibitor binds in the B~, R~, and P~ subsites as expected with the base anti to the ribose. The ribose is in the 3 ' - e n d o conformation as observed for other nucleotide complexes. Lys-41 moves to interact with one of the phosphate oxygens. His-ll9 is observed to be in the B conformation. The ribose 0 - 3 ' may sterically block His-119 from residing in the A conformation. The active site residues, Gln-11, His-12, Thr-45, and Phe-120 show little movement. The N ~ atom of the side chain of Lys-66 moves toward the 0 - 5 ' of
10 Crystallography of RNase Complexes
323
the ribose. Four waters are expelled from the active site on inhibitor binding.
E. 3 ' - C M P - R N a s e
A
Another product complex, 3'-CMP, has been investigated crystallographically (Zegers et al., 1994). This inhibitor binds in the BIR1PI site. The nucleotide in this complex binds in the C - 2 ' - e x o anti conformation. As with other pyrimidines, the base forms two hydrogen bonds with Thr-45 and interacts with the aromatic side chain of Phe-120. The NH2 group forms hydrogen bonds with two waters that in turn interact with the side chains of Thr-45, Asp-121, and Ser-123. The ribose is clearly seen in the electron density forming strong hydrogen bonds between the 2'-OH group and the side chains of His-12 and Lys-41. The phosphate hydrogen bonds to His-12, the carbonyl oxygen of Phe-120, and three water molecules. It also interacts with His-119, but the histidine in this complex is found in the B conformation. Thus, His-119 forms no hydrogen bond with Asp-121. Comparing 3'-CMP with 2'-CMP we see similar interactions; however, several differences are evident. The conformations of the riboses are different, in 2'-CMP only weak interactions are found between the 2 ' - 0 atom and Lys-41, and no hydrogen bond with His-12 is found. Instead, Gln-ll hydrogen bonds to the phosphate.
VII. N O N P R O D U C T I V E L I G A N D - R N a s e COMPLEXES
The high-resolution crystallographic studies of several inhibitor complexes revealed a novel nonproductive mode of binding designated as retro-binding (Aguilar et al., 1989, 1991, 1992). The first two complexes discovered to have this anomalous binding property were cytidylyl-2',5'guanosine (2'5'-CpG) and deoxycytidylyl-3',5'-guanosine (3'5'-dCpdG). These initial reports were carried out by soaking the inhibitor into previously grown crystals. The 1.9 A structure of RNase A that cocrystallized with 3',5'-dCpdG (Lisgarten et al., 1995) is virtually identical to that reported for the soaked crystals. In both complexes, neither the sulfate nor phosphate is displaced from the active site, water in the active site cleft is displaced by the ligand, the guanine is found associated with the
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Gary L. Gilliland
B1 subsite, the ribose electron density is weak, and electron density for the cytosine is not observed. In the P1 subsite of these complexes the sulfate/phosphate ion adjusts its position slightly compared with that observed in other RNase structures (e.g., Wlodawer et al., 1982). This is due in part to two hydrogen bonds formed between the N-1 and N-2 atoms of the guanine with the 0-2' and 0-5' atoms of the anion, respectively. The anion also forms hydrogen bonds with the amino acid residues Gln-ll, His-12, Lys-41, His-ll9, and Phe-120. The three residues, Lys-41, His-ll9, and Phe-120, show significant displacement of the side chain atoms relative to the unligated RNase A. The anion moves by 0.8 A toward the guanine binding a different orientation when compared with the uninhibited structure (Borkakoti et aL, 1983, 1984), resulting in an altered hydrogen bonding pattern. The base forms two hydrogen bonds at the B1 subsite, between the 0-6 and N-7 atoms of the guanine and the N and OG atoms of Thr-45, respectively. Both the phosphate and the ribose atoms of the inhibitors have high temperature factors and the ribose has only one direct interaction with the protein, a hydrogen bond between the 0-2' atom of the ribose and the O of Asp-121. The guanine is anti to the ribose in both complexes. His-119 is found in the A conformation in these inhibitor complexes. When this alternative form of nucleotide binding was found, the original interpretation of the 8-oxoguanosine 2'-phosphate (OS-2'-GMP) RNase A complex structure indicating productive binding (Borkakoti et aL, 1983) was reexamined and found incorrect (Aguilar et al., 1992). This inhibitor was found to bind in a manner similar to that observed for the 2',5'-CpG and 3',5'-dCpdG complexes (Aguilar et al., 1991).
VIII. O T H E R L I G A N D - R N a s e C O M P L E X E S
A number of covalently modified RNase S and RNase A structures have been determined, providing structural details for the interpretation of chemical modification studies and for the investigation of the roles of specific residues. Two crystal structures of RNase covalently modified with dinitrophenol derivatives have been reported. The first describes the chemical addition of 2,4-dintrophenyl group to the N~ atom of Lys41 of RNase S (Allewell et al., 1973). The second describes RNase A
10
Crystallography of RNase Complexes
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that has been chemically cross-linked by the reaction with 1,5-difluoro2,4-dinitrobenzene (7,41-DNP-RNase A) (Weber et al., 1985). The structures of two forms of [N~2-[[[(3'-doexy-3'-thymidinyl)amino]carbonyl] methyl]H12]-RNase A (T-H12-RNase A) and [N81-[[[(3'-deoxy-3 'uridinyl)amino]carbonyl]methyl]H119]-RNase A (U-H119-RNase A), RNase A with deoxynucleosides covalently bound to the active site histidines, His-12 and His-119, have been reported (Nachman et al., 1989, 1990). The structure of derivative I I - R N a s e A, obtained by the reaction of the a-amino group of Lys-1 with 6-chloropurine /3-D-ribofuranosyl 5'-monphosphate, has been determined (Boque et al., 1994). One noncovalent complex of propidium iodide bound to the RNase A-d(pA)4 has been determined (McGrath et aL, 1987). This study was undertaken to assess how this compound, a competitive inhibitor, influences the activity of RNase. A crystallographic summary of this and the covalently modified derivatives mentioned above are presented in Table II.
A. 4 1 - D N P - R N a s e S
One of the first structural investigations of a covalent modification involved the determination of the structure of RNase S modified by the chemical addition of 2,4-dintrophenyl group (DNP) to the N ~ atom of Lys-41 (Allewell et al., 1973). The structure of 41-DNP-RNase S was determined by difference Fourier techniques in the presence and the absence of 3'-CMP at 3.0 and 2.0 A, respectively. In both cases the DNP was not found to occlude the active site, but it was found in a cleft associated with the side chains of residues Gln-11, Asn-34, Asp-38, and Arg-39 and the backbone atoms of Asn-34 and Asp-38. The presence of the DNP moves the side chain of Lys-41 away from the sulfate or phosphate in the active site in a direction opposite from the 3'-CMP binding site. Other side chains in the active site, e.g., His-12 and His119, were not observed to move. The results of the study showed only slight perturbations in the conformation of the overall structure of RNase S and suggested that the reduction in activity and the binding affinity of 3'-CMP may be a result of the loss of the interaction of Lys-41 (a 3.0 ,~ displacement and a loss of the positive charge of the N ;) with the substrate.
TABLE II Crystal Structure Determinations of Ligands Covalently Bound to Ribonuclease
Complex '~ 41-DNP-RNase S 7,41-DNP-RNase A Propidium iodide T-H12-RNase A U-Hll9-RNase A Derivative II
Space group
P3221 P2~212z P2z2t2 P2z2z2z P2z2z2z C222z
Unit cell dimensions a, b, c (A,)
37.1, 44.6, 53.1, 52.8, 75.7,
41.3, 75.2, 64.6, 64.1, 57.9,
75.6 44.7 73.6 73.2 53.3
Resolution (A)
PDB code b
Ref.
-2.0 2.9 1.8 1.8 2.1
---8RSA 9RSA 1RBN
Allewell et al., 1973 Weber et al., 1985 McGrath et al., 1987 Nachman et al., 1990 Nachman et al., 1990 Boque et al., 1994
a Abbreviations for complexes: 41-DNP-RNase S, a covalent complex formed by the reaction of dinitrofluorobenzene with Lys-7; 7,41-DNP-RNase S, a covalent complex formed by the interaction of 1,5-difluoro-2,4-dinitrobenzene with Lys-7 and Lys-41; propidium iodide-d(pA)4-RNase A, propidium iodide noncovalently bound to the d(pA)4-RNase A complex; derivative II, a covalent complex formed by reacting 6-chloropurine 9-fl-D-ribofuranosyl5'-monophosphate with the a-amino group of Lys-1; T-H12-RNase A, [Ne2-[[[(3'-deoxy-3'-thymidinyl)amino]carbonyl]methyl]histone_12]_ribonuclease A; U-H119-RNase A, [N~l-[[[3'-deoxy-3'-uridinyl)amino]carbonyl]methyl]histidine-119]-ribonuclease A. b PDB, Protein Data Bank.
10
Crystallography of RNase Complexes
327
B. 7 , 4 1 - D N P - R N a s e A
The crystal structure determination of 7,41-DNP-RNase A was determined to aid in the interpretation of data that show the thermal stability of this RNase A-dinitrophenyl adduct (Weber et al., 1985). The 2.0 A 7,41-DNP-RNase A structure was compared with the joint neutron and X-ray refined 2.0 ~, RNase A (Wlodawer and Sjolin, 1983). The perturbations in the structure are attributable to three differences: (1) difference in crystallization conditions, (2) the presence and interactions of the cross-linking agent, and (3) difference in crystal packing resulting from different crystal symmetry. The side chains of Lys-7 and Lys-41 reorient because of their linkage to the DNP. The conformations of Gln-ll and Arg-39 are also altered because of the presence of the DNP. His-ll9 moves perhaps because of the presence of the DNP and perhaps because of the absence of phosphate or sulfate in the active site. The analysis of the structure suggested that the structure was very similar to the uncross-linked RNase A except in the region of the crosslink agent, 35 common water sites with the native enzyme; the temperature factor distribution was similar to that observed for other proteins. The analysis of the structure suggests that the difference in stability between the native and cross-linked proteins is a manifestation of the difference in entropic effects of the unfolded states.
C. T - H 1 2 - R N a s e A and U - H l l 9 - R N a s e A
The structures of two forms of ribonuclease A with deoxynucleosides covalently bound to the active site histidines, His-12 and His-119, respectively, have been reported (Nachman et al., 1989, 1990). The His-12 derivative, T-H12-RNase A, is a major product of the reaction of 3'(bromoacetamido)-3'-deoxythymidine with ribonuclease A. The His-119 derivative, U - H l l 9 - R N a s e A, is a major product of the reaction of 3'(bromoacetamido)-3'-deoxyuridine reaction with RNase A. The covalently linked nucleoside in T-H12-RNase A does not bind in the traditional way to the R1 and B~ subsites, but it does interact with atoms of residues Phe-120, Asp-121, and Asn-67 through hydrogen bonds. In this structure the orientation of the nucleoside is fixed; the electron density is clearly defined despite the high temperature factors of the inhibitor
328
Gary L. Gilliland
atoms compared with those of the local protein atoms. The base is in an anti conformation with the ribose in the C - 2 ' - e x o conformation. The position of the inhibitor is consistent with the fact that the enzyme is completely inhibited. The inhibition is a result of steric inaccessibility of the active site. In contrast, the U - H l 1 9 - R N a s e complex is less well defined in the electron density maps. The nucleoside base projects out into the solvent channel of the crystal and forms a contact with a symmetry-related molecule in the crystal lattice. The electron density is very weak, indicating disorder. No apparent intramolecular contacts are evident, possibly accounting for the disorder, and may account for the 1% activity observed for this derivative.
D. Derivative l l - R N a s e
A
The three-dimensional structure of derivative II-RNase A was determined at 2.1 A resolution (Boque et al., 1994). The structure provides information concerning the P2, B3, and R3 subsites and helps explain the catalytic properties of this chemically modified RNase A. The structural interpretation is complicated by the interaction of the modified N terminus with side chains of a symmetry-related molecule. The N ~atom of Lys1 hydrogen bonds with the O r of Tyr-76 of another RNase A molecule in the crystal lattice. The N-7 atom of the purine ring of the derivative is also within hydrogen bonding distance to the O ~ atom of Ser-77 of the same symmetry mate. The investigators argue that the purine/3-Dribofuranosyl-5'-monophosphate substituent is nestled in a pocket that corresponds to the B3 subsite. The conformation of this pocket is close to that observed in the structure of unmodified RNase A. No hydrogen bonds between the base and the protein are present, but there is a covalent link between the protein and the base. At the putative R3 subsite, the ribose is partially disordered, but there is a hydrogen bond between the 0 - 3 ' atom of the sugar and the amine N atom of Ala-4. At the P2 subsite the phosphate of the derivative approaches within 4.6 of the N ~ atom of Lys-7. In this derivative, His-12 is in the normally observed conformation, Lys-41 is in a fully extended conformation within hydrogen bond distance of the Asn-44 side chain, and His-119 is observed in both the A and B conformations assigned 0.7 and 0.3 occupancies, respectively. A phos-
10 Crystallographyof RNase Complexes
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phate is not at the catalytic center, only water molecules forming a complex hydrogen bond network with the active site residues. They argue that the proximity of the negatively charged phosphate of the covalently attached nucleotide to His-ll9 affects catalysis. That is, the interaction with Asp-121 is destabilized, allowing the histidine to exist in a partial conformation that is catalytically incompetent.
E. Propidium l o d i d e - R N a s e A - d ( p A ) 4
The structure of propidium iodide bound to the d(pA)4-RNase A complex has been determined at 2.9 ~, resolution (McGrath et al., 1987). The difference Fourier analysis indicates a single binding site on the side of the molecule opposite from the active side. The residues Phe-8, Met-13, Glu-49, Leu-51, Val-54, and Gln-55 are associated with the binding site. Local conformational changes are observed on binding of the ligand. Changes in residues near the active site residues 12-18, 46-55, and 113-117 are also observed, though they are more than 10 A from the propidium-binding site. No binding of the propidium to the DNA was observed. The authors suggest that the changes in conformation of the enzyme structure near the active site are at least in part responsible for the behavior of propidium as a competitive inhibitor.
IX. SEMISYNTHETIC RNases
The three-dimensional structures of a number of semisynthetic RNases have been determined to aid in the investigation of the roles of residue catalysis, substrate binding, and protein folding. The sensitivity of ribonuclease to different proteases allows removal of either the N or the C terminus, residues 1-20 (Richards and Vithayathil, 1959), and residues 118-124 (Lin et al., 1970), respectively. Studies have shown that synthetic peptides for either the N or the C terminus will associate with the proteolyzed enzyme, forming a productive complex. The properties of a number of noncovalent complexes have been characterized and their crystal structures have been determined (see Table III).
TABLE III Crystal Structure Determination of Semisynthetic Variants of Ribonuclease
Variant a 1-15:21-124 1-15:21-124 1-15(4-F-l-ilisl2) :21-124 1-15:21-124 1-15(M13G) :21-124 1-15(M13A) :21-124 1-15(M13ANB) :21-124 1-15(M13V) :21-124 1-15(M131) :21-124 1-15(M13L) :21-124 1-15(M13F) :21-124 1-118:111-124 1-118: lll-124(DI21N) 1-118:111-124(0121A) 1-118: Ill-124(F120L) 1-118: Ill-124(FI20Y)
Space group
P3121 P3~21 P3~21 P3t21 P3~21 P3~21 P3~21 P3~21 P3~21 P3~21 P3121 P3121 P3121 P3~21 P3~21 P3121
Unit cell dimensions a, b, c (,~) 44.5, 44.5, 44.5, 44.4, 44.5, 44.5, 44.5, 44.3, 44.3, 44.3, 44.3, 67.7, 64.7, 64.7, 67.3, 68.2,
a Synthetic peptides are indicated in boldface type. b PDB, Protein Data Bank.
44.5, 97.2 44.5, 97.6 44.5, 97.2 44.4, 97.2 44.5, 97.3 44.5, 97.0 44.5, 97.2 44.3, 97.6 44.3, 97.7 44.3, 97.9 44.3, 97.9 67.7, 65.0 64.7, 64.9 64.7, 64.9 67.3, 64.9 68.2, 65.2
Resolution (.&) 2.6 1.6 2.6 3.0 1.8 2.0 1.7 1.8 1.8 1.7 1.7 1.8 2.0 2.0 2.0 2.0
PDB code b
1RNV
1RBF 1RBC 1RBD 1RBI 1RBG 1RBH 1RBE 1SRN 2SRN 3SRN 1SSA 1SSB
Ref. Taylor et al., 1981 KJm et al., 1992 Taylor et al., 1981 Taylor et al., 1985 Varadarajan and Richards, Varadarajan and Richards, Varadarajan and Richards, Varadarajan and Richards, Varadarajan and Richards, Varadarajan and Richards, Varadarajan and Richards, Martin et al., 1987 deMel et al., 1992 deMel et al., 1992 deMel et al., 1994 deMel et al., 1994
1992 1992 1992 1992 1992 1992 1992
10 Crystallography of RNase Complexes
331
A. N-Terminus RNase Variants Several RNase structures that include a semisynthetic N-terminal peptide have been determined. The first such report included a comparison of the active 1-15:21-124 structure with 1-1514-F-His12] :21-124 (Taylor et al., 1981) (boldface type indicates synthetic peptides). Residues 16-20 were not included because these four appeared disordered in the RNase S electron density map (Wyckoff et al., 1970). The absence did not affect the activity of the enzyme. A comparison of the active 1-15:21124 with the 2.0 .~ RNase S structure (Wyckoff et al., 1970) showed little change in the region of the active site. The structure of the 1-1514F-His12] :21-124 variant confirmed the modeling predictions, showing that residues 3-13 maintain an c~-helical conformation, that His-12 maintains its orientation, and that the complex-stabilizing electrostatic and nonbonded interactions are maintained. The visibility of the fluorine atom on the imidazole ring also confirmed the orientation of the His12 imidazole ring. A later report describes the three-dimensional structure of a variant that has a peptide engineered to maintain the structural features essential for catalysis, but significantly altered in sequence (Taylor et al., 1985). The N-terminal amino acid sequence of the native enzyme, K E T A A A K F E R O H M D S , was changed to A E A A A A K F A R A H M A A . The structure of the 1-15:21-124 variant confirmed the modeling predictions, showing that residues 3-13 maintain an c~-helical conformation, that His-12 maintains its orientation, and that the complex-stabilizing electrostatic and nonbonded interactions are maintained. The three-dimensional structures of seven RNase S variants with substitutions of M13 for glycine, alanine, c~-amino-n-butyric acid (ANB), valine, isoleucine, leucine, and phenylalanine have been determined to high resolution (1.7-2.0 A) (Varadarajan and Richards, 1992). These mutants are being used to investigate the role in protein folding of interior residues. Each variant has a variety of changes resulting from the substitution. The largest changes are in the surface loop composed of residues 65-69. The largest changes were observed for the M13F and M13G mutants. Surprisingly, the loop moves away from the molecule in both cases. The reason for this change is unclear. Changes are observed in the interactions between Lys-66 and Asp-121. The effects on nucleotide binding and enzymatic activity have not yet been reported.
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Gary L. Gilliland
B. C-Terminus RNase Variants
The structure determination of semisynthetic RNase (1-118:111-124) was carried out to establish a basis for the interpretation of results of structures of variants in which alterations of the amino acid sequence of the tetradecapeptide have been made (Martin et al., 1987). The structure was similar to that of RNase A (Wlodawer and Sjolin, 1983), with no redundant structure visible in the electron density map. The visible structure is composed of the enzyme residues 1-113 and the tetradecapeptide residues 114-124. There are significant deviations in the positions of atoms associated with residues 111-113 of the enzyme and 114-118 of the tetradecapeptide when compared with RNase A. The active site residue positions are in general agreement with those seen in other RNase structures. In the active site, His-12 interacts via a hydrogen bond with the sulfate in the active site, the N; of Lys-41 is involved in a hydrogen bond with the 0 8 atom of Asn-44, and His-ll9 is found in the B conformation. The role of Asp-121 in catalysis was examined by the construction and characterization of two semisynthetic RNase (1-118:111-124) variants in which the aspartate was replaced with asparagine and alanine (deMel et al., 1992). The three-dimensional structures were compared with the unsubstituted semisynthetic RNase (1-118:111-124) (Martin et al., 1987). A sulfate ion is found in the active site of all three enzymes. The orientation and interaction of the sulfate with the protein vary substantially for each variant. His-12 remains virtually unchanged, with the N ':2 atom involved in a hydrogen bond with one of the sulfate ion oxygen atoms. His-119 in all three structures is observed predominantly in the B conformation. The histidine side chain of the alanine and asparagine variants moves 0.5 ,~ closer to the sulfate ion than it does when aspartate is at position 121. At position 121 the backbone atoms are similar in all structures, but the side chain atoms diverge substantially at the C ~ atom of both alanine and asparagine. The divergence increases out at the 0 81 and N 82 atoms for the asparagine. Because His-ll9 is in the B conformer, no direct interaction is observed between His-ll9 and the side chain at position 121. In the structural comparisons the largest difference in the overall structure was observed in the region of residues 65-72. The data suggested that these differences result from alterations in the interactions of Lys-66 with the residue at position 121. Recently the structures of two semisynthetic variants that replace Phe120, a highly conserved residue (Beintema et al., 1988), with tyrosine
10 Crystallography of RNase Complexes
333
and leucine have been determined at 2.0/k resolution (deMel et al., 1994). The active site residues His-12, Lys-41, and His-119 have conformations similar to those observed in the fully active semisynthetic RNase 1118:111-124 (Martin et al., 1987). His-119 is in the B conformation. The hydrogen bond between N ~2 of G l n - l l and 0-3 of the sulfate found in RNase 1-118:111-124 is present in the F120Y variant, but it is not in the F120L. The three hydrogen bonds between residue Lys-66 and Asp121 are similar to those in the F120Y variant, but two of the three hydrogen bonds are absent in the F120L variant as a result of movement of the 65-72 loop with respect to that observed in RNase 1-118:111-124. The movement of this loop is also observed with two other catalytic defective analogs, D121N and D121A. This suggests that the displacement may be one of the reasons for reduction in catalytic efficiency. The sulfate ion in both of the variants is significantly different from that observed for RNase 1-118:111-124. The substitution of phenylalanine with leucine in the F120L variant significantly changes this region of the protein. The side chain of Ser-123 moves toward the leucine side chain and the C ~ and C~ atoms of the leucine move toward the sulfate ion. The substitution of phenylalanine with tyrosine produces changes resulting from an introduction of the hydroxyl group. The O r of Ser-123 forms a hydrogen bond with the tyrosine hydroxyl group, and a bridging water between the O r of Ser-123 and the hydroxyl of Tyr-120 is observed.
X. C O N C L U S I O N S
The crystallographic studies described above reveal a complex picture of enzyme-nucleotide interactions that illustrate both the versatility of the enzyme in binding a spectrum of nucleotides and the specific requirements for catalysis.
A. Substrate Binding and Specificity The mono- and dinucleotides that bind "productively" have provided details of nucleotide interactions at the BIRIP~ and B2R2P2 binding sites, and in a few instances provide details of the P0 subsite (Richards and Wyckoff, 1971; Mitsui et al., 1978; Iwahashi et al., 1981). The structure
334
Gary L. Gilliland
of the derivative I I - R N a s e A complex (Boque et al., 1994) provides a description of some of the interactions at the B3R3P3 binding site. The d ( A p T p A p A p G ) - R N a s e A complex (Fontecilla-Camps et al., 1994) provides more details of nucleotide binding at the BoRoP0, B1R1P1, B2R2P2, and B3R3P3 sites. The structures of most of the nucleotide complexes are the result of having at least tried to take advantage of the pyrimidine specificity of the B~ subsite. The interactions of nucleotides (and analogs) at the B1R1P1 site are now well characterized. The high-resolution studies of nucleotides and analogs have produced a consistent picture of the interactions between the ligand and protein. The ingenious mechanism of binding that allows the binding of either cytosine or uracil at the B~ subsite has emerged (Gilliland et al., 1994). This involves the ability of the hydroxyl group of Thr-45 and two conserved water molecules to change their roles of proton donor or acceptor in hydrogen bonds between the base and protein amino acid residues. This is illustrated in Fig. 6. In contrast, the interactions, conformation, and/or orientation of the sugar and phosphate of the R1 and P1 subsites show considerable variability. The conformation and orientation of the sugar at the R1 ribosebinding subsite are dependent on the type of sugar and the phosphate
Se~ 0 ...... ', "'H
Thr 45\
H 10 " " ,0,,
""
H'"
,," H
',,
.H
.-
Asp83 H/ ' ' ,,0~ H
Uracil
-.
I ribose
Se~.~
0 . . . . .', . . , H~O "'H ,, ~0" H ",
Thr 45 \
.,"
H ""
'i-l~ /H'"
H,I '
" "O~H
Cytosine
N~ N''H'''- .0
I ribose
Fig. 6. The hydrogen bonding pattern observed for the uridine base and proposed for the cytosine base in the RNase B~ subsite (Gilliland et al., 1994).This highlights the change in role from proton donor to proton acceptor for the two conserved water molecules and the Thr-45 O~ in hydrogen bond interactions with the bases.
10 Crystallography of RNase Complexes
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linkage. The sugar always has few direct interactions with the enzyme, and in the higher resolution structure determinations, one to several water molecules may hydrogen bond to the oxygem atoms of the sugar. The orientation of the phosphate in the P1 subsite is dependent on whether it is linked to the sugar via a 2'- or 3'-ester linkage. A hydrogen bond between His-12 and a phosphate oxygen is normally present, and at least one hydrogen bond between the phosphate and side chain of His-119 is present. Studies report either the direct interaction or no interaction of the side chain of Lys-41 with the phosphate moiety. The water structure associated with the phosphate group also varies considerably, depending on the class of ligand and the resolution of the reported study. The interactions of nucleotides at other proposed sites, BoRoP0, BER2P2, and BaR3P3, are not as well defined. At the BoRoP0 site, only the phosphate at the P0 subsite has shown specific interactions. The structures of the ApC-RNase S complex (Mitsui et al., 1978) and the pTp-RNase S complex (Iwahashi et al., 1981) show that the phosphate at the P0 subsite interacts with the positively charged side chain of Lys66. There is currently no information concerning specific interaction of the base with the protein at this position. Unfortunately, the adenosine of the d(ApTpApApG)-RNase A (Fontecilla-Camps et al., 1994) forms specific interactions with a symmetry molecule in the crystal lattice. The interactions between nucleotides and RNase at the B2R2P2 site are based on the results of complexes with adenosine at the B2 subsite. The early difference Fourier studies of nucleotides (Richards and Wyckoff, 1973; Wodak et al., 1977; Pavlovsky et al., 1978) have been augmented with well-defined high-resolution structures (FontecillaCamps et al., 1994; Zegers et al., 1994). The early structures found that adenosine forms hydrogen bonds between the N-1 and N-6 atoms of adenine and oxygen atoms of the side chains of Glu-111, Gin-69, and Asn-71. The base stacks against the side chain of His-119. However, the structure of the high-resolution complex, d(CpA)-RNase A, reveals only two hydrogen bonds with Asn-71, and no other hydrogen bonds with the protein are observed. For the d ( A p T p A p A p G ) - R N a s e A complex, Fontecilla-Camps et al. (1994) observed hydrogen bonds with Gin-69 and Asn-71. Both high-resolution structures confirm the base stacking interaction with His-119. The crystallographic studies of the d ( A p T p A p A p G ) - R N a s e A (Fontecilla-Camps et al., 1994), the d(pA)4-RNase A (McPherson et al., 1986a-c), and the derivative II-RNase A (Boque et al., 1994) complexes
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provide a few details of the B3R3P3 binding site. The adenosine base at this site in the d ( A p T p A p A p G ) - R N a s e A structure stacks with the adenosine base at the B2 subsite, and the electron density for the phosphate and ribose is not observed. No details have been reported for the d(pA)4 interactions. In the derivative II-RNase A complex the adenosine is covalently linked to the c~-amino group of the N-terminal lysine of the enzyme. There are no hydrogen bonds with the B3 adenosine. There is one hydrogen bond to the 0 - 3 ' atom of the ribose with the backbone N atom of Ala-4. Further crystallographic studies with other nucleotides are needed to clarify the current picture of the BoRoPo, B2R2P2, and B3R3P3 binding sites.
B. Catalysis
The catalytic reaction by which RNase A hydrolyzes phosphate-ester linkages of single-stranded RNA is a two-step general acid-general base mechanism. In the first step, the 3'-5' phosphate linkage is cleaved via intramolecular transphosphorylation, leading to a 2',3'-cyclic phosphate intermediate. In the subsequent step, the cyclic phosphate is hydrolyzed by a water molecule within the active site to form a 3'-monophosphate ester as the final product. Both steps in the reaction are known to proceed by an in-line inversion displacement mechanism based on isotopic labeling experiments (Usher et al., 1970, 1972). In this mechanism His-12 and His-ll9 are proposed as the critical catalytic residues in the enzyme active site. These two residues are thought to act as the general base and general acid, respectively, in the first step, reversing their roles in the second step of the reaction. The crystallographic studies previously described consistently show that these residues are well positioned to fulfill their roles in catalysis. Other residues within the active site, such as Lys-41 and Asp-121, are also catalytically important based on chemical modification and site-directed mutagenesis studies (e.g., Hits et al., 1965; Stern and Doscher, 1984); however, their specific role in the catalytic mechanism remains unclear. Of these important residues, the role of Lys-41 has been the most highly debated. The position of Lys-41 within the active site allows direct contact with the substrate, and it has been argued that this residue acts to stabilize anionic intermediates through electrostatic interactions (Haydock et al., 1990). Several RNase complex
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structures show the direct interaction of the Lys-41 side chain with the substrate, supporting this contention.
ACKNOWLEDGMENTS
The author thanks J. Ladner for reviewing the manuscript, and J. Dill for assistance in the preparation of the figures.
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Aguilar, C. F., Palmer, R. A., and Moss, D. S. (1989). An unusual inhibitor complex of ribonuclease exhibiting retro binding: A 1.5 ,~ refinement study of bovine pancreatic ribonuclease A complexed with cytidylyl (2'-5') phosphoryl guanosine. In "Structure and Chemistry of Ribonucleases" (A. Pavlovsky and K. Polyorkov, eds.), pp. 31-40. International Union of Crystallography, Moscow. Aguilar, C. F., Thomas, P. J., Moss, D. S., Mills, A., and Palmer, R. A. (1991). Novel nonproductively bound ribonuclease inhibitor complexes--High resolution X-ray refinement studies on the binding of RNase-A to cytidylyl-2',5'-guanosine (2',5'CpG) and deoxycytidylyl-3',5'-guanosine (3',5'dCpdG). Biochim. Biophys. Acta 1118, 6-20. Aguilar, C. F., Thomas, P. J., Mills, A., Moss, D. S., and Palmer, R. A. (1992). Newly observed binding mode in pancreatic ribonuclease. J. Mol. Biol. 224, 265-267. Alber, T., Gilbert, W. A., Ponzi, D. R., and Petsko, G. A. (1983). The role of mobility in the substrate binding and catalytic machinery in enzymes. Ciba Found. Symp. 93, 4-24. Allewell, N. M., Mitsui, Y., and Wyckoff, H. W. (1973). X-Ray diffraction studies of e41-dinitrophenyl-ribonuclease-S. J. Biol. Chem. 248, 5291-5298. Avey, H. P., Boles, M. O., Carlisle, C. H., Evans, S. A., Morris, S. J., Palmer, R. A., Woolhouse, B. A., and Shall, S. (1967). Structure of ribonuclease. Nature (London) 213, 557-562. Beintema, J. J., Schuller, C., Irie, M., and Carsana, A. (1988). Molecular evolution of the ribonuclease superfamily. Prog. Biophys. Mol. Biol. 51, 165-192. Bernstein, F. C., Koetzle, T. F., Williams, G. J. B., Meyer, E. F., Jr., Brice, M. D., Rogers, J. R., Kennard, O., Shimanouchi, T., and Tasumi, M. (1977). The protein data bank: A computer-based archival file for macromolecular structures. J. Mol. Biol. 112, 535-542. Blackburn, P., and Moore, S. (1982). Pancreatic ribonuclease. In "The Enzymes" (P. D. Boyer, ed.), Vol. 15, pp. 317-433. Academic Press, New York. Boque, L., Coil, M. G., Vilanova, M., Cuchillo, C. M., and Fita, I. (1994). Structure of ribonuclease A derivative II at 2.1 A resolution. J. Biol. Chem. 269, 19707-19712. Borah, B., Chen, C.-W., Egan, W., Miller, M., Wlodawer, A., and Cohen, J. S. (1985). Nuclear magnetic resonance and neutron diffraction studies of the complex of ribonuclease A with uridine vanadate, a transition state analog. Biochemistry 24, 20582067.
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Lindquist, R. N., Lynn, J. L., Jr., and Lienhard, G. E. (1973). Possible transition-state analogs for ribonuclease. The complexes of uridine with oxovanadium(IV) ion and vanadium(V) ion. J. Am. Chem. Soc. 95, 8762-8768. Lisgarten, J. N., Gupta, V., Maes, D., Wyns, L., Zegers, I., Palmer, R. A., Dealwis, C. G., Aguilar, C. F., and Hemmings, A. M. (1993). Structure of the crystalline complex of cytidylic acid (2'-CMP) with ribonuclease at 1.6 ,~, resolution. Conservation of solvent sites in RNase-A at high-resolution structures. Acta Crystallogr. D 49, 541-547. Lisgarten, J. N., Maes, D., Wyns, L., Aguilar, C. F., and Palmer, R. A. (1995). Structure of the crystalline complex of deoxycytidylyl-3',5'-guanosine (3',5'-dCpdG) cocrystallized with ribonuclease at 1.9 ,A resolution. Acta Crystallogr. D 51, 767-771. McGrath, M., Cascio, D., Williams, R., Johnson, D., Greene, M., and McPherson, A. (1987). Propidium binding to a ribonuclease-DNA complex: X-Ray and fluorescence studies. Mol. Pharmacol. 32, 600-605. McPherson, A., Brayer, G. D., and Morrison, R. D. (1986a). Crystal structure of RNase A complexed with d(pA)4. J. Mol. Biol. 189, 305-328. McPherson, A., Brayer, G. D., and Morrison, R. D. (1986b). Structure of the crystalline complex between ribonuclease A and d(pA)4. Biophys. J. 49, 209-219. McPherson, A., Brayer, G., Cascio, D., and Williams, R. (1986c). The mechanism of binding of a polynucleotide chain to pancreatic ribonuclease. Science 232, 765-768. Martin, P. D., Doscher, M. S., and Edwards, B. F. P. (1987). The refined crystal structure of semisynthetic ribonuclease at 1.8 ,~ resolution. J. Biol. Chem. 262, 15930-15938. Mitsui, Y., and Wyckoff, H. W. (1975). The crystal structure of monoclinic ribonucleaseS at six ,~,ngstroms resolution. J. Mol. Biol. 94, 17-31. Mitsui, Y., Urata, Y., Torii, and Irie, M. (1978). Studies on the binding of adenylyl-3',5'cytidine to ribonuclease. Biochim. Biophys. Acta 535, 299-308. Nachman, J., Miller, M., Gilliland, G. L., Carty, R., Pincus, M., and Wlodawer, A. (1989). Crystal structure of two covalent nucleoside derivatives of ribonuclease A. In "Structure and Chemistry of Ribonucleases" (A. Pavlovsky and K. Polyorkov, eds.), pp. 22-30. International Union of Crystallography, Moscow. Nachman, J., Miller, M., Gilliland, G. L., Carty, R., Pincus, M., and Wlodawer, A. (1990). Crystal structure of two covalent complexes of ribonuclease A with nucleosides. Biochemistry 29, 928-937. Pavlovsky, A. G., Borisova, S. N., Borosov, V. V., Antonov, I. V., and Karpeisky, M. Y. (1978). The structure of the complex of ribonuclease S with fluoride analogue of UpA at 2.5 ,~ resolution. FEBS Lett. 92, 258-262. Rasmussen, B. F., Stock, A. M., Ringe, D., and Petsko, G. A. (1992). Crystalline ribonuclease A loses function below the dynamical transition at 220 K. Nature (London) 357, 423-424. Richards, F. M., and Vithayathil, P. J. (1959). The preparation of subtilisin-modified ribonuclease and the separation of the peptide and protein components. J. Biol. Chem. 234, 1459-1465. Richards, F. M., and Wyckoff, H. W. (1971). Bovine pancreatic ribonuclease. In "The Enzymes" (P. D. Boyer, ed.), Vol. 4, pp. 647-806. Academic Press, New York. Richards, F. M., and Wyckoff, H. W. (1973). Ribonuclease-S. In "Atlas of Molecular Structures in Biology" (D. C. Phillips and F. M. Richards, eds.), Vol. 1, pp. 1-75. Oxford Univ. Press (Clarendon), Oxford. Stern, M. S., and Doscher, M. S. (1984). Aspartic acid-121 functions at the active site of bovine pancreatic ribonuclease. FEBS Lett. 171, 253-255.
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Svensson, L. A., Sjolin, L., Gilliland, G. L., Finzel, B. C., and Wlodawer, A. (1986). Multiple conformations of amino acid residues in ribonuclease A. Proteins: Struct. Funct. Genet. 1, 370-375. Svensson, L. A., Sjolin, L., Dill, J., and Gilliland, G. L. (1991). The conformational flexibility of surface residues of bovine ribonuclease A at 1.1 ~, resolution. In "Structure, Mechanism and Function of Ribonucleases" (C. M. Cuchillo, R. de Llorens, M. V. Nogues, and X. Pares, eds.), pp. 31-38. IBF Publications Universitat Autonoma, Barcelona. Taylor, H. C., Richardson, D. C., Richardson, J. S., Wlodawer, A., Komoriya, A., and Chaiken, I. M. (1981). "Active" conformation of an inactive semi-synthetic ribonuclease-S. J. Mol. Biol. 149, 313-317. Taylor, H. C., Komoriya, A., and Chaiken, I. M. (1985). Crystallographic structure of an active, sequence-engineered ribonuclease. Proc. Natl. Acad. Sci. U.S.A. 82, 6423-6426. Torii, K., Urata, Y., Iitaka, Y., Sawada, F., and Mitsui, Y. (1978). Crystal structure of monoclinic ribonuclease-S at 4 ,A resolution. The mode of binding of 4-thiouridylic acid and a fragment of folic acid, p-aminobenzoylglutamic acid. J. Biochem. (Tokyo) 83, 1239-1247. Usher, D. A., Richardson, D. I., and Eckstein, F. (1970). Absolute stereochemistry of the second step of ribonuclease action. Nature (London) 228, 663-665. Usher, D. A., Erenrich, E. S., and Eckstein, F. (1972). Geometry of the first step in the action of ribonuclease-A. Proc. Natl. Acad. Sci. U.S.A. 69, 115-118. Varadarajan, R., and Richards, F. M. (1992). Crystallographic structures of ribonuclease S variants with nonpolar substitution at position 13: Packing and cavities. Biochemistry 31, 12315-12327. Veenstra, T. D., and Lee, L. (1994). NMR study of the positions of His-12 and His-119 in the ribonuclease A-uridine vanadatc complex. Biophys. J. 67, 331-335. Weber, P. C., Salemme, F. R., Lin, S. H., Konishi, Y., and Sheraga, H. A. (1985). Preliminary crystallographic data for cross-linked (lysineT-lysincnl)-ribonuclease A. J. Mol. Biol. 181, 453. Wlodawer, A. (1985). Structure of bovine pancreatic ribonuclease by X-ray and neutron diffraction. In "Biological Macromolecules and Assemblies" (F. Jurnak and A. McPherson, eds.), Vol. 2, pp. 394-439. Wiley, New York. Wlodawer, A., and Sjolin, L. (1981 ). Orientation of histidine residues in RNase A: Neutron diffraction study. Proc. Natl. Acad. Sci. U.S.A. 78, 2853-2855. Wlodawer, A., and Sjolin, L. (1983). Application of joint neutron and X-ray refinement to the investigation of the structure of ribonuclease A at 2.0-,~, resolution. Biochemistry 22, 2720-2728. Wlodawer, A., Bott, R., and Sjolin, L. (1982). The refined crystal structure of ribonuclease A at 2.0 ,~ resolution. J. Biol. Chem. 257, 1325-1332. Wlodawer, A., Miller, M., and Sjolin, L. (1983). Active site of RNase: Neutron diffraction study of a complex with uridine vanadate, a transition-state analog. Proc. Natl. Acad. Sci. U.S.A. 80, 3628-3631. Wlodawer, A., Borkakoti, N., Moss, D. S., and Howlin, B. (1986). Comparison of two independently refined models of ribonuclease-A. Acta Crystallogr. B 42, 379-387. Wlodawer, A., Svensson, L. A., Sjolin, L., and Gilliland, G. L. (1988). Structure of phosphate-free ribonuclease A refined at 1.26 ,~. Biochemistry 27, 2705-2717. Wodak, S. Y., Liu, M. Y., and Wyckoff, H. W. (1977). The structure of cytidilyl-(2',5')adenosine when bound to pancreatic ribonuclease S. J. Mol. Biol. 116, 855-875.
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Wyckoff, H. W., Hardman, K. D., AlleweU, N. M., Inagami, T., Tsernoglou, D., Johnson, L. N., and Richards, F. M. (1967a). The structure of ribonuclease-S at 6 .~ resolution. J. Biol. Chem. 242, 3749-3753. Wyckoff, H. W., Hardman, K. D., Allewell, N. M., Inagami, T., Johnson, L. N., and Richards, F. M. (1967b). The structure of ribonuclease-S at 3.5 ,A resolution. J. Biol. Chem. 242, 3984-3988. Wyckoff, H. W., Tsernoglou, D., Hanson, A. W., Knox, J. R., Lee, B., and Richards, F. M. (1970). The three-dimensional structure of ribonuclease S. Interpretation of an electron density map at a nominal resolution of 2 ,~. J. Biol. Chem. 245, 305-328. Zegers, I., Maes, D., Dao-Thi, M.-H., Poortmans, F., Palmer, R., and Wyns, L. (1994). The structures of RNase A complexed with 3'-CMP and d(CpA): Active site conformation and conserved water molecules. Protein Sci. 3, 2322-2339.
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11 N M R Solution Structures of Ribonuclease A and Its Complexes with Mono- and Dinucleotides CARLOS GONZ/~LEZ, JORGE SANTORO, AND
MANUEL
RICO 1
Instituto de Estructura de la Materia CSIC 28006 Madrid, Spain
I. Introduction II. Structure of Bovine Pancreatic Ribonuclease A in Aqueous Solution A. Assignments and Secondary Structure B. Solution Structure C. The Active Site D. Side Chain Mobility E. Solution Studies on Glycoprotein Ribonuclease B III. Ligand Binding Studies: Complexes with Mono- and Dinucleotides A. Solution Structures of Complexes of RNase A with Pyrimidine Mononucleotides B. Solution Structures of Complexes of RNase A with 2',5'-CpA and 3',5'-d(CpA) IV. Structures of Other Animal Ribonucleases in Solution A. Bovine Seminal Ribonuclease B. Angiogenin References
I. I N T R O D U C T I O N
B o v i n e p a n c r e a t i c r i b o n u c l e a s e A ( R N a s e A , E C 3.127.5; 124 r e s i d u e s , M r 14,000) w a s t h e first e n z y m e t o b e e x a m i n e d b y N M R s p e c t r o s c o p y 1To whom correspondence should be addressed.
RIBONUCLEASES: STRUCTURES AND FUNCTIONS
343 Copyright 9 1997by AcademicPress, Inc. All rightsof reproductionin any form reserved.
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(Saunders et al. 1957). The resolution was such that the observed 40-MHz spectrum showed only four broad peaks, each one encompassing a great number of proton resonances from chemically different groups, from aromatics to aliphatics with no electronegative substituents. The use of NMR to obtain detailed information on protein structure in general had to wait until the middle of the 1960s, when crucial advances in instrumentation opened the way to an ever-increasing number of NMR studies. The studies then provided valuable information about many aspects of protein structure and function. Studies on RNase A structure and folding carried out in the 1960s and 1970s were based mainly on the H ~1 resonances from the four histidine residues whose assignment was reinvestigated and definitively fixed by Markley (1975). It is well known (Richards and Wyckoff, 1971) that two of these histidines, His12 and His-ll9, are of crucial functional significance in the processes of binding and catalysis. On the basis of the enzyme crystal structure and the changes observed in the titration behavior of the histidine H ~1 proton, model structures for the complexes of RNase A with cytidine mononucleotides were reported (Meadows et al., 1969). An effort to assign all aromatic resonances by using a variety of physical and chemical effectors was made by Lenstra et al. (1979). Important issues were addressed by using heteronuclei, e.g., 13C, to study pH-induced conformational transitions (Santoro et al., 1979), tautomeric states of the histidine residues (Waiters and Allerhand, 1980), and assignment and degree of exposure of tyrosine residues (Egan et al., 1978); ~SN was used to examine differences in the binding of nucleotides (Hahn et al., 1985), and 31p was used to determine the ionization state of the phosphate moeity (dianionic) and consequently those of His-12 and His-119 (both protonated) in mononucleotide complexes (Gorenstein et al., 1976; Dobson and Lian, 1987). Most of this early work has been reviewed in Jardetzky and Roberts (1981), Blackburn and Moore (1982), and Eftink and Biltonen (1987). In the second half of the 1980s, NMR was established as an alternative method for the determination of protein structures at the atomic level (Wtithrich, 1986), an advance that was possible thanks to improved developments in instrumentation and methodologies such as high-field (500-600 MHz) and multidimensional NMR spectroscopy. Up to that time, the principal source of structural information was from X-ray diffraction, and it had to be accepted that the protein structure in single crystals, as determined by this technique, was the correct model of a biologically active protein. The NMR technique, which provides struc-
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NMR Structures of RNase A and Its Complexes
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tural information under conditions very close to those in which proteins function, provided the means to check whether solution and crystal structures are identical and to determine the effect of packing on the protein surface. Bovine pancreatic RNase A is one of the best studied proteins by X-ray diffraction on single crystals (Wlodawer et al., 1988, and references therein). In fact, it was the third enzyme whose crystal structure was solved (Kartha et al., 1967). Determining the threedimensional structure of RNase A in aqueous solution (Santoro et al., 1993) was important in order to obtain meaningful conclusions about similarities and differences in the structure and dynamics of the solution and crystal states, as well as to provide an essential reference background for folding, dynamic, and ligand-induced conformational studies in solution. In this chapter, we focus on three-dimensional structures in solutions of RNase A and some of the complexes of RNase A with mono- and dinucleotides.
II. S T R U C T U R E OF BOVINE P A N C R E A T I C R I B O N U C L E A S E A IN A Q U E O U S S O L U T I O N
A. Assignments and Secondary Structure A first attempt to carry out the complete assignment of RNase A was made by Hahn and Rtiterjans (1985), who could assign 21 out of the 124 residues of the enzyme. The complete assignment was performed independently by Rico et al. (1989) and Robertson et al. (1989). Both assignments were carried out using standard homonuclear twoimensional (2D) methods (Wtithrich, 1986), and they are essentialy coincident. A comparison of the few discrepancies found was discussed in Rico et al. (1991) and an extension of the assignment to all residues with long side chains, as well as to primary amide protons of Asn and Gin residues, is given in Rico et al. (1993) and has been deposited in the BioMagRes data bank. The secondary structure in solution can be immediately delineated from the assigned spectra. Typical nuclear Overhauser effect (NOE) patterns for c~ helices were found in segments 4-11, 25-32, and 50-58. The /3 strands could also be identified by the N O E patterns and the low-field shifted H a resonances. The antiparallel alignment between the
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different strands can also be determined from H~-H ~ N O E connectivities. The secondary structure of RNase A in solution is very similar to the one observed in the crystal. A more detailed description is given in the next section in the light of the calculated three-dimensional structure.
B. Solution Structure
The first three-dimensional structure of RNase A in solution was determined on the basis of a small set of distance constraints obtained on a 360-MHz spectrometer (Rico et aL, 1991). A novel approach was used, based on a variation of the variable target function method. This approach makes use of an empirical correlation between the $ and $ angles observed in the Ramachandran diagram of known protein structures. The two torsion angles that define the conformation of the backbone in each residue are substituted for a single parameter. Consequently, the degrees of freedom during the calculation are significantly reduced, and a better convergence of the resulting structures is obtained. This approach allows the calculation of solution structures when the set of distance constraints is too small to use more standard methods. Although many structural features of the RNase A in solution could be determined from this preliminary work, we will discuss a more refined solution structure obtained by Santoro et aL (1993) with a larger set of distance constraints obtained on a 600-MHz spectrometer. Structural determination of highly refined protein structures in solution relies on the acquisition of a large and accurate set of structural constraints. If we consider N O E cross-correlations unambiguously assigned, the main impediment to obtaining reliable distance information from their intensities is the spin diffusion effect. Errors in the interproton distance produced by ignoring this effect can be neglected if the NOESY experiments are performed at short mixing times but, in these conditions, only the most intense cross-peaks are observed, resulting in a loss of valuable information. At longer mixing times, when more cross-peaks are present, the isolated spin-pair approximation leads to systematic errors (Borgias and James, 1988). This problem may be overcome by using a complete relaxation matrix analysis of the N O E S Y spectra, which enables the calculation of accurate interproton distances, because the spin diffusion is duly taken into account. This approach, implemented in the program I R M A (Boelens et aL, 1988, 1989; Koning et aL, 1990),
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which stands for Iterative Relaxation Matrix Approach, was used in the determination of interproton distances from NOE intensities obtained at different mixing times. Because this approximation relies on a quantitative integration of NOE intensities, those peaks that cannot be reliably integrated cannot be included in the analysis (overlapping peaks, peaks located near the diagonal or solvent signal, etc.). In these cases, the standard qualitative classification on strong, medium, and weak crosspeaks was used. Due to the large size of RNase A, serious overlapping effects were present in many regions of the spectra, preventing a reliable integration of many cross-peaks. The final set of NOEs comprised 464 quantitative interproton distance and 999 more approximate constraints. The latter includes only upper distance limits, whereas quantitative constraints include upper and lower limits. In addition, 42 dihedral angle constraints were obtained from the analysis of vicinal 1H-1H J coupling constants, determined from COSY (correlated spectroscopy) experiments. The complete set of constraints is deposited in the Brookhaven Protein Data Bank (NMR data entry 2AAS.MR). The calculation of the three-dimensional structure from the final set of distance and torsion angle constraints was carried out by using restrained molecular dynamics methods as implemented in the package GROMOS (van Gunsteren and Berendsen, 1987), and following an annealing strategy. Starting from the low-resolution structures obtained in previous determinations (Rico et al., 1991, 1993), 16 structures were calculated. The resulting structures display low values of the energy terms and satisfy the distance and torsion angle constraints. The use of the complete relaxation matrix approach allows a direct estimation of the quality of the NMR structures. Several reliability factors (R-factors) have been defined by comparing theoretical and experimental NOE intensities (Gonz~ilez et al., 1991). The two NMR R-factors most commonly used in the literature, the crystallographic-like and the sixth-root R-factor, give remarkably low values (0.47 and 0.16) for the resulting structures of RNase A. The coordinates of the structures are deposited in the Brookhaven Protein Data Bank under filename 2AAS.MR. Contrary to crystallographic structural models, the aim of the computational methods for structural determination in solution is not to obtain a single structure that best satisfies the experimental data. NMR structural determinations, instead, pursue the exploration of all the conformational space consistent with these experimental constraints. The extension of the conformational space depends on the number and quality of the structural restraints obtained from the NMR data, and is normally esti-
348
Carlos Gonz~ilez et al.
mated by the root-mean-square deviation (RMSD) of the atomic coordinates in different resulting structures. The large and accurate set of distance constraints obtained for RNase A gave rise to a remarkably low average pairwise RMSD for the final 16 structures (0.66/k for the backbone and 1.68 ,~ for all heavy atoms, including side chains). The atomic RMSD is even lower (0.4 A) when only the elements of welldefined secondary structure are included. Figure 1 (top) displays the ensemble of the 16 final structures. The high definition of the calculated structures is not only confined to the backbone. Many of the side chains also exhibit well-defined X~ values, with 64 residues having a RMSD of the torsion angle below 10~ The resulting solution structure of RNase A is very similar to the crystal structure, (Fig. 1, bottom). The RMSD of the backbone atoms between the average solution structure and the crystallographic one is 0.92/k, slightly higher than the RMSD between the 16 structures calculated in solution (0.66 ,~). The overall structure consists of two/3 sheets surrounded by three a helices. These elements of secondary structure, which were previously outlined on the basis of the observed patterns of NOE connectivities (Rico et aL, 1989), can be more clearly defined from an analysis of average backbone dihedral angles and hydrogen bond patterns. In Fig. 2 the backbone-backbone hydrogen bonds observed in the solution structures are shown. Figure 2 includes also information on the exchange rates of individual amide protons with solvent deuterons. A discussion of exchange protection factors as related to structural and dynamics features was given in Santoro et al. (1993) and is the subject of a separate review (Neira and Rico, 1996). The helix between residues 4-13 (helix I) is very well defined and regular. Hydrogen bonds of the type COi-NHi+4 are formed throughout the helix, and the ~band q~angles are characteristic of an a helix, except for the ~bvalue in His-12 (-120~ Contrary to helix I, the second helix, comprising residues 24-33 (helix II), is highly irregular. This helix begins with a CO/-NHi+3 hydrogen bond between Asn-24 and Asn-27 and the CO of this last residue does not appear to be involved in any hydrogen bond. At the C end of this helix, Asn-34 is found in the aL helix conformation, reversing the direction of the polypeptide chain. The third helix extends from Leu-51 to Ser-59, with dihedral angles ~b and qJ tightly constrained. It begins with an a-helix turn and continues with two turns of a 3~0 helix. More than one-third of the RNase A residues are forming part of the /3 strands. Seven individual strands can be sketched comprising residues Pro-42 to His-48 (/3~), Lys-61 to Ala-64 (/32), Asn-71 to Gin-74 (/33), Met-
11
NMR Structures of RNase A and Its Complexes
349
Fig. 1. Top: Stereoscopic view of the superposition of the 16 obtained solution structures. Bottom: Stereoscopic view of the superposition of two conformations of RNase A. (a) Solution average structure of the 16 obtained structures (thick line) and (b) X-ray diffraction structure in the crystal state (thin line).
79 to Thr-87 (~4), Ala-96 to Lys-104 (/35), His-105 to G l u - l l l (/36), and V a l - l l 6 to Val-124 (/37). These strands are arranged in an antiparallel way to form two /3 sheets displaying the patterns /31(a)(/34)(a)/35 and
350
Carlos Gonztilez et al.
Fig. 2. Scheme of hydrogen bonds involving the main chain atoms and exchanging properties of amide protons. Solid-line arrows correspond to hydrogen bonds detected in 15 or 16 solution structures and dash-line arrows correspond to those observed in at least 10 of them; arrows point toward acceptor atoms. Black circles correspond to amide protons with an exchange rate (k) lower than 1.5 x 10 -4 min -t, open ones to those with a k greater than 1.5 x 10 2, and shaded ones to those with a k between the two above limits (pH 4, 35~
11
NMR Structures of RNase A and Its Complexes
351
/32(a)f13(a)/36(a)/~7 and show the characteristic right-handed twist. The first sheet is very regular and all standard hydrogen bonds are observed. However, the second one shows two classic/3 bulges in the/37 strand: one involving Val-ll8 and His-ll9 opposite Ala-109, and a second one involving Asp-121 and Ala-122 opposite Ile-107. Residues Cys-65, Gly68, and Gin-69, adjacent to/32 and/~3 strands, have dihedral angles and hydrogen bonding patterns corresponding to a G1 bulge, so that they may be considered as forming part of this second sheet. Several turns are found in RNase A, some of which are distorted to various degrees from the standard geometries (Richardson, 1981; Rose et al., 1985). In the region 14-23, which connects the two first helices, a hydrogen bond between Asp-14 (CO) and Thr-17 (NH) is found, with backbone angles for Ser-15 and Ser-16 corresponding to a type III/3 turn. This turn is followed by an abrupt change in the polypeptide chain at residue 18 and by a distorted hydrogen-bonded type I/3 turn involving residues 22 to 25, just at the beginning of the second helix. Hydrogen bonds between Asp-14 (NH) and Val-47 (CO) and between Val-47 (NH) and His-12 (CO) anchor the "S peptide" to the "S protein" region. The region 33-40 displays the largest dispersion values of the backbone angles. Most of the loss of definition in the atomic coordinates of these residues is due to an increase in the angular RMS difference for the qJ angle of Arg-33, 4) and qJ angles of Asn-34, the q~ angle of Lys-37, and the ~bangle of Asp-38, suggesting the presence of hinge motions involving the peptide planes Arg33-Asn 34 and Lys37-Arg3s rather than an increase in local disorder. The relative high RMSD between the backbone heavy atoms precludes an accurate classification of turns in this region. Residues 65-68 form a well-defined type I /3 turn with the characteristic COiNHi+ 3 hydrogen bond. Turns involving cis-proline are found in the segment 91-94, which forms a type Via/3 turn, and residues 112-115, which form a type VIb/3 turn with no COi+3 hydrogen bond. Finally, the region 75-78 forms an unclassified loop connecting strands/33 and/34.
C. The Active Site
Most of the side chains of residues involved in the active site are welldefined and present X values in good agreement with the crystallographic structure. Only Lys-41 and Gln-ll display a slight increased disorder toward the end of their side chains. The side chain of His-ll9 deserves
352
Carlos Gonzdiez et al.
special attention because it has been a subject of controversy in different crystallographic studies. Early works on RNase S crystals showed the location of this side chain undefined (Wyckoff et al., 1970). Borkakoti et al. (1982) found two alternative sites, A and B, with occupancies of 0.8 and 0.2, respectively. In contrast, only position A prevails in the more refined crystal structure obtained by Wlodawer (1984). The solution studies (Rico et al., 1991, 1993; Santoro et al. 1993) indicate that a dynamic equilibrium exists between two allowed positions for the His119 side chain. This equilibrium is suggested by the conflicting NOEs observed between protons of His-ll9 with protons of Val-ll8 and Asp121. No single conformation satisfying all the constraints could be found. After examination of appropriate molecular models, the conflicting distance restraints could be sorted out in two self-consistent sets, which were used to calculate the two side chain positions. The resulting structures from each set of constraints present 2'1 and X2 angles of - 5 2 ~ and - 6 2 ~ (position B), and 167 ~ and 82 ~ (position A) (see Fig. 3). From the relative N O E intensities, a population of 0.8 and 0.2 for each position could be estimated. These two positions coincide practically with sites B and A described by Borkakoti et al. (1982). In the conditions of the NMR study, the major conformer observed in solution is that corresponding to the site with minor occupancy in the crystal. It has been shown, however, that absence of phosphate or high pH favors the major conformer observed in the crystal (Rico et al., 1991).
D. Side Chain Mobility Although conflicting N O E restraints represent the most direct evidence of multiple conformations in rapid exchange, they can be observed only under very favorable conditions (when the population of the conformer is high enough to detect the corresponding NOE). In RNase A, the only cases observed were His-ll9 (see Section II.C) and His-105, for which the conflicting NOEs can only be satisfied assuming two conformations for the X2 angle of its side chain. In addition, two other experimental indications of side chain mobility can be obtained from homonuclear NMR data: first, the lack of enough NOE constraints to keep the side chain in a well-defined conformation during the restrained molecular dynamics calculation, and, second, intermediate values of the 3j~o and 3j~o, vicinal coupling constant. According
8
Fig. 3. Top: Detail of the active center region showing the most populated His-f19 side chain conformation in solution. Bottom: Average solution structure showing the alternative position of the His-ll9 side chain (broken). The position of the phosphate phosphorus (P) atom in the crystal is indicated.
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Carlos Gonz~ilez et al.
to the Karplus equation, at least one of the two 3j~ coupling constants must be lower than 5 Hz in any of the three most populated alternate conformations of the X1 torsion angle. In the g+ conformation both coupling constants are lower than 5 Hz, and in the other two (g- and t) one constant is lower than 5 Hz and the other larger than 9 Hz (Case et al., 1994). Intermediate values in both constants is possible only when an equilibrium between several exchanging conformations is present. In the case of RNase A, side chains were classified as having one single staggered conformation if the two following requirements were fulfilled. First, X1 is found in the same conformational region for at least 14 of the 16 resulting structures, and either both 3j values were lower than 5 Hz (X~ = 60~ or each differed from the other by more than 5 Hz (Xl = -60~ or 180~ For further torsions in the side chain, only the first criterion was considered. 58 residues show a well-defined gl conformation, and 29 of them are long side chain residues with a well-defined conformation along the entire side chain. Side chain mobility can be related with the percent of solvent accessibility (Richards, 1977), a correlation that is shown in Fig. 4. Residues with well-defined side chain conformations in all their length appear in the interior of the protein (Fig. 4a) whereas those underlying motional averaging (Fig. 4c) are located near the protein surface. Side chain mobility in solution can be compared with the reported highly refined RNase A crystal structure (Svensson et al., 1986, 1991; Kuriyan et al., 1991), where modeling of a number of residues with discrete multiple side chain conformations has been carried out. From 13 residues modeled as mobile in the crystal (Svensson et al., 1986) 9 are found with disordered side chains also in solution. Exceptions are Gin-11, Ser-50, Asn-67, and Lys-104, which show well-defined side chain conformations in solution. Differences found in Gin-11 might reflect the fact that crystal data refer to the phosphate-free RNase A, whereas the solution data belong to the phosphate-bound enzyme (Rico et al., 1989). Removal of phosphate from the active site has been proposed as the cause for the appearance of two conformations for this residue in the crystal phosphate-bound enzyme (Svensson et al., 1986). The hydroxylic Ser-50 (OH) exchanges slowly enough to be observed in H20 solution (6 9.07 ppm) (Rico et al., 1993). Both its decreased exchange rate and chemical shift value suggest its involvement in a hydrogen bond. This bond is in fact detected in 10 out of the 16 converged solution structures, having as acceptor a carboxylate oxygen of Asp-53. The different behavior in mobility of the side chains of Asn-67 and Lys-104 will need further
15
h8l
lo -147 541
89
. ,,,,.
10-
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o d
5 -
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131 39 10 1 50 69 85 17R
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53
43 61
8q
66 91
98
77 94
! 10
20
30
40
50
60
70
80
90
100
% solvent accessibility of residue Fig. 4. Side chain mobility against percentage residue solvent exposure. (a) Residues with a well-defined side chain conformation in all their length, complying with both RMS deviations for torsional angles in the converged structures and 3J, 0 and 3j~, criteria (see text). (b) Residues with a n o n r a n d o m side chain conformation complying with one of the above criteria and showing at least a well-defined conformation around X1. (c) Residues with a motional averaged or multiple side chain conformations.
356
Carlos Gonz~ilez et al.
clarification. In addition to the residues modeled with discrete multiple conformations in the crystal, clear evidence of side chain disorder in residues 1, 7, 15, 18, 24, 37, 59, 66, 94, 123, and 124 was found in the solution studies. Among these, residues 7, 15, 18, 59, 66, and 123 are described in the two more recent crystallographic studies (Svensson et al., 1991; Kuriyan et al., 1991) as showing different side chain conformations.
E. Solution Studies on Glycoprotein Ribonuclease B
Bovine pancreatic RNase is in fact a mixture of unglycosylated RNase A and a collection of glycoforms known as RNase B. This form of the enzyme has five glycoforms consisting of (Man)5(GlcNAc): to (Man)9 (GIcNAc)2 (Man and GlcNAc refer to mannose and N-acetylglucosamine, respectively) at the single glycosylation site, Asn-34. At present, little is understood about the role of carbohydrates on protein structure, dynamics, and function. Effects on protein solubility, protection from proteolytic action, thermal stability, or modulation of enzymatic activity have been proposed. RNase B has been used as a model glycoprotein to obtain insights into these problems. Early studies of RNase B by using natural abundance ~3C NMR (Betman et al., 1981) were first used to determine the oligosaccharide primary structure by comparing the spectra of the enzyme with those of model carbohydrates. Once the major component was identified as (Man)5 (GIcNAc)2, ~3C chemical shifts, spin-lattice relaxation times, nuclear Overhauser enhancements, and linewidths of the oligosaccharide were examined in the free and bound forms. No differences were found on shifts, indicating that the average structure of the oligosaccharide in its free form is not affected by its binding to RNase A. The terminal mannose residues of RNase B exhibited fast internal motion as in the model compounds, as deduced for the remaining NMR measurements. Also, the comparison of chemical shifts of the quaternary carbons of RNase B and RNase A strongly suggests that the carbohydrate region has a negligible effect on the conformation of the enzyme. Just about the same conclusions were drawn from an X-ray diffraction study of RNase B at 2.5 resolution (Williams et al., 1987). Crystals contained two independent molecules of RNase B as the asymmetric unit. The polypeptide backbones of the two molecules in the asymmetric unit were found to be
11
NMR Structures of RNase A and Its Complexes
357
statistically identical and their differences from RNase A to be statistically insignificant. Most of the proton NMR resonances of RNase B have been assigned (Joao et al., 1992) by using standard techniques and by comparison with published assignments of RNase A (Rico et al., 1989; Robertson et al., 1989). A comparison of the ~H NMR spectra of RNase A and RNase B shows that glycosylation of the enzyme has little overall effect on the conformation of the protein in solution. By measuring individual amide H/D exchange rates, some differences could be appreciated. In general, exchange rates are lower for the glycosylated protein by a factor of 1.5to 6.0-fold. The presence of the oligosaccharide is observed to protect against hydrogen exchange not only residues close to the site of glycosylation but also a large number of residues throughout the protein. These data can be explained in terms of enhancing the stability of the native conformation against locally unfolded forms, where exchange occurs. This reduction in dynamic flexibility would extend throughout the entire structure of the enzyme. The exchange properties of the individual glycoforms with one (RNase B-Manl) and five (RNase B-Mans) mannoses (Joao and Dwek, 1993) show also a general enhancement of the protection against exchange as compared to RNase A. Furthermore, these individual glycoforms show additional protection when compared to RNase B (the natural mixture), including amide protons at and around the glycosylation site (residues 32, 34, and 35) as well as residues 10-13 in helix I. This fact may be rationalized by accepting that the glycoforms with a higher content of mannose (Man6 to Mang) must necessarily have higher rates of H/D exchange. The additional protection of the residues, close in sequence (32-35) and spatially (10-13), has no obvious explanation. It could arise from a special conformation for Man5 in which there was an increased steric hindrance between the sugar and the protein, thus reducing solvent accessibility. The overall increase in stability of the glycosylated form of the enzyme is paralleled by small increases in the energy of unfolding by guanidinium chloride (GndCl) (Puett, 1973) or thermal unfolding (Joao and Dwek, 1993), which can be evaluated at 5 kJ/mol. The RNase glycoforms were equally more resistant to pronase than was RNase A (Rudd et al., 1994), suggesting that, in addition to the increase in stability, the oligosaccharide may protect sterically one or more sites susceptible to be cleaved.
358
Carlos Gonz~ilez et al.
III. L I G A N D B I N D I N G STUDIES: C O M P L E X E S WITH MONO- AND DINUCLEOTIDES
The catalytic mechanism of RNase A has been studied extensively (Blackburn and Moore, 1982). It consists of two subsequent steps: the first reaction is a transesterification, which results in the cleavage of the P-O-5' bond and the formation of a 2',3'-cyclic nucleotide at the 3' end of a pyrimidine; the second stage is the hydrolysis of the cyclic nucleotide. The amino acids involved in the catalytic process were determined by chemical modification studies and pH dependence of the enzymatic activity (Blackburn and Moore, 1982). A more detailed description of the geometry of the active site can be achieved by studying the complexes between RNase A and several substrate analogs. Complexes of RNase A with some of the mono- and dinucleotides that are analyzed below have been previously studied by X-ray crystallography (Wodak et al., 1977; Howlin et al., 1987; Lisgarten et al., 1993; Zegers et al., 1994). In early 1D NMR studies, molecular conformations were obtained mainly by monitoring the resonances of histidine H ~2 and H *~ protons of the protein, and the H-5 and Ho6 protons of pyrimidine nucleotides. More recently, two-dimensional NMR was applied to study the 2'- and 3'-pyrimidine nucleotide complexes by Hahn and Rtiterjans (1985). A partial assignment of 21 amino acid residues was used to locate conformational changes in the protein associated with the base type (cytidine or uridine) and the position of the phosphate group of the nucleotide. Our group has carried out an extensive solution study of the complexes between RNase A and four mononucleotides (2'-CMP, 2'-UMP, 3'-CMP, and 3'-UMP) and two dinucleotides [2'-5'-CpA and 3',5'-d(CpA)] by using NMR and restrained molecular dynamics methods (Toiron et al., 1996). Here, a summary of the main results is presented. Standard 2D NMR methodology was used to assign the proton spectra (Wtithrich, 1986). The assignment was greatly facilitated by the previous assignment of the free enzyme (Rico et al., 1989; Robertson et al., 1989; see Section II). Due to serious signal overlapping, the spectra were recorded over a range of temperature and pH. Although the pH for optimal binding is 5.5, the best quality spectra were obtained at pH 4.0. Most of the experimental constraints were derived from spectra acquired in these conditions (pH 4.0, T 380 K), which are identical to those used for the free enzyme. Because the exchange between the free and bound
11
N M R Structures of RNase A and Its Complexes
359
states of mono- and dinucleotides is rapid on the N M R time scale, caution must be taken to extract the intraprotein constraints without any contamination from NOEs belonging to the free enzyme. The same applies to the intranucleotide constraints. Therefore, samples with different stoichiometry were prepared to achieve either complete protein saturation or complete inhibitor saturation. Distance constraints obtained from the analysis of the N O E S Y spectra were subsequently used in a restrained molecular dynamics procedure. The protocol was analogous to the one used for the calculation of the free enzyme. In all the complexes studied the significant chemical shift changes on complex formation are located in well-defined regions of the protein sequence (see Section Ill,A). Because the chemical shift is the N M R parameter most sensitive to structural information, we can conclude that the structure of the protein in the complex is identical to that in the free enzyme except in those regions where a change in the proton chemical shifts was detected. This chemical shift information was implemented in our molecular dynamics calculation by constraining the atomic position in all the regions of the protein where there is no significant variation in chemical shifts. Only the residues with a chemical shift change larger than a certain threshold were allowed to move during the simulation. Thus, the backbone of residues with H N shift deviation lower than 0.1 ppm or H a chemical shift variation lower than 0.05 ppm was kept fixed. A similar criterion was used for the side chains. In order to obtain a starting structure to be used in the molecular dynamics refinement, a preliminary docking of the different inhibitors with the average structure of the free enzyme was carried out. The docking was performed manually with the computer modeling package Insight II. The inhibitor was placed in the interior of the active site in a conformation that roughly satisfies the experimental distance constraints.
A. Solution Structures of Complexes of RNase A with Pyrimidine Mononucleotides As mentioned previously, most of the chemical shift variations on mononucleotide complex formation are located in very restricted regions of the protein sequence (see Figs. 5 and 6). However, the areas affected by the inhibitor binding display large changes in chemical shifts for the H N and H a protons; these areas are the regions 11-12, 41-45, and 119-
AG H~(ppm) 0.4 0.2
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20
30
40
50
60
70
80
90
100
110
120
Sequence Fig. 5. C h e m i c a l shift differences ( p p m ) b e t w e e n the H" r e s o n a n c e s of the different residues in the complexes (1 : l ) with 2 ' - C M P , 2 ' - U M P , 3 ' - C M P , 3 ' - U M P , 2 ' , 5 ' - C p A , and 3 ' , 5 ' - d ( C p a ) and those of free R N a s e A (pH 4, 35~ Differences o b s e r v e d only in the 1:1 c o m p l e x are indicated by asterisks. 360
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~ 80
~ 90
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120
Fig. 6. Chemical shift differences (ppm) between the NH resonances of the different residues in the complexes (1:1) with 2'-CMP, 2'-UMP, 3'-CMP, 3'-UMP, 2',5'-CpA, and 3',5'-d(CpA) and those of free RNase A (pH 4, 35~ #, Unobserved; +, chemical shift measured at pH lower than 5.5. 361
362
Carlos Gonz~ilez et al.
121, which are considered as part of the active site. In addition, the regions 83-85 and 106-108, in the B strand adjacent to the active center, are also affected. In general, chemical shift variations are larger in the CMP complexes than in the U M P complexes. On the other hand, binding of 2'-phosphate nucleotides produces greater changes than does binding of 3'-phosphate nucleotides. Because there is no global change in the structure of the protein, we focused the analysis of the N O E S Y spectra on the differences with the free enzyme more than on the extraction of a complete set of constraints. The pattern of intramolecular N O E restraints in all the complexes is rather similar to that found for free enzyme. Only some differences of NOEs corresponding to the side chain protons were observed. Intramolecular constraints were derived from either protein-saturated or nucleotide-saturated samples. Special attention was paid to detect intermolecular NOEs between the enzyme and the mononucleotide, which were derived from 1:1 samples. The observed intermolecular contacts are shown in Table I. Most of the N O E s involving the base protons (H-5 and H-6) are common to cytidine and uridine rings. Contacts between H - I ' in the sugar ring and the side chain protons of the His-12, Lys-41, and Val 43 are also present in the four complexes. In the 2' complexes, additional NOEs of H - I ' with H" of Asn-44 and H N of Thr45 are observed. The NOEs involving the sugar H-2' resonance in the 3' complexes are not observed in the 2' complexes. On the other hand, NOEs involving H-3' are observed only in the 2' complexes. This fact may not be the result of any structural difference, but a consequence of signal broadening of the H-2' and H-3' resonances in the 2' and 3' complexes, respectively, produced by intermediate exchange between the free and the bound forms. Most of the intramolecular NOEs detected in the free enzyme are also found in the complexes. Particularly interesting is the presence of conflicting NOEs in the side chain of His-ll9. The H s2 proton of His119 shows cross-peaks with protons of Asp-121 and Val-ll8. As in the free enzyme, no position of the histidine ring accounts for all the detected NOEs, and two conformations of the side chain of H i s - l l 9 in rapid exchange must be assumed to satisfy all the constraints. The structure calculations were carried out with the constraints between His-ll9 and Asp-121, which correspond to position A for the side chain of His-ll9 in the free enzyme. The resulting structures obtained from the restrained molecular dynamics protocol present a very low atomic RMSD. Figure 7 shows a
11
NMR Structures of RNase A and Its Complexes
363
superposition of eight structures of the RNase A - 3 ' - C M P complex. Main chain and side chain atom groups in the active site of the four complexes are well-defined, and are shown in Figs. 8 and 9. Some of the hydrogen bond interactions between the protein and its inhibitors are listed in Table II. The structure of the active site is very similar in the four complexes. His-12 maintains the same position in the free enzyme, H 81 forming a hydrogen bond with the carbonyl group of Thr-45. Proton H ~2 in the opposite edge of the imidazole ring forms a hydrogen bond with one of the phosphate oxygens or, alternatively, with 0-2'. Thr-45 is also involved in the binding process, forming two hydrogen bonds. The NH a m i d e proton is hydrogen bonded to the carbonyl oxygen in position 2 of the base in both cytidine and uridine complexes. The hydroxylic oxygen in the side chain of Thr-45 interacts with the electronegative N-3 base atom, acting as a hydrogen donor in the case of the cytidine complexes or as an acceptor in the uridine complexes. At the other side of the active site, the H ~1 proton in the side chain of His-119 forms a hydrogen bond with one of the phosphate oxygens. In the 2'-CMP complex, the H ~1 proton forms hydrogen bonds either with a phosphate oxygen or with 0-3'. The amide proton of Phe-120 is also bound to an oxygen phosphate in all four complexes. All these interactions are also observed in the crystallographic structures of the cytidine complexes (Lisgarten et al., 1993; Zegers et al., 1994). Although N O E cross-correlations between side chain protons of Lys41 and ribose protons are observed in all the complexes, no hydrogen bond involving protons of its amino group results in the final structures. This may be due to the lack of a number of key constraints that would be sufficient to define completely the conformation of this side chain. In any case, the side chain is located closer to the phosphate than in the initial structure. Gin-l l, Lys-66, and Ser-123 are also involved in the binding process but no direct hydrogen bond interaction is observed in the final structures. In the crystallographic structures available, these interactions are often mediated by a water molecule. The chemical shifts of the side chain protons of Gln-11 and Ser-123 change on complex formation, giving additional evidence that these residues may play a role in the binding process. The chemical shifts of the methyl protons of Val43 are largely affected by complex formation, and also they present several N O E cross-peaks with the inhibitor protons. In the calculated structures, this residue is in close contact with the ribose ring. The conformation of the inhibitor molecules is anti in all the complexes, with
TABLE I
Upper Limit Constraints (A) between Pairs of Protons as Derived from Qualitative Evaluation of NOE Cross-Correlations" Proton pair HE' HE'
HE' HE' HE2 HE" c12 (22
HO H@ c12 H" HN HE1 HE1 HE'
12 12 12 41 41 41 43 43 43 43 43 44 45 119 119 119
H-1' H-2'1 H-2'2 H-1' H-1' H-4' H-6 H-1' H-1' H-5 H-5 H-t' H-1' H-3' H-5'1 H-5'2
2'-CMP CYt CYt CYt CYt CYt CYt CYt CYt CYt CYt CYt CYt CYt CYt CYt CYt
3.0 -
2'-UMP
3'-CMP
3'-UMP
2',5'-CpA
4.0
4.0 3.0
5.0 ps 4.9
ps 4.9 5.0 6.0
ps 5.9 6.0 5.0
3.0 4.0 ps 3.9 3.0 ps 5.9 6.2 5.0
4.0
-
3.0 4.0 ps 3.9
6.0 6.0
5.2
4.0 5.0
5.0 5.2 4.5 5.0 5.0
3.0
ps 5.9 ps 4.9 ps 7.2 ps 7.2 5.0 5.0 ps 5.2 5.0 5.0 4.0
5.0 5.2 5.0 5.0 4.0 5.0 5.0
5.0 5.0
5.0
3',5'-d(CpA)
ps 5.9
HE' HE' H6" H" CB CB CB H" HY2 H7' HY2 C2 C l
c12 C2 cy2
He HE' a
119 119 120 122 122 4 109 111 111 111 111 118 118 118 118 118 119 119
H-2'1 H-2'2 H-5 H-5 H-5 H-1' H-2 H-2 H-2 H-2 H-2 H-1' H-2 H-2 H-8 H-2 H-1' H-8
CYt CYt CYt CYt CYt Ade Ade Ade Ade Ade Ade Ade Ade Ade Ade Ade Ade Ade
Restriction refers to the pseudoatom (ps)
4.0
-
-
ps 1.2 3.0
4.5
6.0 4.0 5.0 5.0 5.0 5.0 6.0 6.0 4.0 4.0 5.0
5.0 5.0 3.5 5.0
ps 4.9
6.0 4.0 5.0 5.0
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Carlos Gonziilez et al.
111
Fig. 7. Stereoscopic view of the superposition of eight converged structures of the complex RNase A-3'-CMP.
glycosidic torsion angles X ( 0 - 4 ' - C - 1 ' - N - 1 ' - C - 2 ) ranging from - 1 1 7 (2'-UMP) to -141 (3'-UMP). The pseudorotation phase angles P angles of the riboses are in the general N domain for the 2' complexes (P = 17~ for 2'-CMP and P = 29 ~ for 2'-UMP), and in the S domain for the 3' complexes (P = 147 ~ for 3'-CMP and P = 143 ~ for 3'-UMP). N and S domains correspond to conformations close to C-3'-endo and C-2'-endo, respectively. Although an anti conformation for the glycosidic angle agrees with the conformation observed in the crystallographic structures, pseudorotation phase angles in the S domain are clearly in contradiction with the crystallographic data.
B. Solution Structures of Complexes of RNase A with 2',5'-CpA and 3',5'd(CpA) Similar to the mononucleotide complexes, chemical shift variations on the binding of the 2',5'-CpA and 3',5'-d(CpA) are restricted to specific regions of the sequence. Hence, the complex formation does not affect the global structure of the enzyme. The same areas affected by the mononucleotide binding display the larger chemical shift changes. In addition, some other residues that were not affected in the mononucleo-
11
N M R Structures of RNase A and Its Complexes
367
12
12 12
12 6
6
1 6
7 10
4~
Fig. 8. Stereoscopic view of the average solution structure of the complexes RNase A-2'-CMP (top) and RNase A-2'-UMP (bottom).
tide case show large chemical shift deviations. In the case of 2',5'-CpA, the residues affected are 19-20 and 118-119. In 3'-5'-d(CpA), residue 20 is not affected, but the H N resonances of residues 67, 80, and 122 are largely shifted. It must be pointed out that H N resonances of Thr-45 (in both cases) and Phe-120 [in 3',5'-d(CpA)], which are most greatly affected by the mononucleotide binding, could not be assigned in the spectra, probably due to broadening effects in these specific resonances. The same conclusion obtained in the mononucleotide case is valid here: inhibitor binding does not affect the global structure of the enzyme. Therefore, we will focus the analysis of the N O E S Y spectra on their
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Carlos Gonz~ilez et al.
1 12
--
6 -
12 10
69
"k~
1
I
-
Fig. 9. Stereoscopic view of the average solution structure of the complexes RNase A - 3 ' - C M P (top) and RNase A - 3 ' - U M P (bottom).
differences as compared with the free enzyme. Intermolecular contacts between RNase A and 2',5'-CpA and 3',5'-d(CpA) are shown in Table I. Most of the contacts involving the cytidine in the 2'-5'-CpA complex are common to those found in the 2'-mononucleotide complexes. In the case of 3',5'-d(CpA), some differences with the intermolecular NOEs of the 3'-mononucleotide complexes are observed. Instead of the contact between proton H-2' of the cytidine and H ~ of His-12, as observed in the mononucleotide complexes, a NOE between H-2'2 and the histidine H ~ was detected. On the other hand, the sugar proton H-I' presents in the 2',5'-CpA complex a strong N O E with the side-chain protons H ~ of Lys-41, which is weaker in the 3'-5'-d(CpA) complex. The chemical shift
T A B L E II Hydrogen Bonds between A t o m s of RNase A and Ligand a
Amino acid
Atom
2'-CMP
2'-UMP
3'-CMP
His-12
H e2
O-2'-O-1P
O-2'-O-1P
O-2'-O-1W
Thr-45 Thr-45 Thr-45 Gin-11 His-119 Phe-120 Phe-120 Ser-123
O y2
H-42 N-3 0-2
N-3 H-3
O-2P O-1P
H ~1 H TM H g2 H ~1 HN O H TM
3'-UMP O-2'-O-1P
N-3 H-3
0-2
N-3 0-2
0-2
O-2P O-1P
O-2P O-1P
O-2P O-1P
0-4
0-4
a The bond distances are less than 3.5 ~,. O-1P and O-2P refer to oxygens bound to the phosphorus atom.
2',5'-CpA O-1P-O-2P (30%) H-42 Cyt N-3 0-2 Ade 0 - 5 ' - 0 - 4 ' O-2P Cyt O-3'H
3',5'-d(CpA)
H-42 Cyt N-3 0-2 Cyt 0 - 3 ' Ade 0 - 5 ' - 0 - 4 ' O-2P (20%)
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Carlos Gonz~ilez et al.
dispersion of the two H ~ protons of Lys-41 is lower in the 3' ,5 '-d(CpA) than in the 2',5'-CpA complex. Although the adenine is located on the exterior of the active site cleft, it also presents a number of N O E contacts with several residues of the protein. In both dinucleotide complexes, contacts with residues 4, 111, and 118, and 119 are observed. In the case of 2',5'-CpA, a larger set of NOEs with G l n - l l and Val-ll8 is detected, as well as a N O E between the adenine H-2 proton and the methyl of Ala-109, not observed in the 3',5'-d(CpA) complex. The most important difference between the intramolecular NOEs in the mono- and dinucleotide complexes is the lack of N O E contacts between the side chain protons of His-ll9 and Val-ll8. In the free enzyme and the mononucleotide complexes, conflicting NOEs of His119 with Asp-121 and Val-ll8 were observed. Only the NOEs with Asp-121 remain in the dinucleotide complexes. As a consequence, the resulting structures from restrained molecular dynamics calculations correspond exclusively to conformation A in the free enzyme. Overall, the calculated structures are well defined. The average structures of the two complexes are displayed in Fig. 10. The inhibitor structures are well defined in the bases and sugar moieties of both nucleotides, but not in the connecting regions between the two sugars (the phosphodiester group), due to the absence of NOEs and consequently of experimental constraints in this part of the molecule. Most of the hydrogen bond interactions observed in the mononucleotide complexes are also found in the dinucleotide complexes. However, the interaction between one of the phosphate oxygens and the H ~2proton of His-12 is not observed in all the final structures. Likewise, the population of the hydrogen bond between the amide proton of Phe-120 and one of the phosphate oxygens is lower in the 3'-5'-d(CpA) complex. The H ~ proton of His-ll9 forms a hydrogen bond with the 0 - 5 ' of the adenine, bonded to the phosphorous atom, instead of the O-1P or 0 - 3 ' oxygen. These differences affect only the relative position of the oxygens in the phosphate group. It must be emphasized that no direct experimental information on the phosphate moiety is available from N M R data. The relative position of the oxygen in the phosphate group is mainly driven by the force field potential in the molecular dynamics simulation. The base of the adenosyl residue is involved in several transient hydrogen bonds with Asn-67, Gln-69, and Asn-71. None of these bonds has populations larger than 25% in the final set of calculated structures. It must be pointed out that the position of the adenine base is almost identical in
11
NMR Structures of RNase A and Its Complexes
371
12
12
12
12 69
69
6
,-,r
1
/',~.\
Fig. 10. Stereoscopic view of the average solution structure of the complexes RNase A-2'-,5'-CpA (top) and RNase A-3',5'-d(CpA) (bottom).
all the structures. The different hydrogen bonds arise from the various positions of the side chains of residues 67, 69, and 71, which are very flexible in the solution structures. Glycosidic torsion angles in the 2 ' , 5 ' - C p A - R N a s e A complex are in the anti conformation in both nucleosides ( - 1 2 3 ~ for the cytosine and - 7 7 ~ for the adenosine). The sugar conformations are in the general N domain, with pseudorotation phase angles of 80 ~ (cytosine) and 42 ~ (adenine), corresponding to the Co4'-exo region. Both bases are also in the anti conformation in the 3',5'-d(CpA) complex, with values for the glycosidic angle of - 1 4 2 ~ for the cytosine and - 1 0 7 ~ for the adenosine.
372
Carlos Gonz~ilez et al.
Sugar conformations are in both cases in the general S domain with pseudorotation phase angles of 124 ~ (C-l'-exo) and 81 ~ (C-2'-endo), respectively. Some distortion is observed in the geometry of the cytidine sugar ring, arising probably from internal inconsistencies in the set of NOEs. Remarkably, the sugar conformations obtained for the 3',5'-d(CpA)-RNase A complex by X-ray crystallography are in an Ntype domain. To further check this discrepancy between the crystal and solution structures, we analyzed the 3j coupling constants between the sugar protons in the two complexes. N O E intensities between sugar protons are not very sensitive to the ring conformation, which can, however, be accurately determined from 3j values by means of the Karplus equation (Wijmenga et al., 1993). There are several methods to measure 3j coupling values. In our case, we have obtained the coupling constants by analyzing the DQF (double quantum filtered)-COSY spectra. To improve the accuracy of the estimation of 33, couplings, we have carried out computer simulations of the COSY cross-peaks. In these simulations, the experimental conditions, such as line widths or apodization functions, are reproduced by the program, and the values for the 3j involved are varied manually until a good matching with the experimental cross-peak is achieved. Because 3JHI,H2, is very small for rings with low pseudorotation phase angles, this coupling constant is very informative to discard N-type sugar conformations. In the case of the 3',5'-d(CpA) complex, the two ribose rings show values of 3JH~,2H2,~ around 7.5 Hz and 3JH~,H2,2 of 6.5 Hz. In both cases, the values are consistent only with sugar puckers in the general S domain. In the case of 2',5'-CpA complex, 3JHI,H2, coupling constants are small in both ribose rings, as confirmed for the lack of H - I ' - H - 2 ' cross-peak in the phase sensitive COSY spectrum. As mentioned before, the cytidine mononucleotides and the two dinucleotides have been studied by X-ray crystallography. Many of the structural features observed in the solution studies are common to the crystal structures, although some interesting differences there exist. Especially remarkable is the presence of two conformers of the side chain of His119 in the solution structures of all mononucleotide complexes. Only one of these positions is observed in the crystallographic studies of 2'CMP (conformation A) (Lisgarten et al., 1993) and 3'-CMP (conformation B) (Zegers et aL, 1994). In the case of the dinucleotide complexes, only one position is observed, in complete agreement with the X-ray results. Sugar puckers in solution in some cases differ from the ones in crystal state. In the 3'-mononucleotide complexes, the ribose pseudorotation
11
NMR Structures of RNase A and Its Complexes
373
phase angles are large (S domain), whereas in the crystal structure of the 3'-CMP RNase complex (the only 3' complex available) this pseudorotation phase angle is small (the sugar is in the N domain). In the case of 2'-mononucleotide complexes, the solution and crystallographic results agree and in the two states the riboses are in an N-type conformation. However, the deoxyribose sugar rings in the 3',5'-d(CpA)-RNase A complex adopt S-type conformations, in contrast to the crystallographic results (see Note added in proof). Comparison of the dihedral angles involving the phosphate moiety of the dinucleotide complexes is not possible due to the poor definition of the solution structures in this region.
IV. S T R U C T U R E S O F O T H E R A N I M A L R I B O N U C L E A S E S IN S O L U T I O N
A. Bovine Seminal Ribonuclease
Bovine seminal ribonuclease is the only dimeric protein of the pancreatic-type superfamily, with the subunits linked by two disulfide bridges (D'Alessio et al., 1975, 1992; Di Donato and D'Alessio, 1972). Its subunit, monomeric bovine seminal ribonuclease (mBS-RNase), is an unusual monomer with a high structural stability (D'Alessio et al., 1972) and higher catalytic activity than that of the parent dimeric enzyme (Piccoli et al., 1988). mBs-RNase is homologous to bovine pancreatic ribonuclease A, with a primary structure more than 80% identical, including the residues found at the active site of RNase A. The assignment of the ~H NMR spectrum of mBS-RNase in solution and its secondary structure determination have been reported (D'Ursi et al., 1995). The general pattern of sequential and medium-range connectivities is the same for mBS-RNAse and RNase A (Rico et al., 1991), indicating similar secondary structures for both proteins. The same conclusion is obtained by comparing H a conformation shifts for both proteins. The tertiary structure determination and refinement of monomeric bovine seminal ribonuclease are being carried out by the same research group using an iterative approach based on a full relaxation matrix interpretation of NMR spectra and several molecular dynamics (MD) and energy minimization procedures. The starting model for molecular dynamics and energy minimization procedures has been the X-ray struc-
374
Carlos Gonz~ilez et al.
ture of the A chain of dimeric BS-RNase. The calculation procedures employ a growing number of restraints, up to 1000; many of them have been also quantitatively evaluated (200). Three final 500-psec MD simulations were significant: without restraints, with fixed restraints, and with time-averaged restraints (Torda et al., 1990). The resulting trajectories have been analyzed with particular attention to (1) the residues responsible for dimerization and tertiary arrangement of the dimeric mBS-RNase, (2) the residues involved in the catalytic activity, and (3) the flexible loops, where a high number of substitutions with respect to the RNase A are present. The conformation of region 16-22 (hinge peptide), connecting the first c~-helical region (which in the native dimeric BS-RNase interacts with the main body of the other subunit) with the second helix, is characterized by a high degree of flexibility, confirmed by the lack of a welldefined pattern of NOEs between strands 13-16 and 43-48 as observed in RNase A. This indicates that the N-terminal tail of mBS-RNase interacts less strongly with the body of the protein than it does in the case of RNase A, and favors the swapping of the S peptide. The flexibility of the "hinge peptide" determines also a slightly higher mobility of the residues involved in the catalytic activity. In particular, the flexibility of the His-ll9 side chain is retained, while the orientation of the His-12 side chain is less rigidly determined. A possible origin of this flexibility is the lack of a partially extended segment around residue 13, which is present both in RNase A and in dimeric BS-RNase. This segment interacts in a "parallel/3-sheet-like" fashion with the 43-49 strand, stabilizing the folding of the "hinge peptide" and the position of His-12. In mBSRNase, the 12-16 segment exhibit a pseudohelical conformation, and no H bonds with the 43-48 strand are present. The region around Cys-31 and Cys-32 is responsible for the disulfide bridges between the two subunits in dimeric BS-RNase (Mazzarella et aL, 1993), and exhibits considerable sequence difference from RNase A. As expected, large conformational differences are detected by comparison with the two RNases: the regularity of the helical structure in mBSRNase is already lost around residues 28, giving rise to a semiextended pseudohelical region. Characteristic conformations are found for several loops, where many substitutions in the primary structure are concentrated compared to the RNase A. In particular the conformation of loop 65-72 shows the more significant peculiarity due to the deamidation of the Asn-67 that leads to a prevalence of an isopeptidic bond generated during the deamidation process.
11
NMR Structures of RNase A and Its Complexes
375
B. Angiogenin Angiogenins are 14-kDa proteins able to induce blood vessel growth in various preparations and are thought to be involved in the development of solid tumors. They belong to the pancreatic ribonuclease superfamily as shown by sequence similarities with RNase A. These proteins possess a ribonucleolytic activity but they are extremely weak ribonucleases toward conventional RNase A substrates, and they differ as well in specificity. However, this ribonucleolytic activity appears to be crucial for angiogenesis. In addition, they are able to induce secondmessenger pathways, enter endothelial cells, and translocate to the nucleus. The solution structure of bovine angiogenin by ~H NMR spectroscopy has been determined by the group of Lallemand (Reisdorf et al., 1994; Lequin et al., 1996). Backbone conformations of the 10 structures of bovine angiogenin are shown superimposed in Fig. 11. Residues 1-3 and 123-125, which are not constrained, have been omitted. The average RMS deviation for the backbone atoms of residues 5-121 is 0.71 ~, (1.39 ,~ for all backbone atoms). The NMR samples were prepared in
Fig. 11. Stereoscopic view of the three-dimensional solution structure of bovine angiogenin. Superposition of 10 final structures.
376
Carlos Gonz~ilez et al.
50 mM sodium phosphate, pH 4.5 or 5.0, in either H20/D20 (9:1, by volume) or D20. The final concentration was 4 mM. Structures were calculated using 1117 distance constraints, including 490 long-range constraints and 78 constraints to specify hydrogen bonds. Structure calculations were performed with a procedure combining minimization in the dihedral space with the DIANA program, followed by simulated annealing and a final minimization at 0 K with the XPLOR program. The general fold is reminiscent of that of RNase A, including the three helices and the antiparallel twisted/3 sheet with three strands on one side and four strands on the other side. The major discrepancy concerns loop 67-69, which is longer in RNase A and stabilized by a fourth disulfide bridge. In angiogenins, this loop is involved in binding to cellular receptors. The regular secondary structure elements are well defined, the average RMS deviation value is only 0.43 ~, for backbone atoms of helices and the/3 sheet. But helix II is rather loose and loops display an increased disorder, as illustrated in Fig. 11. In particular, the loop connecting strands 4 and 5 (residues 86-94) as well as the region 36-41 shows the largest dispersion. During the refinement of the solution structure of bovine angiogenin, the crystal structures of human and bovine angiogenins were reported (Acharya et al., 1994, 1995). Because the atomic coordinates of bovine angiogenin have not yet been released to the Protein Data Bank, we cannot give a detailed comparison of both structures. However, the solution structures of bovine angiogenin seem to be similar to that of crystal state, in terms of global shape, backbone overall fold, and definition of secondary structure elements. The spatial positions of the three RNase A residues His-12, His-ll9, and Lys-41, known to be important in ribonucleolytic catalysis, are conserved in angiogenins. In the solution structure of bovine angiogenin, the active site residues His-14, His-115, and Lys-41 are rather well defined and correspond to the observed positions in RNase A (see Fig. 12). A dynamic equilibrium between two positions of the His-119 side chain of Rnase A has been underlined (Rico et al., 1991). Observation of the alternative position in angiogenin would require identification of NOEs between His-ll5 and methyl protons of Val-114 (Val-ll8 in RNase A). These NOEs could not be identified owing to the strong overlap in the corresponding region of the NOESY spectra. Therefore an alternative conformation of His-ll5 in bovine angiogenin cannot be excluded. In addition, crystallographic and mutagenesis data (Acharya et al., 1994, 1995; Russo et al., 1994) have led to the proposal that the position of the side chain of Gln-ll7 or Glu-ll8 in human or bovine angiogenin,
11
377
NMR Structures of RNase A and Its Complexes
,.%');'
His 14
Glu 118
Phe 116
His 115 His 1|
Fig. 12. Stereo view of the active site of bovine angiogenin: site P1 (His-14, Lys-41, His-ll5), site BI (Thr-45), and Glu-ll8, which obstructs the B1 site.
found to obstruct the p u t a t i v e p y r i m i d i n e binding site, could account for the low r i b o n u c l e a s e activity, and that a c o n f o r m a t i o n a l c h a n g e would be necessary prior to or during catalysis. N O E c o r r e l a t i o n s w e r e o b s e r v e d b e t w e e n the side chain H~ p r o t o n s of Glu-118 and the imidazole ring H ~ p r o t o n of His-14, which orients the side chain of G l u - l l 8 in the active site. In all structures it obstructs the putative p y r i m i d i n e binding site; h o w e v e r , it a p p e a r s to be r a t h e r flexible. T h e a u t h o r s did not find any additional N O E s that would define a n o t h e r position of this side chain. T h e y suggest that e i t h e r a different c o n f o r m a t i o n would be a very m i n o r c o n f o r m e r or that a c o n f o r m a t i o n a l c h a n g e would occur during, r a t h e r than prior to, catalysis.
ACKNOWLEDGMENTS Thanks are given to Drs. J. L. Neira, M. Bruix, and C. Toiron for providing original data on the assignment and structure determination of RNase A and its complexes, as well as to the research groups of Profs. Lallemand (Paris) and Piccone (Naples) for sharing structural data on angiogenin and monomeric bovine seminal RNase prior to publication.
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The excellent technical assistance of A. G6mez, C. L6pez, L. de la Vega, and D. A. Santos, as well as the revision of the manuscript by Dr. G. M. Langdon, is gratefully acknowledged.
NOTE
Both here and in our recently published work on the solution structure of the complexes of RNase A with 2',5'-CpA and 3',5'-d(CpA) (Toiron et al., 1996), the different sugar conformations in the complexes of RNase A with 3'-CMP and 3',5'-d(CpA) have been noted as the main discrepancy between the solution and the crystal structures (Zegers et al., 1994). After publication of our paper on the solution structure, we found an error in the calculation of the sugar pseudorotation phase angles reported in the paper by Zegers et al., (1994). According to the PDB coordinates (entry 1RPG), the deoxyriboses do not adopt a N-type conformation but a S-type conformation in both nucleotides, with phase angles very similar to the ones observed in solution. This makes the solution and crystal structures more closely similar.
REFERENCES
Acharya, K. R., Shapiro, R., Allen, S. C., Riordan, J. F., and Vallee, B. L. (1994). Crystal structure of human angiogenin reveals the structural basis for its functional divergence from ribonuclease. Proc. Natl. Acad. Sci. U.S.A. 91, 2915-2919. Acharya, K. R., Shapiro, R., Riordan, J. F., and Vallee, B. L. (1995). Crystal structure of bovine angiogenin at 1.5 ,~, resolution. Proc. Natl. Acad. Sci. U.S.A. 92, 2949-2953. Berman, E., Waiters, D. E., and Allerhand, A. (1981). Structure and dynamic behaviour of the oligosaccharide side chain of bovine pancreatic ribonuclease A. J. Biol. Chem. 256, 3853-3857. Blackburn, P., and Moore, S. (1982). Pancreatic ribonuclease. In "The Enzymes," (P. D. Boyer, ed.), Vol. 15, pp. 317-433. Academic Press, New York. Boelens, R, Koning, T. M. G., and Kaptein, R. (1988). Determination of biomolecular structures from proton-proton NOEs using a relaxation matrix approach. J. Mol. Struct. 173, 299-311. Boelens, R., Koning, T. M. G., van der Marel, G. A., van Boom, J. H., and Kaptein, R. (1989). lterative procedure for structure determination from proton-proton NOEs using a full relaxation matrix approach. Application to a DNA octamer. J. Magn. Res. 82, 290-308. Borgias, B. A., and James, T. L. (1988). COMATOSE. A method for constrained refinement of macromolecular structure based on two-dimensional nuclear Overhauser effect spectra. J. Magn. Res. 79, 493-512. Borkakoti, N., Moss, D. S., and Palmer, R. A. (1982). Ribonuclease A: Least-squares refinement of the structures at 1.45,~ resolution. Acta Crystallogr. B 38, 2210-2217. Case, D. A., Dyson, H. J., and Wright, P. E. (1994). Use of chemical shifts and coupling constants in nuclear magnetic resonance structural studies on peptides and proteins.
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pp. 393-416. Academic Press, San Diego. D'Alessio, G., Floridi, A., De Prisco, R., Piguero, A., and Leone, E. (1972). Bull semen ribonucleases. 1. Purification and physico-chemical properties of the major component. Eur. J. Biochem. 26, 153-161. D'Alessio, G., Maloni, M. C., and Parente, A. (1975). Dissociation of bovine seminal ribonuclease into catalytically active monomers by selective reduction and alkylation of the intersubunit disulphide bridges. Biochemistry 14, 1116-1122. D'Alessio, G., Di Donato, A., Parente, A., and Piccoli, R. (1992). Seminal RNase: A unique member of the ribonuclease superfamily. Trends Biochem. Sci. 16, 104-106. D'Ursi, A., Oschkinat, H., Cieslar, C., Piccone, D., D'Alessio, G., Amodeo, P., and Temussi, P. (1995). Assignment and secondary structure determination of monomeric bovine seminal ribonuclease employing computer-assisted evaluation of homonuclear three-dimensional IH-NMR spectra. Eur. J. Biochem. 229, 494-502. Di Donato, A., and D'Alessio, G. (1973). Interchain disulphide bridges in ribonuclease BS-1. Biochem. Biophys. Res. Commun. 55, 919-928. Dobson, C. M., and Lian, Y. L. (1987). A alp MAS NMR study of cytidine 2'-phosphate bound to ribonuclease A in the crystalline state. F E B S Lett. 225, 183-187. Egan, W., Shindo, H., and Cohen, J. S. (1978). On the tyrosine residues of ribonuclease A. J. Biol. Chem. 253, 16-17. Eftink, M. R., and Biltonen, R. L. (1987). Pancreatic ribonuclease A: The most studied endo-ribonuclease. In "Hydrolitic Enzymes" (A. Neuberger and K. Brocklehorst, eds.), pp. 333-376. Elsevier, New York. Gonz~ilez, C., Rullmann, J. H. A., Bonvin, A. M. J. J., Boelens, R., and Kaptein, R. (1991). Toward an NMR R factor. J. Magn. Res. 91, 659-664. Gorenstein, D. G., Wyrwicz, A. M., and Bode, J. (1976). Interaction of uridine and cytidine monophosphates with ribonuclease A. IV. Phosphorus-31 NMR studies. J. Am. Chem. Soc. 98, 2308-2314. Hahn, U., and Rtiterjans, H. (1985). Two-dimensional IHNMR investigation of ribonuclease A and ribonuclease A-pyrimidine-nucleotide complexes. Eur. J. Biochem. 152, 481-491. Hahn, U., Desai-Hahn, R., and Rtiterjans, H. (1985). ~H and ~SN investigation of the interaction of pyrimidine nucleotides with ribonuclease A. Eur. J. Biochem. 146, 705-712. Howlin, B., Harris, G. W., Moss, D. S., and Palmer, R. A. (1987). X-Ray refinement study on the binding of cytidylic acid (2'oCMP) to ribonuclease A.J. Mol. Biol. 196,159-164. Jardetzky, O., and Roberts, G. C. K. (1981). "NMR in Molecular Biology." Academic Press, New York. Joao, H. C., and Dwek, R. A. (1993). Effects of glycosylation on protein structure and dynamics in ribonuclease B and some of its glycoforms. Eur. J. Biochem. 218, 239-244. Joao, H. C., Scragg, I. G., and Dwek, R. A. (1992). Effects of glycosylation on protein conformation and amide proton exchange rates in ribonuclease B. F E B S Lett. 307, 343-346. Kartha, G., Bello, J., and Harker, D. (1967). Tertiary structure of ribonuclease. Nature (London) 213, 862-865. Koning, T. M. G., Boelens, R., and Kaptein, R. (1990). Calculation of the nuclear Overhauser effect and the determination of proton-proton distances in the presence of internal motions. J. Magn. Res. 90, l l 1-123. Kuriyan, J., Osapay, K., Burley, S. K., Brtinger, A. T., Hendrickson, W. A., and Karplus, M. (1991). Exploration of disorder in protein structures by X-ray restrained molecular dynamics. Proteins 10, 340-358.
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Lenstra, J. A., Bolscher, G. J. M., Stob, S., Beitema, J. J., and Kaptein, R. (1979). The aromatic of bovine pancreatic ribonuclease studied by ~H NMR. Eur. J. Biochem. 98, 385-397. Lequin, O., Albaret, C., Bontems, F., Spik, G., and Lallemand, J.-Y. (1996). Solution structure of bovine angiogenin by lH nuclear magnetic resonance spectroscopy. Biochemistry 35, 8870-8880. Lisgarten, J. N., Gupta, V., Maes, D., Wyns, L., Zegers, I., Palmer, R. A., Dealwis, C. G., Aguilar, C. F., and Hummings, A. M. (1993). Structure of the crystalline complex of cytidylic acid (2'-CMP) with ribonuclease at 1.6A resolution. Conservation of solvent sites in RNase A high-resolution structures. Acta Crystallogr. D 49, 541-547. Markley, J. L. (1975). Correlation proton magnetic resonance studies at 250 MHz of bovine pancreatic ribonuclease. I. Reinvestigation of the histidine peak assignments. Biochemistry 14, 3546-3554. Mazzarella, L., Capasso, S., Demasi, D., Di Lorenzo, G., Mattia, C. A., and Zagari, A. (1993). Bovine seminal ribonuclease: Structure at 1.9A resolution. Acta Crystallogr. D 49, 389-402. Meadows, D. H., Roberts, G. C. K., and Jardetzky, O. (1969). NMR studies of the structures and binding sites of enzymes. VIII. Inhibitor binding to ribonuclease. J. Mol. Biol. 45, 491-511. Neira, J. L., and Rico, M. (1996). H/D exchange and NMR studies of stability and folding of RNase A, submitted. Piccoli, R, DiDonato, A., and D'Alessio, G. (1988). Co-operativity in seminal ribonuclease function. Biochem. J. 253, 329-336. Puctt, D. (1973). Conformational studies on a glycosylatcd bovine pancreatic ribonuclcasc. J. Biol. Chem. 248, 3566-3572. Reisdorf, C., Abergel, D., Bontems, F., Lallemand, J. Y., Decottignies, J., and Spik, G. (1994). Proton rcsonancc assignmcnts and sccondary structure of bovine angiogcnin. Eur. J. Biochem. 224, 811-822. Richards, F. M., and Wyckoff, H. W. (1971). Bovine pancreatic ribonuclease. In "The Enzymes" (P. D. Boycr, cd.), Vol. 4, pp. 647-806. Academic Press, New York. Richards, F. M. (1977). Areas, volumes, packing and protein structure. Annu. Rev. Biophys. Bioeng. 6, 151-176. Richardson, J. S. (1981). The anatomy and taxonomy of protein structure. Adv. Protein Chem. 34, 167-339. Rico, M., Bruix, M., Santoro, J., Gonz~ilez, C., Neira, J. L., Nieto, J. L., and Herranz, J. (1989). Sequential ~H-NMR assignment and solution structure of bovine pancreatic ribonuclease A. Eur. J. Biochem. 183, 623-638. Rico, M., Santoro, J., Gonz~ilez, C., Bruix, M., Neira, J. L., Nieto, J. L., and Herranz, J. (1991). 3D structure of bovine pancreatic ribonucleasc A in aqueous solution: An approach to the tertiary structure determination for a small basis of ~H NMR NOE correlations. J. Biomol. N M R 1, 283-298. Rico, M., Santoro, J., Gonzfilez, C., Bruix, M., Neira, J. L., and Nieto, J. L. (1993). Refined solution structure of bovine pancreatic ribonuclease A by ~H NMR methods. Sidechain dynamics. Appl. Magn. Reson. 4, 385-415. Robertson, A. D., Purisima, E. D., Eastman, M. A., and Scheraga, H. A. (1989). Proton NMR assignments and regular backbone structure of bovine pancreatic ribonuclease A in aqueous solution. Biochemistry 28, 5930-5938. Rose, G. D., Gierasch, L. M., and Smith, J. A. (1985). Turns in peptides and proteins. Adv. Protein Chem. 37, 1-109.
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Rudd, P. M., Joao, H. C., Coghill, E., Fiten, P., Saunders, M. R., Opdenakker, G., and Dwek, R. A. (1994). Glycoforms modify the dynamic stability and functional activity of an enzyme. Biochemistry 33, 17-22. Russo, N., Shapiro, R., Acharya, K. R., Riordan, J. F., and Vallee, B. L. (1994). Role of glutamine-ll7 in the ribonucleolytic activity of human angiogenin. Proc. Natl. Acad. Sci. U.S.A. 91, 2920-2924. Santoro, J., Juretschke, H. P., and Rtiterjans, H. (1979). ~3C NMR investigations on ribonuclease A. Biochim. Biophys. Acta 578, 346-356. Santoro, J., Gonz~lez, C., Bruix, M., Neira, J. L., Nieto, J. L., Herranz, J., and Rico, M. (1993). High-resolution three-dimensional structure of ribonuclease A in solution by NMR spectroscopy. J. Mol. Biol. 229, 722-734. Saunders, M., Wishnia, A., and Kirkwood, J. G. (1957). The nuclear magnetic resonance spectrum of ribonuclease. J. Am. Chem. Soc. 79, 3285-3290. Svensson, L. A., Sj01in, L., Dill, J., and Gilliland, G. L. (1986). Multiple conformations of amino residues in ribonuclease A. Proteins 1, 370-375. Svensson, L. A., Sj01in, L., Dill, J., and Gilliland, G. L. (1991). The conformation flexibility of surface residues of bovine pancreatic ribonuclease A at 1.1A resolution. In "Structure, Mechanism and Function of Ribonucleases, Proceedings of the Second International Meeting" (C. M. Cuchillo, R. de Llorens, M. V. Nogu6s, and X. Par6s, eds.), pp. 31-38. Universitat Aut6noma de Barcelona, Barcelona. Toiron, C., Gonz~ilez, C., Bruix, M., and Rico, M. (1996). Three-dimensional structure of the complexes of ribonuclease A with 2',5'-CpA and 3',5'-d(CpA) in aqueous solution, as obtained by NMR and restrained molecular dynamics. Protein Sci. 5, 1633-1647. Torda, A. E., Scheck, R. M., and van Gunstcrcn, W. F. (1990). Time averaged NOE distance restraints applied to Tendamistat. J. Mol. Biol. 214, 223-235. Van Gunstcrcn, W. F., and Berendsen, H. J. C. (1987). "Groningcn Molecular Simulation (GROMOS) Library Manual." Biomos, Groningcn, The Netherlands. Waiters, E., and Allcrhand, A. (1980). Tautomcric states of the histidinc residues of bovine pancreatic ribonuclease A. J. Biol. Chem. 255, 6200-6204. Williams, R. L., Green, S. M., and McPhcrson, A. (1987). The crystal structure of ribonucicase B at 2.5A resolution. J. Biol. Chem. 262, 16020-16031. Wijmcnga, S., Moorcn, M., Hilbcrs, C. W. (1993). NMR of nucleic acids; from spectrum to structure. In "NMR in Macromolecules," (G. C. Roberts, ed.). IRL Press, Oxford. Wlodawcr, A. (1984). Structure of bovine pancreatic ribonuclease by X-ray and neutron diffraction. In "Biological Macromolecules and Assemblies. Volume II. Nucleic Acids and Interactive Proteins," (F. Jurnak and A. McPherson, eds.), pp. 395-439. Wiley, New York. Wlodawer, A., Svensson, L. A., Sj61in, L., and Gilliland, G. L. (1988). Structure of phosphate-free ribonuclease A refined at 1.26A. Biochemistry 27, 2705-2717. Wodak, S. Y., Lie, M. Y., and Wyckoff, H. W. (1977). The structure of cytidyl (2'-5') adenosine when bound to pancreatic RNase S. J. Mol. Biol. 116, 855-875. Wtithrich, K. (1986). "NMR of Proteins and Nucleic Acids." Wiley, New York. Wyckoff, H. W., Tsernoglou, D., Hanson, A. W., Knox, J. R., Lee, B., and Richards, F. M. (1970). The three dimensional structure of ribonuclease S. Interpretation of an electron density map at a nominal resolution of 2~,. J. Biol. Chem. 245, 305-328. Zegers, I., Maes, D., Dao-Thi, M., H., Poortmans, F., Palmer, R. A., and Wyns, L. (1994). The structure of RNase A complexed with 3'-CMP and d(Cpa): Active site conformation and conserved water molecules. Protein Sci. 3, 2322-2339.
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12 Seminal Ribonuclease" The Importance of Diversity G I U S E P P E D'ALESSIO,* A L B E R T O DI D O N A T O , * LELIO
MAZZARELLA,t
AND
RENATA
PICCOLI*
* Dipartimento di Chimica Organica e Biologica tDipartimento di Chimica Universith Degli Studi di Napoli Federico II 80134 Naples, Italy
I. Introduction II. Isolation and Production of Seminal RNase A. Preparation from Natural Sources B. Production of Recombinant BS-RNase III. Structures A. Covalent Structure B. Three-Dimensional Structure C. Folding Pathway D. Chemical Modifications and Engineering IV. Functions A. Catalytic Function B. Special Biological Actions References
I. I N T R O D U C T I O N
A person or an object is described as "diverse" as it is unlike, different f r o m the other m e m b e r s o f its group; and can be "diverse" because it is heterogeneous and exists in various forms. Seminal ribonuclease is a "diverse" ribonuclease on both counts.
383 RIBONUCLEASES: STRUCTURES AND FUNCTIONS
Copyright 9 1997 by Academic Press, Inc. All rights of reproduction in any form reserved.
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For many years the term "ribonuclease" has been synonymous with bovine pancreatic ribonuclease A, and for a long time the RNases discovered in mammals and molds were all monotonously monomeric proteins, with a small molecular size, typically Michaelian enzymes, with no biological action other than their catalytic degradation of RNA. Then a new RNase was discovered in bull seminal fluid (D'Alessio and Leone, 1963) and seminal vesicles (Hosokawa and Irie, 1971); it was much larger in size than RNase A (Forlani et al., 1967), and dimeric (D'Alessio et al., 1972b). To date, bovine seminal RNase (BS-RNase) is still a diverse RNase: "different" from the historic prototype RNase A m a n d from all other RNases of the vertebrate superfamilymfor its dimeric structure, for its non-Michaelian kinetics (Piccoli et al., 1982), and for its special, i.e., noncatalytic, biological actions (D'Alessio et al., 1991). But BS-RNase is also a diverse RNase because it exists in a multiplicity of structural forms, and is endowed with a multiplicity of biological actions. Two quaternary conformations (Piccoli et al., 1992) and three isoenzymatic subunit compositions (Di Donato and D'Alessio, 1981) have been described for BS-RNase. Furthermore, BS-RNase performs a surprising array of biological actions: aspermatogenic, antitumor, immunosuppressive, and antiviral (see Section IV,B). BS-RNase may not be the only seminal RNase: an RNase has been purified from human semen (De Prisco et al., 1984) and low levels of RNase activity have been detected in the semen of several mammals, including mouse, rabbit, and sheep (G. D'Alessio, unpublished). Certainly, it would be surprising not to find RNase activity in a biological fluid, especially because all vertebrate RNases studied so far are proteins destined to be exported as extracellular enzymes. However, what makes BS-RNase again diverse, in this case with respect to the other seminal RNase activities from other mammals, is its great abundance. Its concentration in bull semen (about 1.5 mg/ml of plasma) is three orders of magnitude higher than that of the RNase purified from human semen, and attests to a great effort supported by the protein synthesis machinery of bull seminal vesicle cells. Furthermore, BS-RNase appears to be the result of some consequential evolutionary events that provided the bovine genome with a specific gene for the protein (Confalone et al., 1995), to be expressed specifically in the seminal vesicle glands (Sasso, 1993). The only two species in which a functional seminal RNase gene has been found belong to the same family and are strictly related: B o s taurus, from which BS-RNase is isolated, and B u b a lu s bubalus, the water buffalo (Confalone et al., 1995). All this suggests that BS-RNase serves a definite purpose in the bovine reproductive physiology and performs a task that
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in other, even closely related, species is not needed, or is carried out by non-RNase molecule(s). Another peculiarity about BS-RNase is that it has been repeatedly and independently discovered and rediscovered in different laboratories. After the first preliminary note announcing the finding of copious RNase activity in bull semen (D'Alessio and Leone, 1963), and a study of its molecular weight (Forlani et al., 1967), seminal RNase was isolated in Kyoto from bull seminal vesicles (Hosokawa and Irie, 1971), and then a detailed report was produced by the group in Naples on its purification from seminal plasma (D'Alessio et aL, 1972a). At the same time Dost~l and Matou~ek (1972) isolated from bull semen an "aspermatogenic substance" that was soon identified as BS-RNase (Matougek et al., 1973a; Leone et al., 1973). Next, an apparently different RNase was isolated from bull semen (Reddy et al., 1979), but it was later determined that this was the same enzyme previously isolated in other laboratories (D'Alessio et al., 1981). It should also be mentioned that originally both of the groups in Naples and in Kyoto identified two RNase activity peaks in the final chromatographic separation of the purification procedure. These were termed RNase Vs~ and Vs2 from seminal vesicles (Hosokawa and Irie, 1971), and RNase BS-1 and BS-2 from seminal plasma (D'Alessio et al., 1972a), with VSl and BS-1, respectively, as the major RNase component. Later, it was reported that RNase BS-2, the minor RNase component of bull semen, was not a distinct RNase, nor a homogenous protein, but rather a mixture of RNases, including pancreatic RNase A and at least one hybrid RNase made up of a subunit of seminal RNase linked through disulfide(s) to an unidentified peptide moiety (D'Alessio et al., 1981). Thus BS-1 RNase was identified as the only RNase proper of bull semen, and became simply BS-RNase. BS-RNase has been briefly reviewed by Blackburn and Moore (1982), and by D'Alessio et al. (1991).
II. ISOLATION A N D P R O D U C T I O N OF SEMINAL RNase
A. Preparation from Natural Sources
Different purification protocols were used by the various groups who independently isolated seminal RNase, as reported in Section I. Since
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then, simpler and more rapid procedures have been proposed, based on ion-exchange chromatography (Tamburrini et al., 1986), or affinity chromatography with agarose-linked nucleotides (Krietsch et al., 1983), or cellulose-linked DNA (Murti and Pandit, 1983). Ion-exchange chromatography remains the most effective, rapid, and inexpensive procedure for extracting BS-RNase from seminal plasma. More than 20 mg of protein can be prepared by cation-exchange chromatography in 2 days from 20 ml of plasma, and 1 mg in 15 min from 1 ml, with carboxymethyl-cellulose, or Mono S columns, respectively (Tamburrini et al., 1986). The same chromatographic protocols can be applied to a 50-100% saturated ammonium sulfate fraction from a bull seminal vesicle homogenate. It has been found (N. Russo, unpublished) that when the final 100% saturated ammonium sulfate supernatant, as recovered after centrifugation of the precipitate, is brought to pH 9.1 by careful addition of sodium hydroxide, a residual fraction of homogeneous BS-RNase can be precipitated, with a significant improvement in the total yield of BS-RNase from seminal vesicle tissue.
B. Production of Recombinant BS-RNase
A cDNA encoding the subunit sequence of BS-RNase was cloned by Preuss et al. (1990) by screening a cDNA library from bovine seminal vesicle tissue, and a synthetic cDNA coding for the protein was constructed by Kim and Raines (1993). A semisynthetic cDNA (de Nigris et al., 1993) coding for the same sequence was also constructed by ligating a synthetic DNA segment coding for residues 1-48 of the BS-RNase amino acid sequence to a cDNA segment obtained from BS-RNase mRNA (Palmieri et al., 1985), and coding for residues 49 through 124. Both the fully synthetic and the semisynthetic cDNAs encoding BSRNase subunit sequence were cloned and expressed in E s c h e r i c h i a coli (de Nigris et al., 1993; Kim and Raines, 1993) with the T7 expression system (Studier et al., 1990). The semisynthetic cDNA was expressed also in Chinese hamster cells (Russo et al., 1993). When expressed in prokaryotic cells, recombinant BS-RNase is sequestered in inclusion bodies, from which it can be solubilized with acid and/or denaturants in the presence of thiol reagents. When these
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are removed by dialysis or gel filtration, the fully reduced and denatured protein can be refolded through air oxidation (Kim and Raines, 1993) or in the presence of a glutathione redox buffer (de Nigris et al., 1993). In the former procedure only a fraction of the oxidized protein was obtained as a dimer, whereas the remaining was monomeric, or with an unexpected behavior on a gel-filtration column, which could suggest the molecular size of a trimer (Kim and Raines, 1993). On further analyses, it has been proposed that this component of the reoxidation mixture is a dimer in which one of the two subunits is not correctly folded, and one of the two intersubunit disulfides is not reoxidized (Kim and Raines, 1994). Furthermore, on air reoxidation of unfolded recombinant BS-RNase, a series of active components, suspected to be mixed disulfides between a BS-RNase subunit and peptides (Kim and Raines, 1993), and a poorly active monomeric component (Kim and Raines, 1994) were also produced. When refolding and reoxidation of fully reduced and denatured recombinant BS-RNase is carried out in the presence of a glutathione redox buffer (Smith et al., 1978) made up of 3 mM glutathione and 0.6 mM oxidized glutathione, about 15% of the protein folds and associates into dimeric BS-RNase; the remainder refolds into a monomeric, active form, in which Cys-31 and Cys-32, responsible for the two intersubunit disulfide bridges of dimeric BS-RNase, are linked into mixed disulfides with glutathione moieties (de Nigris et al., 1993). On removal of glutathione by selective reduction of the mixed disulfides (D'Alessio et al., 1975), followed by gel filtration and by air reoxidation of the intersubunit disulfides, all the monomeric protein is associated into dimeric BS-RNase. The final yield of dimeric BS-RNase is about 8 mg/liter of original cell culture. This procedure does not give any by-products, likely because it mimics a physiological pathway (see Section Ill,C). Fully reduced and unfolded BS-RNase may refold and then associate into only one of the dimeric protein forms, the form termed M - - M (see Sections III,B,2 and III,B,3,a), whereas two-thirds of the protein isolated from natural sources is in the M • M form, characterized by the exchange of the N-terminal domains between subunits, and only one-third is in the M - - M form, with no exchange. Thus the recombinant newly refolded and dimerized BS-RNase has to be incubated at 37~ to speed up the lengthy transformation of M = M to M • until the equilibrium ratio typical of natural BS-RNase is reached (see Section III,B,2). Likely, during the long incubation to equilibrate M - - M with M • M, the selective
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deamidation of Asn-67 (see Section III,A,3) occurs spontaneously, just like in the seminal vesicle tissue. After this treatment, the recombinant protein expressed in prokaryotes is catalytically as active as natural BS-RNase and has the same dual quaternary conformation and isoenzymatic pattern. It differs from the natural product, however, both in structure, for an extra Met at its N terminus, and in function, for its low activity in the antitumor assay, apparently dependent on the presence of the N-terminal methionine (Adinolfi et al., 1995). Met -1 can be eliminated either with an appropriate aminopeptidase or by expression of the cDNA encoding the protein ligated at its 5' end to a leader peptide that contains a cleavage site for Factor Xa (Adinolfi et al., 1995). Expression of recombinant BS-RNase with a eukaryotic system yields instead a protein with all the structural and functional features of the natural product, including full antitumor activity and no extra Met at the N terminus (Russo et al., 1993). This was obtained by expressing in Chinese hamster cells under the metallothionein IIA promoter (Friedman et al., 1989) the semisynthetic cDNA encoding the subunit sequence of BS-RNase (see above) previously ligated at its 5' end to a synthetic DNA segment coding for the signal sequence of BS-RNase (Preuss et al., 1990). The recombinant protein, secreted by the hamster cells, is easily concentrated and purified from the cell culture medium. However, as is often the case with eukaryotic expression systems, the yield (about 0.5 mg/liter) was much lower than those reported for the prokaryotic expression systems.
III. S T R U C T U R E S
As isolated from either bovine seminal plasma or from seminal vesicle tissue, BS-RNase displays a complex microheterogeneity, i.e., a variety of structures, which are in fact isomers, chemical or conformational, of the protein (see below). However, on first inspection, BS-RNase appears as a very basic protein with a pl of 10.3 (D'Alessio et al., 1972a) with a formula weight of 27,218 Da (Suzuki et aL, 1987), made up of two subunits linked by two disulfide bridges as well as by noncovalent forces (D'Alessio et al., 1991).
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A. C o v a l e n t Structure
1. Significant Sequence Identities and Differences of B S - R N a s e within the Phylogenetic R N a s e Tree The amino acid sequence of the BS-RNase subunit chain (Suzuki et al., 1987) is more than 80% identical to that of RNase A (see Fig. 1). This makes BS-RNase a member of the vertebrate RNase superfamily (Beintema et al., 1988), which has RNase A as its historical prototype. When its amino acid sequence is compared to those of the other members of the superfamily (see Chapter 8), BS-RNase clearly appears to belong to the subfamily of pancreatic-type mammalian RNases, with which it shares at identical sequence positions the eight Cys residues that pair to form the four intrachain disulfides, and the residues that make up the catalytic site. Clearly, the gene duplication that originated the evolutionary history of BS-RNase occurred much later than the duplications lead-
Fig. 1. Amino acid sequence of the BS-RNase subunit. The intrachain disulfide links are shown, as well as the Cys residues at positions 31 and 32, which form the intersubunit disulfides.
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et
al.
ing to the divergence of frog RNases, of eosinophil RNases, and of angiogenins and reptile RNases. When the reconstruction of the immediate ancestor of BS-RNase and of bovine pancreas RNase is attempted by the inferential method (Libertini and Di Donato, 1994), or by the reconstruction strategy of Jermann et al. (1995), it appears that once duplicated, the ancestor RNase gene accumulated in one of its duplicates a series of mutations that led to BS-RNase. These mutations brought about the acquisition of 13 new residues, 5 of which are basic, including 4 residues that have been found to play a decisive role in the making of a dimeric ribonuclease featuring the exchange of the N-terminal c~ helices between subunits: Pro-19, Leu28, Cys-31, and Cys-32 (Di Donato et al., 1994, 1995; Mazzarella et al., 1995). However, with the exception of Val-102, already present in the putative ancestor sequence, and of G l y - l l l , Cys-31, and Cys-32, one or more of these residues are present in at least one other protein member of the vertebrate RNase superfamily. Although the uniqueness and significance of Gly-111 and Val-102 remain to be explained, the couple of adjacent Cys residues at positions 31 and 32 appear instead clearly related to the uniqueness of BS-RNase dimeric structure in the whole RNase superfamily. These Cys residues, which pair with the corresponding Cys residues of the other subunit (Cys-31 with Cys-32 and vice versa) to form the intersubunit disulfides (Di Donato and D'Alessio, 1973), have been shown to be both necessary and sufficient to determine the dimerization of an RNase. RNase A, engineered with Cys residues at positions 31 and 32 of its chain, spontaneously dimerizes (Di Donato et al., 1995). However, Leu-28 (which in the three-dimensional structure generates an intersubunit hydrophobic contact with the homologous residue of the partner subunit) and Pro-19 also have roles in maintaining a dimeric RNase, and significantly increase the proclivity of RNase monomers to exchange their N-terminal a helices (Di Donato et al., 1995). These and other findings can help us envision a possible evolutionary route from a monomeric to a dimeric RNase. Let us consider the following: (1) mutational events positioning one (Kim et al., 1995a) or two adjacent Cys residues (Di Donato et al., 1994) on a turn of the second RNase a helix can by themselves transform a monomeric RNase into a dimeric one; (2) other substitutions in nearby and distant regions of the molecule are, however, necessary to provide the dimeric construct with a significant and stable N-terminal exchange feature (see Section III,B,3); (3) seminal-like RNase genes with a single Cys have been found in the genomes of species evolutionarily related to the bovine group, such as
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the giraffe (Breukelman et al., 1993), the sheep, and the goat (Confalone et al., 1995). However, these have been positively identified as pseudogenes, i.e., vestigial traces of aborted evolutionary attempts, or relics of abandoned, unused genetic devices. In any case it would be arduous to conceive that these pseudogenes emerged later than the gene for BS-RNase. Based on these findings, the most plausible evolutionary route from a monomeric to a dimeric RNase seems to be the most obvious: one that goes through successive mutations, occurring on a duplicate RNase ancestor gene, and confers some advantage to the new RNase, hence to its host (D'Alessio, 1995)~just variations on an RNase theme, which happened to be much appreciated by some Bovidae. It does not seem easy to accommodate the above findings with the alternative hypothesis advanced by Bennett et al. (1994), suggesting that the potential ability to exchange domains between protomers is a basis for an evolutionary jump from a monomeric to a dimeric RNase. It should be added that the ability to "swap domains" is present in RNase A (Crestfield et al., 1962), and to a lesser extent in BS-RNase monomers (Parente et al., 1977), but both these proteins can associate into unstable dimers only, and only under conditions rather distant from those of a physiological environment, the proper conditions in which a potential candidate to evolutionary success must be tested. Of course the evolutionary success of a biomolecule is inexorably dependent on some advantage that the biomolecule provides to the host organism. It has been proposed (D'Alessio, 1995) that in the case of BS-RNase this advantage may be identified in some special bioactions (see Section IV,B) that the seminal RNase, and no other of its several other RNases, grants B o s taurus. As discussed in the following sections, two quaternary forms coexist in the protein, only one of them capable of the catalytic, RNA-degrading, bioaction, the other endowed also with special bioactions. Thus it may be that we are observing the slow birth of a fitter biomolecule, and that eventually only one of the two BS-RNase forms will survive, likely that richer in bioactions, because several genes coding for merely RNA-degrading enzymes are already available in the bovine genome. In conclusion, this may be a case of "evolution in progress" (D'Alessio, 1995). Confalone et al. (1995) have advanced a different hypothesis for the evolutionary success in some Bovidae of BS-RNase. Considering the embryotoxic effects of BS-RNase (see Section IV,B,3), and considering that fertilization can occur in cows in the absence of most seminal plasma
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components, including BS-RNase, the "mating of a pregnant cow with a bull carrying BS-RNase [may have led] to abortion of the previous embryo. If a new fertilization [occurred], the progeny would result BS-RNase+. ''
2. Intersubunit Disulfides The double covalent linkage (Di Donato and D'Alessio, 1973) between subunits is another unusual feature of the structure of BS-RNase; it is unusual not only for an RNase, but also in general for an enzyme or a small protein. It could be assumed that two solid, covalent bonds were better fit to protect the evolutionary advantage of a dimeric RNase than were noncovalent forces. On the other hand, although covalent, a disulfide bond can be easily cleaved in a controllable fashion by enzymes or by suitable concentrations of cellular or extracellular thiols. In fact, the intersubunit disulfides of BS-RNase are very sensitive to reduction, so much so that they are instantly reduced by approximately stoichiometric concentrations of dithiothreitol at room temperature (D'Alessio et al., 1975). Under these conditions the four intrasubunit disulfides present in each subunit remain completely resistant to dithiothreitol. A kinetic analysis of the reactivity toward model disulfide compounds of Cys-31 and Cys-32 monomeric BS-RNase showed two- to fourfold higher reaction rates than those determined for free cysteine (Parente et al., 1985). This superreactivity was found to be due to both the adjacency of the two cysteins in the peptide chain and to the presence of nearby positively charged groups, those of Arg-33 and Lys-34. Thus, it can also be hypothesized that the intersubunit disulfide bonds of BS-RNase, rather than strengthening the dimeric structure, may have instead the function of permitting a controllable dissociation-association of the protein monomers. In this respect, it should be noted that the monomeric derivatives of BS-RNase are more active catalytically than the parent dimeric enzyme (see Section IV,A), but do not possess the special, noncatalytic actions of native, dimeric BS-RNase (see Section IV,B).
3. Deamidation of Asn-67 In the m R N A encoding the BS-RNase subunit chain, the triplet coding for the residue at position 67 designates an Asn (Palmieri et al., 1985; Preuss et al., 1990). But in about 25% of the subunits of BS-RNase, as isolated from seminal plasma or seminal vesicles by a variety of methods,
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an Asn is not to be found at that position, because it is deamidated. This was determined when purified BS-RNase was separated by ionexchange chromatography into three components, each with different subunit compositions: a2, a/3, and/32. The B-type subunit was found to have a full complement of amide groups, while the c~ type is selectively deamidated at Asn-67 (Di Donato and D'Alessio, 1981; Di Donato et al., 1986). The three subforms of BS-RNase may not represent typical isoenzymes, in that they share the same functional properties, both catalytic (Piccoli et al., 1988) and noncatalytic (G. D'Alessio, unpublished data). The finding that newly synthesized BS-RNase is identified with the /32 subform (Quarto et al., 1987) indicates that the selective deamidation occurs, as a spontaneous or catalyzed process, during the prolonged storage of the protein in the seminal vesicle gland. Hence the physiological significance of the selective deamidation of Asn-67 in BS-RNase must be related to the aging of the protein. In fact, the description of the selective deamidation of Asn-67 as a precise molecular process coincident with the aging of BS-RNase was a direct and definite indication, later confirmed for crystallins by the findings of Voorter et al. (1987), that selective deamidation can be a molecular signal to mark aged proteins: a "biological clock," as proposed by Robinson and Rudd (1974). In this respect, it should also be noted that although an Asn-67 is present in all known pancreatic RNases, with the exception of turtle pancreas RNase, which can be assigned to a distinct subfamily (Beintema et al., 1988), deamidations have been detected in RNases only as a result of some severe treatment (Beintema et al., 1982). In native RNase A, Asn-67 deamidates at a very low rate (Di Donato et al., 1993), which approximates that of Asn-67 in BS-RNase only under rather harsh conditions (Manjula et al., 1976) or after denaturation of the protein (Thannhauser and Sheraga, 1985). The striking difference in deamidation rates of Asn-67 in native BSRNase and RNase A is surprising; even more surprising is the finding that in monomeric, stable, and active BS-RNase the deamidation rate of Asn-67 is much lower than in the native dimeric enzyme (Di Donato et al., 1986), and very similar to the rate measured for RNase A (Di Donato et al., 1993). Clearly, no primary structure determinants can be responsible for these effects, so that one has to speculate (1) that in solution, just as has been observed for the protein in the crystalline lattice (Mazzarella et al., 1993), the loop 65-72, containing Asn-67, can adopt two different interconvertible conformations, one of them identical to the conformation of the same loop in RNase A; and (2) that the
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conformation that is different from that of RNase A creates a more acidic or a more basic microenvironment, either of which could favor a higher deamidation rate. The mechanism of deamidation of Asn-67 of BS-RNase was determined to be based, as first proposed by Bornstein and Balian (1977), on the formation of a cyclic succinimide intermediate. Because the cycle can then be hydrolyzed on either side of the imide nitrogen, both Asp and isoAsp residues are generated at the site of deamidation. In fact, conclusive evidence for this mechanism was obtained studying the case of BS-RNase, with the first direct demonstration of the presence of isoAsp in the deamidated protein (Di Donato et al., 1986). It has also been shown that the isoAsp residue generated through deamidation of Ash-67 of BS-RNase in an isolated peptide fragment, or in the denatured protein, is a good substrate for protein carboxyl Omethyltransferases (Di Donato et al., 1986). Methylation occurs at the free a-carboxyl of isoAsp, and the methyl group can then spontaneously hydrolyze. It has been demonstrated that repeated cycles of methylationdemethylation can "cure" a peptide of the anomalous isoAsp structure (Galletti et al., 1988). However, because no methylation was found to occur in the native selectively deamidated protein, the physiological significance of a methylation of the a-carboxyl of isoaspartyl residues in BS-RNase remains to be ascertained.
B. Three-Dimensional Structure
1. X-Ray Investigation of BS-RNase Structural details of BS-RNase have been obtained by an extensive X-ray diffraction analysis of the native enzyme (Capasso et al., 1983; Mazzarella et al., 1993). The coordinates have been deposited with the Brookhaven Protein Data Bank (PDB) (Bernstein et al., 1977). The PDB file name is 1RBS. Crystals, grown from concentrated ammonium sulfate at pH 5.1, diffract to at least 1.9 A resolution. They are orthorhombic with space group P22121, and unit-cell parameters a = 36.5, b = 66.7, c = 107.5 A, Z = 4. There is one dimer in the asymmetric unit, therefore the potential twofold symmetry of the molecule is not a symmetry element of the crystal. This means that the two subunits are not forced to be identical
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and differences between them may give valuable information regarding the flexibility of the molecule and may possibly suggest the structural source of the half-site reactivity of the enzyme (see Section IV,A,2). A first model of the seminal enzyme was based on a low-resolution isomorphous electron density map; the interpretation of the map was greatly facilitated using as a guide the structure of RNase S. As expected, the two chains of the model were related by a dyad axis almost parallel to the c axis, in agreement with pseudosystematic absences of some loworder reflections; these absences can be noted by visual inspection of the diffraction pattern and can be interpreted in terms of a pseudo-I222 space group symmetry. A stepwise rebuilding of the chains, alternated with several cycles of stereochemically restrained least-squares refinement, produced a model with good stereochemical parameters and a low value of the overall disagreement index R between observed and calculated structure amplitudes. However, a most fundamental aspect of the structure concerning the conformation of the peptide 16-21 of the two chains was still unsatisfactorily modeled, because the electron density map was not clearly defined in this region (Capasso et aL, 1983). For this reason a new data set was collected on an area detector, which allowed pushing the resolution further to 1.9 A and a more accurate list of observed structure amplitudes; in particular, the number of accurately measured reflections [/obs > 3~r(1)] was almost doubled. Using this new data set, a second examination of the resultant BS-RNase model was performed using 2F,,-Fc and F,,-Fc omit Fourier maps. Revisions at this point were minor, in general involving side chains and the structure of the bound solvent. Electron density was better defined and also allowed a more precise definition of the structure of the 16-22 peptide. The revised model was subjected to further refinement in which the contribution of the disordered solvent was added to the calculated structure factors. The final standard crystallographic R was 17.7% on 16,492 reflections, with I > 3~r(1). Throughout the refinement, restraints related to the local twofold symmetry were not applied (Mazzarella et al., 1993).
2. Structural Unit of BS-RNase As expected on the basis of the high level of sequence similarity, the secondary structure of BS-RNase is very similar to that of RNase A. It presents, however, a marked difference in region 16-22, which determines a completely different orientation of the N-terminal peptide, thus altering the tertiary structure of the chain and conferring a unique feature
Giuseppe D'Alessio et al.
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to the overall structure of the seminal enzyme. In order to simplify the discussion, let us partition each chain (called S1 and $2, respectively) into a tail T (residues 1-15), a body B (residues 23-124), and a hinge peptide (residues 16-22). Quantitative comparisons of the seminal and pancreatic enzymes have been performed using coordinates of the phosphate-free structure of RNase A (Wlodawer et al., 1988) refined at 1.26 A resolution (PDB file name 7RSA). The two models of ribonucleases were best superimposed using the B core (N, C a, C, and O backbone atoms of residues 25-35, 40-60, 72-86, 95-110, and 116-123), which includes almost all residues in the helical and/3-sheet regions of the B domain. The C a atom model of seminal ribonuclease is shown in Fig. 2; the local molecular dyad axis is vertical and the two chains, S1 and $2, have different colors. They are linked by two interchain disulfides that form a 16-membered cycle. The model of the seminal enzyme can be divided into two parts (left and right with respect to the noncrystallographic dyad axis), each including a T and a B moiety of two different chains, T1/B2 or T2/B 1. These parts are called the structural units of the seminal enzyme. Aside from the loop 65-72, the two structural units are identical within experimental error, as can be seen from the plot in Fig. 3, which shows the root-mean-square (rms) deviation per residue between T1/
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Residue number Fig. 3. R o o t m e a n square deviation (rmsd) b e t w e e n the two structural units of BSRNase M• form as a function of the residue n u m b e r .
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B2 and T2/B1 as a function of the residue number. The two units have been superimposed using othe B core residues, and the rms deviation for these residues is 0.19 A (0.30 A for residues 2-15, not used for the superposition). As for RNase A, the folding of the structural unit can be classified into the type c~B roll of the ~ § class (Orengo and Thornton, 1993), and is characterized by a central core of/3 structure surrounded by three helices, H1, H2, and H3. The tail T is partially folded as an c~ helix (H1), which spans residues 3-13; the essential His-12 is located at the C end of the helix. H2 begins at Asn-24, has normal c~-type 5----> 1 hydrogen bonds, and ends with a 6 ~ 1 (~ type) and a 4 --> 1 bond between residues 30 and 35 and 31 and 34, respectively, with the C-terminal residue Lys34 in the Lc~ conformation. This pattern of H bonds, commonly found at the C terminus of an c~ helix (Schellman, 1980), is also present in RNase A, which has in position 34 an Asn, a residue frequently found in the Lc~ conformation. Interestingly, the replacement of Asn for Lys, which only rarely adopts a left-handed conformation (Richardson and Richardson, 1989), does not alter the C-terminal feature of the helix. The third helix H3 (residues 50-60) starts as an c~ helix and ends with one turn of a 3~0 helix. Apart from the helices H2 and H3, the body B is formed of an extensive /3 structure comprising two antiparallel /3 sheets: one sheet has three strands spanning residues 42-48, 79-91, and 94-104, with residues 88-89 bulging out. The second sheet is more irregular and is formed by two strands encompassing residues 105-113 and 114-124, flanked on one side by two short strands (residues 61-64 and 71-75, respectively). Although the long/~-strand 94-113 is involved in the formation of both sheets, the two sheets roll up at the level of residues 104-105 to form a partial barrel, which, together with the N-terminal c~ helix H1, delimits a cleft where the substrate binds. The two sheets are also joined by the helix H3. With the exception of G l n - l l and His-12, all residues known to be important for the catalytic activity of the enzyme are located close to or within the/3-sheet structure: Lys-41 is at the beginning of the first sheet, which includes Asn-44 and Thr-45; the essential His-119, Asp121, and Ser-123 belong to the second sheet. The pyrimidine-preferring binding site B1 (Richards and Wyckoff, 1971) is located between the two sheets, whereas the purine or pyrimidine site B2 is formed only by residues of the second sheet; the binding site p l of the phosphate, involved in the catalytic breakdown of the phosphodiester bond, is located between the two histidines.
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Of the four proline residues also present in pancreatic ribonuclease, Pro-42 and P r o - l l 7 are in the trans configuration, whereas Pro-93 and P r o - l l 4 are located in sharp turns of the first and second sheet, respectively, and adopt the cis configuration. Pro-19 (Ala in RNase A) is an important structural determinant for the swapping of the N termini between subunits. The four intrachain disulfides are conserved in both the seminal and the pancreatic enzyme and present closely related structural features: two of them (Cys-26/Cys-84 and Cys-58/Cys-110) fix firmly the two helices H2 and H3 to the ends of the/3 core and are left-handed (g3 "~ -90~ a third bond (Cys-40/Cys-95) is also left-handed and connects two exposed loops. The fourth disulfide bond between Cys-65 and Cys-72 is characterized by a right-handed conformation (g3 "~ 90~ with similar parameters in both subunits, though the structure of the loop 65-72 is different. In the chain S1 it adopts the conformation found in RNase A, whereas a similar conformation for $2 would give rise to serious overlaps with atoms of the same loop of S1 of a screw-related molecule. These packing interactions may then justify the different conformation adopted by the $2 loop. It must be recalled that this loop includes the most labile Asn67, a residue that deamidates selectively (see Section III,A,3) via a succinimide derivative, to an c~-linked or a/3-1inked Asp (isoaspartyl) residue (Di Donato et al., 1986). The structure of the isoaspartyl derivative of RNase A, recently determined at high resolution (Capasso et al., 1996), shows a large rearrangement of the loop 65-72 on the insertion of the extra methylene group in the main chain; this loop is shifted toward the main body of the molecule in the region of His-ll9, with the consequence of reducing slightly the space available to the base in the purine-binding site. The conformation of the $2 loop of BS-RNase resembles that found for the isoaspartyl derivative of RNase A at least for residues 67 to 69: on the other hand, a chromatographic analysis of dissolved crystals of the seminal enzyme has indicated the presence of a conspicuous amount of the c~/3 subform (see Section III,A,3). The electron density map in this region is somewhat less clearly defined with respect to other regions and, on the basis of the previous considerations, it may actually represent an average of the amidated and deamidated subforms. A detailed scheme of the hydrogen bonding network between main chain atoms is shown in Fig. 4. Except for a few additional hydrogen bonds, the major difference between this and a similar diagram for RNase A is that the T and B domains belong to different chains. A
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Fig. 4. Backbone hydrogen bonding scheme for the structural unit of the quaternary form M x M of BS-RNase. Residues 1-18 and 18-124 belong to two different chains. Bonds shorter than 3.2 ,& are indicated by solid lines; those between 3.2 and 3.35 ,~ and with N - - H - - O angles greater than 110~ are indicated by broken lines. Sequence differences compared to RNase A are indicated by shaded circles.
quantitative comparison with pancreatic ribonuclease (Wlodawer et al., 1988) shows that, with the exception of the hinge peptide and of the loop 65-72 of $2, the differences are well within those observed among various models of RNase A. The rms deviation for the B core is 0.30 and 0.31 A for the T2/B 1 and T1/B2 unit, respectively; the rms deviation
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for residues 2-15 (0.39 A for T2 and 0.56 A for T1) decreases only slightly when these residues are used for superposition, indicating that the T/B spatial relationship in RNase A is practically undistinguishable from that found in the two structural units of BS-RNase. The largest differences (> 1 ,A,) are observed for residues 37-38, 113-114, and 87-95. In the first case the two residues are located in a partially exposed short loop that connects the end of helix H2 with the B-sheet structure. Despite the large number of residue substitutions between the seminal and pancreatic enzyme in this region, the structure is highly conserved, except for residues 37-38, where the sequence Lys-Asp is replaced by Gln-Gly in the seminal enzyme. In RNase S (Kim et al., 1992) the peptide plane 37-38 has a different orientation with respect to RNase A, and in BSRNase it is flipped over by almost 180 ~ With this orientation of the peptide plane, the conformation of residue 38 is allowed for glycine; in addition, the CO group of Gin-37 can make a hydrogen bond with the NH group of Gly-ll2 of a screw-related molecule. These packing interactions may cause the differences observed for residues 113-114. Residues 87-95 form an exposed turn between the second and third strand of the first/3 sheet and are also in close contact with the loop 35-40. Differences greater than 1 A are also observed for the first two residues of the chain, which are involved in different packing contacts. 3. Quaternary Structure a. X - R a y Structure o f M • The structural unit of the seminal enzyme is very similar to RNase A, although in the former the interacting T and B moieties belong to different chains. That is, the N-terminal segment of each chain is detached from the position occupied in RNase A, where it interacts with the body of its own chain, is rotated 180 ~ about the molecular dyad axis, and takes over the equivalent position on the body of the second chain. In order to underline this unique feature of the solid-state structure of the seminal enzyme, the dimer has been termed M x M to distinguish it from the dimer M - - M , in which no swapping occurs (Piccoli et al., 1992). The swapping of the T domain between subunits necessarily requires a new folding of the hinge peptide with respect to the pancreatic molecule. Its conformation in BS-RNase is also different for the two chains; in both cases, Pro-19 is trans and is positioned at the first corner of a type I/3 turn stabilized by a 4--> 1 hydrogen bond between the carbonyl group of Ser-18 and the amide group of Ser-21; however, the orientation of the turn in the two chains
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is considerably different. At the end of the hinge peptide, Ser-21 through Ser-23 form one turn of a 310 helix, which then proceeds with the helix H2. The quaternary structure of the seminal enzyme is stabilized by the interchain disulfides between Cys-31 and Cys-32 of one chain with Cys32 and Cys-31 of the second chain, respectively; the two bonds form a 16-membered cycle that concatenates the C ends of the H2 helices of the two subunits. The relative orientation of the helices, which form an angle of approximately 40 ~ leaves space for a favorable interaction between the side chains of Leu-28 of the two chains; this residue, which replaces a Gin in pancreatic ribonuclease, is exposed to the solvent in this molecule and becomes buried at the subunit interface in the seminal enzyme. Despite this hydrophobic interaction, the contact area provided by the interactions between the two helices would not be large enough to stabilize the dimer in the absence of the covalent linkages. Because of the extensive T/B interactions (Fig. 2), the swapping of the N-terminal segments produces a large increment of the subunit interface, which justifies the persistence of a dimeric species in solution when the interchain disulfides are selectively reduced. The two structural units T1/B2 and T2/B 1 are related by a 180~ rotation about the molecular axis, plus a small but significant translation of--~0.4 parallel to the same axis. For the M x M dimer this implies that, after superposit!on of the two bodies, the two tails are consistently shifted by about 0.8 A relative to each other in the direction of the molecular axis. This shift is effective in maintaining the T1/B2 and T2/B1 domains in the same relative position found in RNase A: on the other hand, it necessarily requires a readjustment of the hinge peptide structure and may well explain the different conformation adopted by this peptide in S1 and $2. The swapping of the T domains has another interesting consequence: each active site, located at the T/B interface, is actually formed by residues belonging to different chains, and, in turn, each chain is involved in both active sites. Therefore, the departure from the exact twofold symmetry can also affect the structure of the active sites and may provide a plausible mechanism for the allosteric properties displayed by the M x M dimer (see Section IV,A,2). A recently performed energetic analysis of the hinge peptide (Mazzarella et aL, 1995) successfully predicts the two conformations found for this peptide in BS-RNase and also indicates that the peptide is scarcely prone to small adjustments, which would prevent the propagation of structural modifications from one active site to the other.
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The interchange of the N termini is the result of a preordinated scaffold that places in the correct orientation and position the N-terminal end of the helix H2 with respect to the C terminal end of the helix H1 of the second chain; the interchain disulfides provide a first important anchorage of the two chains. The interactions across the molecular axis between the side chains of Leu-28, sticking out from the helix H2 one turn of the helix from the interchain disulfide bridges, are also important for a fine adjustment of the relative position of the bodies of the two chains. Within this well-defined quaternary structure, the swapping of the T domains depends on the relative propensity of the hinge peptide to adopt either of the two conformations characteristic of RNase A and of the M x M dimer of BS-RNase. In this context, the replacement of Ala for Pro at position 19 of RNase A plays a key role in the swapping. This conclusion is supported by the results of energy calculations (Mazzarella et al., 1995) that indicate that the replacement of Pro for Ala in an RNase A mutant induces a more external position of the hinge peptide with a reduced number of interactions with the rest of the protein. The trans form of Pro was also predicted to be more stable in the mutant than the cis form. Moreover, the NMR analysis of the monomeric species of BS-RNase has shown that its tertiary structure is very close to that of RNase A, and that the hinge peptide, with Pro-19 in the trans configuration, does not display definite signals indicative of strong interactions with the protein matrix (D'Ursi et al., 1995). This finding is in line with the observation that the monomeric derivative is more sensitive than native seminal and pancreatic enzymes to thermal denaturation, and that dissociation of BS-RNase into monomers leads to exposure of a tyrosine residue (Grandi et al., 1979). Indeed, Tyr-25, deeply buried in the seminal enzyme, is well protected in RNase A by the tightly bound hinge peptide and may become more exposed in the monomeric species of BS-RNase if the latter peptide assumes a more external position.
b. D u a l Quaternary Structure o f B S - R N a s e . The M x M quaternary structure, determined by X-ray studies and described in Section III,B,3,a, although prevalent, is not the only quaternary structure for BS-RNase. The protein also has access to a conformation denoted as M - - M, adopted by about one-third of the protein molecules, in which no swapping of N-terminal segments between subunits takes place (see Fig. 2). Under the conditions employed for crystallization of the protein, only the M x M form was found to be present in the crystals (Piccoli et al., 1992). Thus,
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evidence for the existence of the M - - M form was acquired through biochemical methodologies in solution (Piccoli et al., 1992). One of the lines of evidence, and an effective method for separating the two quaternary forms of BS-RNase, stems from, and explains, the early finding (D'Alessio et al., 1975) that when the intersubunit disulfides of BS-RNase are cleaved, only one-third of the protein is recovered as monomeric on gel filtration, whereas the remaining two-thirds elutes as noncovalent dimers. It is now clear that on cleavage of the intersubunit disulfides the M - - M form readily dissociates into monomers, whereas in the M • form the noncovalent interactions between the N-terminal a helix of each subunit and the main body of its partner hold together the two subunits in a noncovalent dimeric association. In fact, on air reoxidation of the intersubunit disulfides, the dissociated monomers and the noncovalent dimers reform only the M = M or the M x M type of dimer, respectively (Piccoli et al., 1992). The M• M dimers of BS-RNase are closely reminiscent of, and represent the "natural" counterpart of, the noncovalent artificial dimers of RNase A described by Crestfield et al. (1962). When their classical pioneering experiment, which revealed the interchange of N-terminal segments between monomers of aggregated RNase A, was reproduced on BS-RNase, further evidence was obtained on the existence of the M - - M subform. The experiment consisted of the aggregation of two types of monomers, each inactivated by alkylation of one of the two active site His residues: His-12, located in the interchangeable N-terminal segment, and His-ll9, located in the main body of the monomer. The observed recovery of activity on association into dimers of the two inactive monomers could only result from the formation of composite active sites, through interchange of the N-terminal segments between monomers. However, quantitation of the recovered activity showed that a fraction of the monomers associated without interchange (Piccoli et al., 1992). The two quaternary structures of BS-RNase have different functional properties. The M - - M form, without exchange, seems to be capable only of RNA degradation. The finding that it has no allosteric properties (Piccoli et aL, 1992) supports an earlier proposal that the two composite active sites of the M • M structure form the molecular basis for site-site communication, hence for the allosteric properties of BS-RNase (Piccoli et al., 1988; see Section IV,A,2). Furthermore, it has been found that the M x M form is the most, and possibly the only, quaternary form endowed with antitumor action (Cafaro et al., 1995; Di Donato et al., 1995; Kim et aL, 1995a) and with aspermatogenic action (Kim et al., 1995a).
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The intriguing finding that BS-RNase, as isolated by a variety of purification procedures, consistently contains the M x M and M - - M forms in a constant 2:1 ratio was explained when the isolated forms were incubated under pseudophysiological conditions, at pH 7 and 37~ Each form was found to convert into the other until an equilibrium was reached, when about two-thirds of the protein was in the M x M "exchanging" conformation and one-third was in the M - - M form (Piccoli et al., 1992), which is the equilibrium ratio observed for M x M and M - - M in native BSRNase. The process is extremely slow, with half-times of the order of days, but more recent data (Murthy et aL, 1996) indicate that the interconversion process is much faster under reducing conditions, i.e., when the intersubunit S-S bridges are cleaved and the equilibrium species are noncovalent dimers and free monomers. This is not surprising, considering that the reciprocal movement of the displaced N-terminal arms, moving from one subunit to the other, may be facilitated by the absence of the covalent constraints created by the two disulfides at the intersubunit interface. Two different approaches have been exploited for analyzing the structural determinants responsible for the exchange of the N-terminal helices in the M x M conformation. Kim et aL (1995a) substituted a Ser residue for Cys-31 or Cys-32, the residues responsible for the intersubunit disulfides of BS-RNase. They found that the engineered reconstructed dimers, each with a single intersubunit disulfide, were mostly M - - M dimers, with M x M dimers present to a much lesser extent than in native BS-RNase. Thus the copresence of both disulfides covalently linking the two subunits to each other has an impact on the ability of BS-RNase monomers to exchange their N-terminal domains. Di Donato et al. (1994, 1995) followed an alternative approach. By site-directed mutagenesis they equipped monomeric RNase A with significant residues from the BS-RNase sequence. When two cysteine residues were inserted in the RNase A sequence at positions 31 and 32, substituting for a Lys and a Ser, respectively, the dimer obtained had a significant, albeit modest, propensity to exchange the N-terminal domains. However, when either a Leu at position 28 or a Pro at position 19 was also substituted for the original Gin and Ala, respectively, of the RNase A sequence, the fraction of the M x M form at equilibrium in the dimeric RNase A mutant was significantly higher, and even higher when all four substitutions (Cys-31, Cys-32, Leu-28, and Pro-19) were introduced. In the latter case the same equilibrium ratio between the M x M and the M - - M form was the same as that found for native BS-RNase (Di Donato et al., 1994, 1995).
12 Seminal RNase: Importance of Diversity
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The general conclusion that can be reached from these studies is that one intersubunit disulfide is enough for the making of a dimeric RNase of the vertebrate superfamily. However, to make a dimeric RNase with a high propensity toward stable N-terminal exchanges, such as BSRNase, a tight interface, stabilized by two adjacent disulfides and by the hydrophobic contact at Leu-28, is necessary. This would provide a good fulcrum for the levering up, and then into position, of the exchangeable a helices. The Pro at position 19 would then lower the energy of the "exchange" conformation of the hinge peptide linking the N-terminal a helix to the H2 a helix.
C. Folding Pathway
Refolding studies of fully denatured and reduced BS-RNase have been carried out following two experimental approaches, both based on the reformation of correctly paired disulfide bondsmas facilitated by a glutathione redox system (Smith et al., 1978) or by air regeneration (Parente and D'Alessio, 1985). With the former system no dimeric protein is obtained, not even traces, and the unfolded chains refold into monomers with Cys-31 and Cys-32 blocked by glutathione moieties into mixed disulfides. In fact, whether glutathione is used to assist the reformation of intrasubunit disulfides, or to dissociate the native dimer (Smith and Schaffer, 1979) through selective cleavage of the intersubunit disulfides, in the presence of a glutathione redox buffer the stable form of BSRNase is monomeric. Also, in air regeneration experiments, BS-RNase chains first reorganize into monomers, but then, albeit very slowly and with a rather low yield, a fraction of the protein associates into nativelike dimers (Parente and D'Alessio, 1985). These two sets of findings appear contradictory, and even more surprising (Jaenicke, 1987) is the finding of an oligomeric protein for which a stable, active monomeric form is recognized. However, contradictions and surprises may be dealt with by considering both the evolutionary history of the protein and its presumable refolding pathway in vivo. If we accept that the BS-RNase ancestor was monomeric (as are all its offshoots except BS-RNase), the first step in the refolding pathway of BS-RNase in the cell may be reminiscent of its previous evolutionary experience as a monomer. This step would consist in the refolding of a monomeric intermediate, assisted by a microsomal redox system based
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on glutathione or cysteine (Hwang et al., 1992). As described in Fig. 5, which illustrates a hypothetic refolding pathway for BS-RNase in vivo, the product of this first step would be monomeric BS-RNase with mixed disulfides between Cys-31 and Cys-32, and glutathione or cysteine. The second step, consisting of the deblocking of Cys-31 and Cys-32 and of their orderly pairing into intersubunit disulfides, may require an environment with a different redox potential, and probably the assistance of a protein disulfide isomerase, as well (Freedman, 1984). An earlier proposal of a refolding pathway for BS-RNase (Parente and D'Alessio, 1985) must be redesigned after the findings: (1) that BSRNase has access to two quaternary conformations, and that freshly refolded dimers are in the "nonexchanging" M - - M conformation (Piccoli et al., 1992) (see Section III,B,2); (2) that the bond between Ser-18 and Pro-19 is a trans bond also in BS-RNase monomers (D'Ursi et al., 1995). Thus, in the scheme illustrated in Fig. 5, in the third and final step of BS-RNase refolding, the M - - M dimers partially transform into, and then equilibrate with, the M x M form.
D. Chemical Modifications and Engineering 1. M o n o m e r i c D e r i v a t i v e s
Due to the high, differential sensitivity to thiol reagents of BS-RNase intersubunit disulfides (see Section III,A,2), folded and stable monomeric derivatives of BS-RNase can be readily prepared by treatment of the dimeric protein with a moderate excess of dithiotreitol (D'Alessio et al., 1975). As described in Section III,B,3, only about one-third of the protein is monomerized as M(SH)2 (where M stands for monomer and the two sulfhydryls are those of Cys-31 and Cys-32). Most of the protein, being in its M x M form, on cleavage of the intersubunit disulfides, remains a noncovalent dimer. M(SH)2 is quite stable at pH 5 and can be firmly stabilized as a monomer either by alkylation of the exposed sulfhydryls with iodoacetic acid, iodoacetamide, or ethyleneimine (D'Alessio et al., 1975; Parente et al., 1977), or by reaction with glutathione (Smith and Schaffer, 1979) or with thiosulfonate reagents (Tamburrini et al., 1989). With the latter procedures, two mixed disulfides [M(SSR)2] are formed between half-cystine-31 and -32 and glutathione,
Unfolded chains +
R-SH/R-SS-R
I M(SSR)~[ R-SH and/or Factor(s)
I ~ ~ ' 2 R-SH
I M=MI N-terminal exchange
MxMI Fig. 5. A model for the refolding of denatured and reduced BS-RNase chains into the dimeric M • and M - - M forms.
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or the thio moiety of the thiosulfonate reagent. This gives the advantage of reversibility over the alkylation procedures, because the mixed disulfides are as differentially labile with respect to the intrasubunit disulfides as the original intersubunit disulfides (Tamburrini et al., 1989; Piccoli et aL, 1992). All monomeric derivatives of BS-RNase are more active catalytically than the native dimeric enzyme, with the M(SH)2 derivative being the most active. The basis for this superreactivity is discussed in Section IV,A,2. By contrast, monomeric derivatives have no detectable special biological actions (see Section IV,B), unless special conditions are employed (Wu et al., 1995). 2. Hybrid, Monofunctional, Superactive D i m e r As described above, the stable monomers M(SSR)2, produced by the reaction of BS-RNase with thiosulfonate reagents (see above and Fig. 5), can be readily reassociated into nativelike dimers by selective reduction of the mixed disulfides, followed by air-facilitated reformation of the intersubunit disulfides. This procedure can also be used for the production of a hybrid dimer, by performing the reassociation step on a mixture made up of equimolar amounts of M(SSR)2 and of catalytically inactivated M(SSR)2, as obtained by carboxymethylation of an active site His residue. This hybrid derivative of BS-RNase, structurally dimeric but functionally monomeric, has been prepared and found to have a surprisingly high catalytic activity and no allosteric properties (Tamburrini et al., 1989). These results were tentatively explained by proposing that the carboxymethyl group linked at His-119 mimicked the binding of a quasi-substrate at the active site, thus freezing the enzyme in its active conformation (see Section IV,A,2).
IV. FUNCTIONS
A. Catalytic Function BS-RNase is virtually undistinguishable from RNase A when its main enzymatic parameters are analyzed (Floridi et al., 1972; Irie and Hosokawa, 1971). The internucleotide bond is cleaved in two steps: a trans-
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phosphorolytic step, with the intermediate formation of 2',3'-cyclic nucleotides, followed by hydrolysis of the resulting cyclophosphates in the rate-determining step. The bond specificity is also the same as that of RNase A, with a strong preference for phosphodiester bonds linking pyrimidine nucleotides on the 3' side. It has been reported that BSRNase can cleave also polyadenylic chains, but does not hydrolyze adenosine 2',3'-cyclic phosphates (Floridi et al., 1972). A major difference between BS-RNase and RNase A, and all other mammalian RNases, is its resistance (Murthy and Sirdeshmukh, 1992) to the protein RNase inhibitor (Lee and Vallee, 1993). The reason for this resistance has been presumed to be its dimeric structure, given the inhibitor sensitivity of monomerized BS-RNase (Murthy and Sirdeshmukh, 1992). This hypothesis has been definitely confirmed by the illuminating X-ray structure of the inhibitor complexed to RNase A (Kobe and Deisenhofer, 1995), which shows that a dimeric RNase would just not fit in the horseshoe structure of the protein inhibitor. The inhibitor insensitivity has been demonstrated for both M • and M - - M forms, as well as for the noncovalent dimers, produced after cleavage of the intersubunit disulfides (Murthy et al., 1996). 1. A c t i o n
on Double-Stranded
RNA
and on DNA
9R N A
Bovine pancreatic RNase A is virtually inactive on double-stranded RNA (Barnard, 1969) under pseudophysiological conditions of ionic strength and pH, which stabilize the secondary structure of the nucleic acid. Under the same conditions, BS-RNase can effectively degrade double-stranded RNA and poly(A), poly(U) complexes (Libonati and Floridi, 1969) and the RNA strand of D N A . RNA hybrids (Taniguchi and Libonati, 1974). Interestingly, dimers of bovine pancreatic RNase A obtained by lyophilization (Libonati, 1969), as well as by cross-linking with bisimido esters (Bartholeyns and Moore, 1974), are able to degrade double-stranded RNA and DNA 9RNA hybrids (Libonati et al., 1975a). However, the correlation between the dimeric structure of these RNases and their ability to degrade double-stranded structures was soon recognized as merely apparent, when a monomeric derivative of BS-RNase and monomeric whale pancreatic RNase were found to be capable of degrading double-stranded RNA (Libonati et al., 1975b; Libonati and Palmieri, 1978). Further investigations led to the proposal that the degradation of double-stranded structures can be explained in terms of destabilization of the polynucleotide secondary structure by RNases with a
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higher charge density (Libonati et aL, 1975a), in discrete regions of the protein (Sorrentino and Libonati, 1994). The hypothesis finds support also in the finding that BS-RNase destabilizes double-stranded DNA (Libonati and Beintema, 1977; Pandit and Ramakrishna, 1986) and poly[d(A-T), d(A-T)] (Pandit and Ramakrishna, 1986). The physiological or biological relevance of the action of BS-RNase on double-stranded RNA structures is not clear. However, it has been reported (Schein et aL, 1990) that y-interferon (IFN-y) activates the degrading action of BS-RNase on double-stranded RNA. These data can be related to the reported finding that a double-stranded specific RNase is coinduced in cells treated with interferon-inducing factors (Meegan and Marcus, 1989), and might be of interest, given the role of interferon in the induction of an antiviral state. A relationship has been found between the BS-RNase activating effects of IFN-), molecules in which the positively charged C-terminal segment is missing, such as the murine IFN-y and a proteolytic fragment of human IFN-),, and their antiviral action (Schein and Haugg, 1995). Furthermore, antibodies capable of neutralizing the antiviral action of human IFN-y were found to block its ability to activate the BS-RNase degradation of doublestranded RNA. 2. AIIosteric Properties
BS-RNase is an allosteric enzyme, and to date the only RNase for which allosteric behavior has been described; however, it is difficult to conceive of a physiological significance for this property of the enzyme, given its high abundance in semen, and in the gland where it is produced, and its unlikely involvement in any metabolic pathway. Although seminal RNase cleaves susceptible dinucleotides, as substrates of the first step of its reaction, with classical Michaelian kinetics (Piccoli et al., 1982), the hydrolysis of the 2',3'-cyclic phosphates (products of the first step, and substrates of the second, rate-limiting step) occurs in a rather complicated kinetic fashion (Piccoli et al., 1982, 1988). At low concentrations, the substrate induces a negative cooperativity; at higher concentrations, a positive cooperativity. This results in a mixedtype cooperativity, with a pronounced "bump" in the substrate saturation curve, located in the concentration range between the positive and the negative cooperativity regions (see inset of Fig. 6). An extensive examination of the reaction over three orders of magnitude of substrate concentration, followed by analysis of the data with the Hill equation, confirms
411
12 Seminal RNase: Importance of Diversity
0.S
h - 0.87
>
//
-0.S
h-
I
1.27
> O
m
-1.0
h- o.s8 e,S.0
-1 .S
h - 0.99
~z.s I.IJ
>( N
0 -2.0 -4.0
-3.0
0
4 [S] (mM) -2.0
8 -1.0
log IS] (M) Fig. 6. A Hill plot of kinetic data obtained with BS-RNase and cytidine 2',3'-P (cyclic) as a substrate in the concentration range 0.1 to 100 mM. The values (h) of the Hill number calculated for the relevant segments of the curve are indicated. Inset: Data at low substrate concentrations showing the characteristic bump in the saturation curve.
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the different effects of the substrate at low (with a Hill number lower than 1) and high (with a Hill number higher than 1) concentrations, respectively, with the expected approach to unity of the Hill number at very high and very low substrate concentrations (Fig. 6). As anticipated by the allosteric theory, competitive inhibitors, such as the 3'-phosphate nucleoside reaction products, activate the enzyme at low concentrations and shift the substrate saturation curve toward higher activity and hyperbolicity (Piccoli et al., 1988). When tested in binding studies, these molecules titrate only one of the two structurally available active sites of the enzyme (Di Donato et al., 1987). Anomalous kinetics have also been reported for monomeric RNase A (Walker et al., 1975), but at much higher substrate concentrations (30-50 mM), compared with those (3-5 mM) arousing the allosteric behavior of seminal RNase. Comparative analyses of the kinetic behavior of BS-RNase and its monomeric form, and of RNase A and its dimeric derivative, as prepared by Crestfield et al. (1962), revealed that the monomeric RNases, natural or artificial, have no allosteric properties, whereas the dimeric RNases, natural BS-RNase and the RNase A dimers, both display the "bumpy" saturation curve (Piccoli and D'Alessio, 1984). However, the dimeric structure of BS-RNase and of the RNase A dimers was later shown to be essential, but not sufficient, to explain their allosteric properties, because another structural feature, common to both BSRNase and the RNase A Crestfield-type dimers, plays a more crucial role, i.e., that of possessing the M • conformation. BS-RNase in the M = M conformation has no allosteric properties; the M x M conformation instead, characterized by composite active sites, made up with one catalytic His from one subunit and the other from the partner subunit, appears to be suited for direct site-site communications, essential for generating homotropic substrate effects. (Piccoli et al., 1992). A model for the allosteric mechanism of BS-RNase has been proposed (Piccoli et al., 1988). The essential features of the model are as follows: (1) at low substrate concentrations, occupation of one of the two structural active sites results in a conformational transition, with the "switching off" of the second site, which produces negative cooperativity and half-site reactivity, and the "switching on" of noncatalytic binding sites, or subsites; (2) once these subsites are saturated, a second conformational change makes available to the substrate the second catalytic site, which engenders positive cooperativity. The model is supported by binding studies: those mentioned above, carried out with the 3'-nucleotide reaction products, and others, also
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showing half-site reactivity of the enzyme, performed with bromoacetyluridine, an affinity probe (Donadio et al., 1986). When the model was tested by computer simulation, using the available experimental constants and constants optimized by iterative methods, and a number of 4 subsites, derived by studies with RNase A (Pares et al., 1980), the resulting saturation curve fitted the experimental data very satisfactorily (Piccoli et al., 1988). It may be noteworthy to mention that the fit was lost when a number of sites lower or higher than 4 was used.
B. Special Biological Actions In the story of BS-RNase, the chapter on its biological actions is perhaps the most intriguing and volatile, because the protein is endowed with an astonishing array of biological activities and effects. As an enzyme, BS-RNase is by definition a biological effector, performing the biological action of RNA degradation, so that the other, noncatalytic bioactions of BS-RNase have been denoted as special biological actions (D'Alessio et al., 1991), and the acronym RISBASES has been proposed for all (Rlbonucleases endowed, like BS-RNase, with Special Bioactions). Although BS-RNase is not the only RNase for which unpredictable special biological actions have been uncovered, as reviewed in chapters 13, 14, and 15, it is unusual to find in a single RNase or protein so many and such diverse and disparate special bioactions: aspermatogenesis and antitumor, embryotoxic, immunosuppressive, and antiviral effects. The physiological role of BS-RNase is still mysterious, yet the available data allow some rational hypotheses. The first is based on the finding that when all BS-RNase is removed from bull seminal plasma, the plasma is deprived of its immunosuppressive activity (Tamburrini et al., 1990). All mammalian seminal fluids contain some immunosuppressive agents that aid in the suppression of immune responses of the female organism toward the nonself sperm cells. Thus the hypothesis was advanced (D'Alessio et al., 1991) that the physiological role of BS-RNase is that of an immunosuppressive agent; the hypothesis would also explain the high concentration of BS-RNase in bull semen, given the high dilution of semen in the recipient female tract. This hypothesis has also been considered by Kim et al. (1995a).
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Another hypothesis can be advanced based on the ability of BS-RNase to degrade both single- and double-stranded (see Section IV,A,1) R N A effectively and to bind D N A effectively (Libonati and Beintema, 1977). Given its high concentration in semen, even after dilution in the female tract, BS-RNase could actively degrade any R N A or complex any D N A that resulted from contaminating, decayed prokaryotes or eukaryotes. These activities could effectively protect the zygote from the potentially deleterious transforming effects of nucleic acids. The possibility should be considered that all special bioactions of BSRNase are in fact the reflection of some yet unknown action that produces different effects just because the action is exerted on different biological systems, e.g., the various assay systems that the investigators use to study the enzyme. This unitary hypothesis for BS-RNase biological function is suggested by the following findings: all special BS-RNase bioactions depend (1) on the integrity of the RNase function of the enzyme; (2) on the integrity of its dimeric structure; and (3) on the fraction of the protein in the M • M conformation. BS-RNase catalytically inactivated by alkylation of an essential His residue has no antitumor (Vescia et al., 1980) or immunosuppressive (Tamburrini et al., 1990) activities; BS-RNase mutants with an Asp residue substituting for the catalytically essential His-119 residue have no antitumor or aspermatogenic action (Kim et al., 1995b). Likewise, monomeric derivatives of BSRNase do not display any special activities: aspermatogenic (Kim et al., 1995a), antitumor (Vescia et al., 1980; Kim et al., 1995a), immunosuppressive (Tamburrini et al., 1990; Kim et al., 1995a), or anti-HIV (G. D'Alessio, P. La Colla, and R. J. Youle, unpublished). Finally, the M • M conformation of BS-RNase is important if not essential for its antitumor (Kim et al. 1995a; Cafaro et al., 1995; Di Donato et al., 1995) and for its aspermatogenic (Kim et al., 1995a) actions. An interesting effect of BS-RNase, that may not be properly described as a biological action, is that exerted on actin (Simmet al., 1987). BSRNase binds actin and promotes the bundling of the protein in orderly filaments. This effect appears to be interesting also because it is lost on dissociation of the protein into monomers, as happens for all bioactions of BS-RNase. Furthermore, this observation can be correlated to the finding that the angiogenin-binding protein on the surface of endothelial cells is an actinlike molecule (Hu et al., 1993). However, at least for the antitumor action of BS-RNase, no correlations were detected between the actin-binding effect of BS-RNase and its antitumor action (M. R. Mastronicola, unpublished results).
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1. Aspermatogenic Action As reported in Section I, Matou~ek and colleagues discovered the aspermatogenic action of seminal RNase while searching for an aspermatogenic factor that they had detected in bull semen (Dost~il and Matougek, 1972). One of the most distinctive characters of BS-RNase antispermatogenic action is the selectivity with which the enzyme exerts a potent cytotoxic action on the cell line that develops from spermatogonia, but not on the spermatogonia (Matougek and Grozdanovi~, 1973). On its administration to the animal by intratesticular injection, the testis seminiferous tubules become empty, as spermatids, spermatocytes, and mature spermatozoa disappear. But spermatogonia, which originate the sperm cell line, still form a single layer on the basal lamina of the tubules (Hlinak et al., 1981). Thus, when the protein is cleared from the animal, maturation of spermatogonia, with the eventual production of spermatozoa, can resume (Matougek and Grozdanovi~, 1973). This indicates that the aspermatogenic effect of BS-RNase is reversible. Furthermore, in treated animals no adverse effects are detected on the morphology of testis Leydig cells, nor on androgen secretion (Hlinak et al., 1981), which suggests that the hormonal testicular function is not influenced by the enzyme. In fact, when BS-RNase was administered to rats in a single intratesticular dose, the animals became infertile but their sexual behavior was totally unaffected (Hlinak et al., 1981). The finding that the aspermatogenic effect of BS-RNase is produced also when the protein is administered subcutaneously, although at higher doses (Matou~ek and Grozdanovi~, 1973), suggests that BS-RNase can overcome the blood-testis barrier. In this respect it is of interest that the aspermatogenic effect of BS-RNase can be exerted on several mammals, including mice, rats, rabbits, and rams (see Hlinak et al., 1981), but not on bulls (J. Matougek, personal communication). It should be noted that, given the high potential of this protein as a male contraceptive, and the earnest efforts that both research institutions and industry devote to the search for such agents, it is really surprising that the suggestive, promising lead contributed by the Czech group with the discovery of the aspermatogenic action of BS-RNase does not seem to have been followed by any other specialized investigators in the field.
2. Immunosuppressive Action That the RNase from bull seminal fluid has an immunosuppressive action was first reported by Sou~.ek et al. (1983), who found that a
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preparation (termed ZS RNase) obtained by acidification of seminal plasma, followed by ammonium sulfate precipitation, inhibited the incorporation of labeled thymidine into stimulated human lymphocytes. Furthermore, the RNase preparation could also (1) depress the potency of mouse spleen cells in a regional graft-versus-host assay and (2) prolong the survival of skin grafts in allogeneic mice, but (3) did not affect the colony-forming activity of mouse bone marrow cells. Later Sou~.ek et al. (1986) showed that a homogeneous preparation of seminal RNase was as immunosuppressive as ZS RNase, and that the enzyme did not inhibit either the killer or the natural killer activities of normal peripheral lymphocytes, whereas it could enhance the colony formation activity in semisolid agar of bone marrow cells. The immunosuppressive activity of BS-RNase was studied also by Tamburrini et al. (1990), who used as an assay the inhibitory effect exerted by BS-RNase on the proliferation of activated lymphocytes. They also found that although the enzyme had no effect on the secretion of interleukin-2 by the activated T cells, it decreased drastically the expression of the c~ chain of the interleukin-2 receptor on the activated T cell membrane. Furthermore, it was found that both the dimeric structure and the integrity of the enzyme catalytic action are essential prerequisites for BS-RNase immunosuppressive activity. The latter finding was recently confirmed by Kim et al. (1995b), who showed the loss of this biological action in a catalytically inactive BS-RNase mutant in which the essential His-119 was replaced by an Asp residue. 3. E m b r y o t o x i c
Activity
One of the first biological effects noticed in the aspermatogenic substance that turned out to be identified with BS-RNase was a pronounced embryotoxic effect (Matou~ek et al., 1973a). When BS-RNase was injected subcutaneously into pregnant female mice, a drastic reduction in the number of embryos and in that of the offspring was observed. The antiembryonic action of BS-RNase was also examined in vitro on twoblastomere embryos, which were arrested in their development at the four-blastomere or the morula stage (Matou~ek et al., 1973a). The antiembryonic effect of BS-RNase has been confirmed using pregnant female rabbits, pigs, and guinea pigs (Matou~ek et al., 1973b). The embryotoxic effect of BS-RNase also depends on the integrity of its catalytic action (Kim et al., 1995b), and on the MxM conformation of the protein (Kim et al., 1995a).
12 Seminal RNase: Importance of Diversity
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4. A n t i v i r a l Action
An antiviral action may not be surprising in a ribonuclease. RNases might degrade the genome of R N A viruses, or complex with the D N A of D N A viruses. Tested on the HIV virus (Youle et al., 1994), the enzyme was found to have no effect on isolated virus particles, but inhibited virus multiplication and the formation of syncytia in H9 virus-infected cells. Interestingly, onconase, a homologous antitumor RNase from frog eggs (see Chapter 15), displays the same antiviral activity, and to a greater extent, whereas other homologous RNases, such as bovine pancreatic RNase A and EDN, a human eosinophil-derived RNase with a selective neurotoxic activity (see Chapter 13), have no antiviral action. On the one hand this suggests that the antiviral action of BS-RNase and onconase is selective; on the other, it hints at a possible correlation between the antitumor property of BS-RNase and onconase and their antiviral action. The H9 cells used in the investigation are in fact tumor cells, although resistant to the cytotoxic action of the antitumor RNases at the concentrations used for the antiviral tests. Possibly, the virus may activate the cytotoxic potential of the RNases by facilitating their intracellular transport to the cytosol, where the antitumor RNases appear to exert their cytotoxicity through degradation of cytosolic R N A (see Chapter 15). Similar results have been obtained by infecting MT-4 or C8166 cells with human immunodeficiency virus type 1 (HIV-1) or HIV-2 viral strains and evaluating the antiviral action of BS-RNase from the extent of inhibition of viral multiplication and of syncitia formation in infected cell cultures, and from the prevention exerted by the enzyme of the virus-induced cytopathogenicity (P. La Colla and G. D'Alessio, unpublished results). 5. A n t i t u m o r Action
The antitumor action of BS-RNase is reviewed in Chapter 15 of this volume.
REFERENCES
Adinolfi, B. S., Cafaro, V., D'Alessio, G., and Di Donato, A. (1995). Full antitumor action of recombinant seminal ribonuclease depends on the removal of its N-terminal methionine. Biochem. Biophys. Res. Commun. 213, 525-532.
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13 Eosinophil-Associated Ribonucleases MELISSA R. S N Y D E R *
AND GERALD
J. GLEICHt
* Departments of Biochemistry and Molecular Medicine t Departments of Immunology and Medicine Mayo Clinic and Foundation Rochester, Minnesota 55905
I. Introduction A. Historical Perspective B. The Eosinophil C. Gordon Phenomenon D. Eosinophilia II. The Ribonuclease Superfamily A. Purification of Eosinophil-Derived Neurotoxin B. Sequence Information III. Eosinophil-Derived Neurotoxin A. General Sequence Characteristics B. Glycosylation C. Enzymatic Activity D. Helminthotoxicity E. Neurotoxicity F. Inhibition of Protein Synthesis and Cytotoxicity IV. Eosinophil Cationic Protein A. General Sequence Characteristics B. Glycosylation C. Enzymatic Activity D. Helminthotoxicity E. Neurotoxicity F. Inhibition of Protein Synthesis and Cytotoxicity V. Conclusion References
425 RIBONUCLEASES: STRUCTURES AND FUNCTIONS
Copyright 9 1997 by Academic Press, Inc. All rights of reproduction in any form reserved.
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Melissa R. Snyder and Gerald J. Gieich
I. I N T R O D U C T I O N
A. Historical Perspective In 1879, Paul Ehrlich observed a leukocyte that possessed a strong affinity for acidic dyes (Gleich and Adolphson, 1986). Due to the avidity of this cell for eosin, he named this leukocyte the eosinophil. Shortly after this discovery, increased numbers of eosinophils were associated with bronchial asthma and parasitic infections (Gleich and Adolphson, 1986). Following anaphylaxis, eosinophils infiltrate the lungs of guinea pigs, suggesting a link between eosinophils and asthma (Gleich et al., 1995). Originally, it was hypothesized that the eosinophil modulated hypersensitivity reactions by neutralizing molecules associated with mast cells, such as histamine (Gleich et al., 1995). However, it is now thought that the eosinophil functions as a proinflammatory cell and is responsible, in part, for tissue damage that occurs during allergic responses. The eosinophil granule proteins possess cytotoxic and cytostimulatory activities and have the potential to cause bronchial tissue damage in asthma. The eosinophil functions in a similar manner during certain infections, although the cytotoxic activity is directed toward the invading organism.
B. The Eosinophil The eosinophil is characterized by a bilobed nucleus and large cytoplasmic granules (Gleich and Adolphson, 1986; Gleich et al., 1995). These granules consist of an electron-dense, crystalline core and electron-lucent matrix. In response to a parasitic infection or hypersensitivity reaction, the eosinophil releases the contents of these granules into the extracellular environment (Gleich and Adolphson, 1986; Gleich et al., 1995). The proteins found within the granule are cytotoxic and likely mediate many of the biological functions of the eosinophil. This review will examine two of these proteins, eosinophil-derived neurotoxin (EDN) and eosinophil cationic protein (ECP), with respect to their physiology and biochemistry.
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C. Gordon Phenomenon
In 1933, M. H. Gordon published a report in which he claimed to have discovered the agent responsible for the development of Hodgkin's disease. He observed that rabbits, after an intracerebral injection of lymph node homogenates from persons with this disease, developed muscle stiffness and ataxia. This observation was confirmed by others (Durack et aL, 1979), and it was found that the neurotoxic response produced in the rabbits was caused by eosinophils present in the lymph node homogenates. This unusual neurotoxicity of the eosinophil is known as the Gordon phenomenon. The Gordon phenomenon can be induced in experimental animals, such as rabbits, by an intracerebral or intrathecal injection of purified eosinophils (Durack et al., 1979). Several days after the injection, the animal develops muscle stiffness and mild ataxia, followed by incoordination and more severe ataxia. Eventually, these animals experience severe muscle wasting and, in some cases, paralysis. Histologically, these symptoms are associated with loss of Purkinje cells in the cerebellum and a vacuolation of the white matter in the cerebellum, spinal cord, and brain stem. The white matter vacuolation, which occurs several days after the loss of Purkinje cells, seems to be due to disruption of the myelin sheath around the axon (Durack et al., 1979).
D. Eosinophilia
Although no direct counterpart to the Gordon phenomenon in humans is known, there are diseases associated with peripheral blood eosinophilia that affect the nervous system, including the eosinophilmyalgia syndrome (EMS) and the hypereosinophilic syndrome (HES). EMS is characterized by a moderate to marked peripheral blood eosinophilia in the absence of infection or allergy (Hertzman et al., 1990; Belongia et al., 1993). EMS affects many systems of the body, although the most common symptoms are severe myalgia and muscle weakness (Butterfield et al., 1995; Belongia et al., 1993; Hertzman et al., 1990). The 1989 EMS epidemic was associated with over-the-
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counter consumption of L-tryptophan, manufactured by Showa Denko KK (Belongia et al., 1993), containing contaminants including 1,1'ethylidenebistryptophan (EBT) (Mayeno et al., 1990). A recent report claims that EBT can induce interleukin (IL)-5 production by T cells (Yamaoka et al., 1991), and IL-5, in turn, is able to induce eosinophil growth, differentiation, and release of granular proteins (Kita et al., 1992). Another report claims that EBT can act directly on the eosinophil, resulting in up-regulation of the IL-5 receptor and degranulation (Yamaoka et al., 1994). Although these claims pose an attractive pathophysiology for EMS, our laboratory has not been able to substantiate these findings (Kita et al., 1995). Patients with EMS have elevated blood levels of eosinophil proteins (Martin et al., 1990; Varga et al., 1992), indicating that degranulation has occurred in vivo. The symptoms from the EMS outbreak caused by contaminated L-tryptophan were similar to those of the toxic oil syndrome (Butterfield et al., 1995; Silver, 1993). This epidemic occurred in Spain beginning in 1981, and was caused by ingestion of adulterated rapeseed oil. Further investigation into the L-tryptophan epidemic led to the discovery of a second contaminant, 3-(phenylamino)alanine, which is similar to the contaminant 3-phenylamino-l,2-propanediol isolated from the adulterated rapeseed oil (Mayeno et al., 1992). It has been suggested, based on structural similarities, that these contaminants may stimulate similar disease mechanisms, resulting in marked eosinophilia and neurological manifestations. Another condition, HES, is also characterized by an elevated peripheral blood eosinophil count that persists longer than 6 months in the absence of any known cause (Butterfield et al., 1995; Spry, 1993). Patients with HES show widespread organ involvement, particularly within the heart, skin, and nervous system. Neurological problems are very common and approximately 65% of HES patients display nervous system dysfunction (Chusid et al., 1975; Moore et al., 1985) ranging from muscle ataxia to behavioral changes, confusion, and memory loss (Moore et al., 1985; Prick et al., 1988). It is clear that the eosinophil is responsible for the Gordon phenomenon in experimental animals, but it is difficult to demonstrate conclusively a similar relationship between the eosinophil and the neurological abnormalities in EMS and HES. However, the similarities in the clinical manifestations of these two syndromes, unrelated except for eosinophilia, suggest that the eosinophil plays a role in human neurological disease.
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II. THE RIBONUCLEASE SUPERFAMILY A. Purification of Eosinophil-Derived Neurotoxin and Eosinophil Cationic Protein
Fractionation of eosinophil organelles led to the discovery that the Gordon phenomenon is induced by the cytoplasmic granule (Durack et al., 1979). The eosinophil granule contains several proteins, including major basic protein (MBP), eosinophil-derived neurotoxin (EDN), eosinophil cationic protein (ECP), and eosinophil peroxidase (EPO) (Ackerman et al., 1983). Through immunoelectron microscopy, these proteins have been localized to the granule. EDN, ECP, and EPO are located in the granule matrix, whereas MBP is found in the crystalline core (Peters et aL, 1986). MBP is toxic to many mammalian cells and other organisms, including parasites and bacteria (Gleich and Adolphson, 1986; Gleich et al., 1995). Because of its potent cytotoxicity, it was surprising that purified MBP is not able to induce the Gordon phenomenon (Durack et al., 1981). On the other hand, EDN and ECP, the two matrix proteins, produce the Gordon phenomenon (Durack et al., 1981; Fredens et al., 1982). In addition to being neurotoxic, ECP is a potent helminthotoxin and bactericide with activities similar to MBP (Yazdanbakhsh et al., 1987; Lehrer et al., 1989; Ackerman et al., 1985; Hamann et al., 1987, 1990a; Molina et al., 1988). However, EDN displays no bacterial and only weak parasitic toxicity when compared to MBP and ECP (Ackerman et al., 1985; Hamann et al., 1987, 1990a; Molina et al., 1988); its most obvious activity is potent neurotoxicity.
B. Sequence Information
The first sequence information for EDN and ECP was obtained from purified proteins (Gleich et al., 1986). Although the sequence was incomplete, a high degree of homology between these two proteins and human pancreatic ribonuclease (HPRNase) was observed (Beintema et al., 1984). When the genes for EDN (Hamann et al., 1989; Rosenberg et al., 1989b) and ECP (Barker et al., 1989; Rosenberg et al., 1989a) were cloned, it was observed that, in addition to displaying a 65% sequence identity, they were each approximately 30% identical to HPRNase (Bein-
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tema et al., 1984). Subsequently, many other proteins that display sequence homology to HPRNase have been identified, and these proteins comprise the ribonuclease superfamily. Angiogenin, a protein originally purified from the media of a human colon adenocarcinoma cell line and able to induce vascularization of tumors, is 30% identical to E D N (Strydom et al., 1985; Kurachi et al., 1985). EDN is also about 30% identical both to bovine seminal ribonuclease (Palmieri et al., 1985), originally purified from bull semen, and to onconase (Ardelt et al., 1991), isolated from R a n a pipiens oocytes. In addition to being homologous to a variety of proteins, EDN is completely identical in sequence to human urinary nonsecretory ribonuclease (HUNRNase) (Beintema et al., 1988), and is identical through the first 30 residues of human liver ribonuclease (HLRNase) (Sorrentino et aL, 1988) and human spleen ribonuclease (Yasuda et aL, 1990). As suggested by the sequence homology, all of these proteins, including EDN and ECP, are enzymatically active as ribonucleases. The genes for EDN ( R N S 2 ) and ECP ( R N S 3 ) , located on chromosome 14 (Mastrianni et al., 1992; Hamann et al., 1990b), are each composed of two exons separated by a single intron in the 5' untranslated region (Hamann et al., 1990a, 1991). BPRNase also contains a single intron in this region (Carsana et al., 1988), and the angiogenin gene is composed of an intronless coding sequence (Kurachi et al., 1985). Because the angiogenin gene has not yet been completely sequenced, it is not known whether its 5' untranslated region contains an intron. In any case, the intronless coding sequence may be a characteristic of the ribonuclease superfamily. The high level of nucleotide identity in the coding regions between EDN and ECP is also observed in the 5' untranslated regions, the single intron, and 75 base pairs into the 3' untranslated regions (Hamann et al., 1990b, 1991). The EDN and ECP genes may have arisen from a gene duplication event that occurred approximately 25 to 40 million years ago (Hamann et aL, 1990b). In addition to the eosinophil, other leukocytes contain varying amounts of EDN and ECP. Based on immunological assays, EDN is found in basophils and neutrophils, although at levels 15- to 50-fold lower than in the eosinophil (Abu-Ghazaleh et al., 1992). Basophils and neutrophils also contain ECP at levels 70- to 100-fold below that found in the eosinophil (Abu-Ghazaleh et al., 1992). Analyses of mRNA from various cells and cell lines also indicate the presence of EDN and ECP. EDN mRNA was found in the uninduced promyelocytic leukemia line, HL-60 (Rosenberg et al., 1989a,b); the basal level of mRNA was up-regulated when
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cells were induced to eosinophilic or neutrophilic differentiation, but it was down-regulated after differentiation toward monocytes (Rosenberg et al., 1989a,b). EDN mRNA was also detected in human blood eosinophils and neutrophils, but not monocytes (Rosenberg et al., 1989b). In contrast to the mRNA findings with EDN, ECP mRNA could be detected only in mature peripheral blood eosinophils and in HL-60 cells induced toward eosinophilic differentiation (Rosenberg et al., 1989a). No mRNA was found in uninduced HL-60 cells or in those induced toward monocytic or neutrophilic differentiation (Rosenberg et al., 1989a). Despite the similarities of the EDN and ECP genes, regulatory mechanisms controlling their differential transcription must exist.
III. EOSINOPHIL-DERIVED NEUROTOXIN
A. General Sequence Characteristics EDN is synthesized as a preprotein with a 27-amino-acid signal peptide. Cleavage of this peptide results in a mature protein of 134 residues (Hamann et al., 1989; Rosenberg et al., 1989b). EDN is a cationic protein with a calculated isoelectric point of 8.9. There are many similarities between the primary sequences of EDN and HPRNase. In HPRNase, there are eight cysteine residues forming four disulfide bonds (Blackburn and Moore, 1982). All of these residues are conserved in EDN. EDN possesses the three catalytic residues, histidine-15, lysine-38, and histidine-129, required for ribonuclease activity, and many of the residues known to be important for substrate binding in HPRNase (Blackburn and Moore, 1982; Beintema, 1989). One particular difference in the substrate-binding residues occurs at position 130 in EDN, where the phenylalanine in HPRNase has been replaced with leucine. Alterations within the substrate-binding pocket may explain the differences in enzymatic and biological activities among the members of the ribonuclease superfamily.
B. Glycosylation Many members of the ribonuclease superfamily possess potential sites for N-linked glycosylation, whose consensus site is Asn-X-Thr/Ser. Gly-
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cosylation typically introduces heterogeneity into a population of protein molecules, and because glycosylation is tissue specific, it may be one way that the various ribonucleases achieve structural diversity. This is evident in the comparison of HPRNase and E D N sequences. HPRNase has three N-linked glycosylation consensus sites located at positions 34, 76, and 88 (Beintema et al., 1984). The mature ribonuclease is actually a mixture of differentially glycosylated proteins ranging from 14 to 30 kDa (Ribo et al., 1994). In almost all HPRNase molecules, N-34 is glycosylated, whereas only 50% of the molecules are glycosylated at N76. Glycosylation of the final site, N-88, is rarely observed (Ribo et al., 1994). EDN contains five potential sites for N-linked glycosylation (Hamann et al., 1989; Rosenberg et al., 1989b), although none is conserved from HPRNase. Although it is not known which of these sites is glycosylated, it is clear that native EDN possesses a certain amount of carbohydrate. According to the amino acid sequence, the molecular mass of mature EDN is 15.5 kDa. However, as analyzed by SDS gel electrophoresis, EDN exists as a heterogeneous population of molecules with molecular masses between 18 and 21 kDa (Gleich et al., 1986). Treatment of EDN with endoglycosidase F results in a decrease in the molecular mass of the protein, whereas a similar treatment with endoglycosidase H does not (Gleich et al., 1986). Because endoglycosidase F cleaves either complex or high-mannose oligosaccharides, and endoglycosidase H cleaves only the high-mannose type, it seems likely that EDN contains complex oligosaccharides. In addition to the conventional Nlinked carbohydrates, it has been reported that tryptophan-7 of HUNRNase is modified with an aldohexopyranosyl residue at the C-2 position of the indole ring (Hofsteenge et al., 1994). Because EDN and HUNRNase have identical primary sequences, it is tempting to hypothesize that EDN is also modified in the same manner. This is particularly attractive in light of the observation that position seven of EDN results in a blank during amino acid sequencing (Gleich et al., 1986). However, this type of modification may be tissue specific and found only in HUNRNase. Further characterization of native EDN is required.
C. Enzymatic Activity Despite the sequence homology, there are differences between the activity and substrate specificity of HPRNase and EDN. HPRNase is
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active on most R N A substrates, including dinucleotide monophosphates and double-stranded R N A (dsRNA). The only known substrate specificity of HPRNase is a preference for pyrimidine residues, which is also shared by E D N (Weickmann et al., 1981; Sorrentino et al., 1992). With natural substrates, such as yeast R N A or wheat germ RNA, E D N is approximately threefold lower in activity than HPRNase (Sorrentino et al., 1992; Slifman et al., 1986). However, with synthetic polymers of single-stranded RNA, E D N can be anywhere up to 30-fold lower in activity, depending on the sequence of the R N A (Sorrentino et al., 1992). E D N also displays no detectable activity toward small substrates, including cyclic 2',3'-phosphates and dinucleotide phosphates, although HPRNase is quite active on these substrates (Sorrentino et al., 1992). As far as can be assayed, HLRNase and E D N are very similar in their enzymatic activities (Sorrentino et al., 1992), suggesting that any tissuespecific alterations, such as glycosylation, are not affecting enzymatic activities. As may be expected from the identity of the catalytic residues, most ribonucleases in the superfamily lose enzymatic activity after treatment with any reagent that modifies histidine residues, such as iodoacetic acid. Treatment of BPRNase with this reagent results in carboxymethylation of catalytic residues histidine-12 or histidine-119 (Plapp, 1973; Crestfield et al., 1963), but never both residues in the same molecule. By analogy, it is hypothesized that treatment of EDN with iodacetic acid probably results in carboxymethylation at histidine-15 or histidine-129. The enzymatic activity of EDN, HLRNase, and HPRNase can also be inhibited by the placental ribonuclease inhibitor (Sorrentino et al., 1992). The inhibitor and ribonuclease form a one-to-one complex, although little is known about the details of the structure.
D. Helminthotoxicity
The only two members of the ribonuclease superfamily known to be toxic to parasites are the eosinophil proteins. EDN typically displays an LDs0 between 0.1 and 0.2 m M and generally is less potent than MBP, although this is dependent on both the type of parasite and the time of incubation. For example, EDN is only 10-fold lower than MBP in its toxicity toward Trichinella spiralis newborn larvae (Hamann et al., 1987), although it is essentially inactive toward microfilariae of Brugia p a h a n g i
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and Brugia malayi (Hamann et al., 1990a). The toxicities of these two proteins for T r y p a n o s o m a cruzi tryptomastigotes are almost identical (Molina et al., 1988), although the mechanisms seem to differ. The toxicity of MBP is inhibited by heparin whereas the activity of E D N is not (Molina et al., 1988). However, addition of yeast R N A or placental ribonuclease inhibitor abolishes E D N toxicity toward the parasite (Molina et aL, 1988). Although this observation suggests that, in the case of EDN, the helminothotoxicity is related to its ribonuclease activity, it is interesting that BPRNase is not toxic at similar concentrations (Molina et al., 1988).
E. Neurotoxicity As mentioned before, E D N is a potent neurotoxin. At a dose of 7 ~g, E D N is capable of inducing the Gordon phenomenon in rabbits within 3 to 5 days after an intrathecal injection (Gleich et al., 1986). Even a 20-fold lower dose of the protein is still neurotoxic, although the time before symptom onset is longer (Gleich et al., 1986). Several other members of the ribonuclease superfamily, including HLRNase (Sorrentino et aL, 1992) and onconase (Newton et al., 1994b), are also neurotoxic. In light of the complete primary sequence identity, it is not surprising that HLRNase displays the same neurotoxic potency as EDN. On the other hand, onconase, which is only 30% identical to EDN, can also induce the Gordon phenomenon with the same efficacy as EDN (Newton et al., 1994a). HPRNase is also neurotoxic, although it requires threefold to fourfold higher doses than E D N (Sorrentino et al., 1992). Many experiments, primarily involving chemical modification, have been performed in an attempt to demonstrate a link between ribonuclease activity and neurotoxicity. When EDN or HLRNase is treated with iodoacetic acid (in the absence of a denaturant), the resulting carboxymethylated proteins are not enzymatically active, but they are still recognized by antibodies made to the native protein (Sorrentino et al., 1992). In contrast, carboxymethylation of denatured and reduced E D N results in an enzymatically inactive protein that is not recognized by the same antibodies (Sorrentino et al., 1992). Thus, the carboxymethylated E D N retains some native structure. Carboxymethylated E D N and HLRNase are not able to induce the Gordon phenomenon (Slifman et a/.,1986), despite the retention of nativelike structure. This suggests that the enzymatic activity
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is crucial for neurotoxicity. However, it is clear that ribonuclease activity is not sufficient for this particular biological function, because BPRNase has no detectable neurotoxic activity at doses 1000-fold higher than E D N (Sorrentino et al., 1992).
F. Inhibition of Protein Synthesis and Cytotoxicity Ribonucleases are capable of inhibiting protein synthesis by degradation of RNA, leading to cell death. When injected into X e n o p u s oocytes, E D N inhibits total protein synthesis with an ICs0 of 10 -5 mg/ml (Saxena et al., 1992). The loss of protein synthesis corresponds to a degradation of total cellular RNA. In this system, inhibition of protein synthesis requires microinjection of E D N because there is no mechanism for its transport into the cytoplasm. This problem can be overcome by taking advantage of receptor-mediated endocytosis. When injected into X e n o p u s oocytes, BPRNase inhibits protein synthesis with an efficacy similar to E D N (Saxena et aL, 1992; Rybak et al., 1991). However, with cultured K562 human erythroleukemia cells, the presence of BPRNase in the media has no effect on the protein synthesis or growth of the cells (Rybak et al., 1991). To determine if cytoplasm accessibility is the problem, BPRNase was linked through disulfide bonds to transferrin. This hybrid protein is capable of inhibiting protein synthesis in K562 cells expressing the transferrin receptor with an ICs0 of 10 -v M (Rybak et al., 1991). Transferrin binds to its receptor on the extracellular surface of the membrane, resulting in endocytosis of the fusion protein. The BPRNase, which is still enzymatically active, can then degrade the R N A in the cytoplasm of the cell, resulting in cell death. It is also possible to construct fusion proteins through the use of recombinant D N A technology. Recombinant E D N was expressed as a fusion protein with the single-chain antibody to the transferrin receptor (Newton et al., 1994a). Although this fusion protein was only 10% as enzymatically active as native EDN, it was toxic to tumor cells expressing the transferrin receptor with the ICs0 of 0.5 nM (Newton et al., 1994a). A recombinant form of EDN lacking the antibody was not toxic to these cells, nor was the fusion protein toxic to cells that did not express the receptor (Newton et al., 1994a). If the ribonuclease activity of EDN is responsible for its helminthotoxicity or neurotoxicity, specific mechanisms for its transport into the cell must exist. As a purified protein,
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without any transport facilitation, onconase inhibits protein synthesis and cell growth of 9L glioma cells with an ICs0 of l0 -7 M (Wu et aL, 1993). Onconase exhibits saturable binding to these cells, and its toxic effect is lost after treatment with iodoacetic acid (Wu et al., 1993). The toxicity of onconase toward these cells can be increased 100-fold by retinoic acid (Wu et al., 1995). Retinoic acid disrupts the Golgi apparatus, facilitating transport of the protein to the cytosol. The use of retinoic acid makes the 9L glioma cells susceptible to otherwise nontoxic enzymes, such as HPRNase (Wu et al., 1995). These findings suggest the hypothesis that onconase binds to a specific receptor on the surface of these cells, and is internalized through endocytosis. Onconase then travels through the Golgi apparatus before entering the cytosol. In other words, onconase is toxic only when it can degrade intracellular RNA. The 9L glioma cells must have a receptor that allows for specific interaction with onconase because HPRNase, even in the presence of retinoic acid, is 10,000-fold less toxic than onconase (Wu et al., 1995). A similar mechanism may be utilized for neurotoxicity, in which there is internalization of specific ribonuclease by the Purkinje cell, leading to cell death.
IV. E O S I N O P H I L C A T I O N I C P R O T E I N
A. General Sequence Characteristics In its primary amino acid sequence, ECP is quite similar to EDN. Mature ECP is composed of 133 amino acids, although it is synthesized as a preprotein with a signal peptide of 27 residues (Barker et aL, 1989; Rosenberg et al., 1989a). The level of identity between EDN and ECP is 70%, although their signal peptides share approximately 90% identity. ECP is even more cationic than EDN, with an esimated isoelectric point of 10.8 (Barker et al., 1989; Rosenberg et al., 1989a). In fact, almost 50% of the amino acid differences between the two proteins result from the change of a neutral or negatively charged residue in EDN to an arginine or lysine in ECP. ECP also possesses the eight conserved cysteine residues that probably form four disulfide bonds. The three catalytic residues required for ribonuclease activity are also present in ECP, as well as the same substrate-binding residues found in EDN. Although the primary sequence similarities between EDN and ECP are quite extensive, there
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must be differences in the secondary and tertiary structures of the two proteins to account for the differences in biological functions.
B. Glycosylation Five consensus sites for N-linked glycosylation are found in ECP, although only two are conserved in EDN. Mature ECP has a molecular mass estimated from the amino acid sequence of 15.5 kDa; however, several protein species between 16 and 21 kDa can be observed on SDS gels (Gleich et al., 1986; Peterson et aL, 1988). These various species disappear after ECP is treated with endoglycosidase F, but not with endoglycosidase H (Gleich et al., 1986). Again, this behavior is similar to EDN and suggests that ECP is glycosylated with complex or hybrid oligosaccharides, although it is not known which sites within ECP are modified. It is possible that the glycosylated sites in EDN and ECP are not conserved between the two proteins, thus leading to a greater level of structural diversity. ECP lacks tryptophan-7, which is the site of the potential aldohexopyranosyl modification in HUNRNase (Hofsteenge et al., 1994). If this modification is present in EDN, it may also have an impact on the three-dimensional structure of the protein, which may in turn affect its biological activities.
C. Enzymatic Activity In general, the RNase activity of ECP resembles that of EDN. ECP, like EDN, prefers pyrimidine residues, with the highest activity being displayed toward poly(U) (Sorrentino and Glitz, 1991). With the poly(U) substrate, EDN is approximately 10-fold more active than ECP (Slifman et al., 1986; Sorrentino and Glitz, 1991). ECP is most active on natural substrates, although it is still 100-fold less active than EDN on tRNA and yeast RNA (Gullberg et al., 1986). ECP, like EDN, is not active toward very small substrates such as cyclic 2',3'-phosphates and dinucleoside phosphates, or dsRNA (Gullberg et al., 1986). The reasons for the differences in the specific activities of EDN and ECP are not yet understood.
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D. Helminthotoxicity ECP and E D N are the only members of the ribonuclease family known to be helminthotoxic. ECP typically displays an LDs0 between 0.1 and 6 mM, which is approximately 2-fold to 30-fold more potent than E D N (Ackerman et al., 1985; Hamann et aL, 1987, 1990a; Molina et aL, 1988). In fact, this level of parasitic activity is comparable to that of major basic protein (MBP). Although E D N and ECP are very similar in primary sequence and enzymatic activity, their individual mechanisms of parasitic toxicity may differ. The addition of total yeast R N A or placental ribonuclease inhibitor prevented E D N from killing T r y p a n o s o m a cruzi tryptomastigotes, but had no effect on the toxicity of ECP (Molina et aL, 1988). On the other hand, the toxic effects of ECP and MBP were inhibited by heparin, but that of E D N was not (Molina et aL, 1988). If ribonuclease activity is responsible for the helminthotoxicity, it appears contradictory that EDN, which has the higher specific activity, is less toxic to parasites. This would suggest that either the transport of ECP into the parasite is more efficient than for EDN, or that ECP simply uses another mechanism of action that is more efficient than degradation of cellular RNA. In addition to being helminthotoxic, ECP is also toxic to bacteria (Lehrer et aL, 1989). The mechanism of this toxicity was investigated using recombinant protein methods (Rosenberg, 1995). A mutant ECP protein was constructed in which two catalytic residues were altered, resulting in a loss of ribonuclease activity. Despite the lack of enzymatic activity, the mutant ECP was as toxic to bacteria as was the wild-type protein (Rosenberg, 1995). It has been reported that ECP is able to form membrane channels (Young et al., 1986), although this has not been demonstrated in bacteria. However, it is clear that the antibacterial property of ECP is independent of its ribonuclease activity, although the mechanism is not known.
E. Neurotoxicity ECP is capable of inducing the Gordon phenomenon in rabbits, although it is approximately 10-fold less active than E D N (Gleich et al., 1986). Treatment of ECP with iodoacetic acid, which is thought to car-
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boxymethylate the active site histidine residues and to abolish ribonuclease activity, results in a loss of neurotoxic activity (Gleich et al., 1986). This result suggests that both ECP and E D N cause the Gordon phenomenon through their ribonuclease activity. Thus, ECP may mediate its toxicity through different mechanisms, depending on the target cell. ECP has also been implicated in human neurological diseases. Levels of ECP in the cerebrospinal fluid of patients with various central nervous system diseases are significantly higher than those in normal individuals without neurological diseases (Hallgren et al., 1983). Given that ECP is capable of inducing the Gordon phenomenon in animals, it is reasonable to suspect that ECP may be involved in the pathogenesis of human neurological disease.
F. Inhibition of Protein Synthesis and Cytotoxicity The eosinophil appears to be the primary leukocyte responsible for tissue damage in bronchial asthma, which occurs when the granule proteins, including ECP, are released into the extracellular space. ECP is thought to be a mediator of tissue damage based on protein levels in serum and bronchoalveolar lavage fluid. On exposure to allergen, serum and bronchoalveolar lavage fluid ECP levels increase, correlating with the severity of the asthmatic response. After corticosteroid treatment, ECP levels begin to decrease (Ahlstedt, 1995; Venge, 1993, 1994; Woolley et al., 1995). More direct evidence for the involvement of ECP in bronchial tissue damage comes from the observation that ECP is toxic to guinea pig tracheal epithelium in vitro (Motojima et al., 1989). Somewhat surprisingly, EDN, even at concentrations twofold higher than ECP, was not toxic to the epithelial cells (Motojima et aL, 1989).
V. C O N C L U S I O N
EDN and ECP, two members of the ribonuclease superfamily, are extracellular ribonucleases having cytotoxic capabilities. Although this is important for immunity to parasites, it may have deleterious effects, similar to the Gordon phenomenon, in diseases associated with increased
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numbers of eosinophils. Despite the observation that E D N can cause cell death through R N A degradation and inhibition of protein synthesis, additional experiments are needed to conclude that neurotoxicity and helminthotoxicity are caused by ribonuclease activity. In addition, evidence exists indicating that a ribonuclease-inactive mutant of ECP is bactericidal. If ribonuclease activity is responsible for E D N toxicity, there must be specific mechanisms by which E D N can enter the cell. Through this type of specific interaction, many proteins with similar enzymatic activity could have very diverse biological functions.
REFERENCES
Abu-Ghazaleh, R. I., Dunnette, S. L., Loegering, D. A., Checkel, J. L., Kita, H., Thomas, L. L., and Gleich, G. J. (1992). Eosinophil granule proteins in peripheral blood granulocytes. J. Leukocyte Biol. 52, 611-618. Ackerman, S. J., Loegering, D. A., Venge, P., Olsson, I., Harley, J. B., Fauci, A. S., and Gleich, G. J. (1983). Distinctive cationic proteins of the human eosinophil granule: Major basic protein, eosinophil cationic protein, and eosinophil-derived neurotoxin. J. Immunol. 131, 2977-2982. Ackerman, S. J., Gleich, G. J., Loegering, D. A., Richardson, B. A., and Butterworth, A. E. (1985). Comparative toxicity of purified human eosinophil granule cationic proteins for schistosomula of Schistosoma mansoni. Am. J. Trop. Med. Hyg. 34, 735-745. Ahlstedt, S. (1995). Clinical application of eosinophilic cation protein in asthma. Allergy Proc. 16, 59-62. Ardelt, W., Mikulski, S. M., and Shogen, K. (1991). Amino acid sequence of an antitumor protein from Rana pipiens oocytes and early embryos: Homology to pancreatic ribonucleases. J. Biol. Chem. 266, 245-251. Barker, R. L., Loegering, D. A., Ten, R. M., Hamann, K. J., Pease, L. R., and Gleich, G. J. (1989). Eosinophil cationic protein cDNA: Comparison with other toxic cationic proteins and ribonucleases. J. Immunol. 143, 952-955. Beintema, J. J. (1989). Presence of a basic amino acid residue at either position 66 or 122 is a condition for enzymatic activity in the ribonuclease superfamily. FEBS Lett. 254, 1-4. Beintema, J. J., Wietzes, P., Weickmann, J. L., and Glitz, D. G. (1984). The amino acid sequence of human pancreatic ribonuclease. A n a l Biochem. 136, 48-64. Beintema, J. J., Hofsteenge, J., lwama, M., Morita, T., Ohgi, K., Irie, M., Sugiyama, R. H., Schieven, G. L., Dekker, C. A., and Glitz, D. G. (1988). Amino acid sequence of the nonsecretory ribonuclease of human urine. Biochemistry 27, 4530-4538. Belongia, E. A., Mayeno, A. N., Gleich, G. J., and Kita, H. (1993). Eosinophilia-myalgia syndrome. In "Eosinophils: Biological and Clinical Aspects" (S. Makino and T. Fukuda, eds.), Chapter 23, pp. 421-438. CRC Press, Boca Raton, Florida.
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Blackburn, P., and Moore, S. (1982). In "The Enzymes" (P. D. Boyer, ed), 3rd Ed., Vol. 15, pp. 317-433. Academic Press, New York. Butterfield, J. H., Leiferman, K. M., and Gleich, G. J. (1995). Eosinophil-associated diseases. In "Samter's Immunologic Diseases" (M. M. Frank, K. F. Austen, H. N. Claman, and E. R. Unanue, eds.), 5th Ed., Vol. 1, Chapter 29, pp. 501-527. Little, Brown, Boston, Massachusetts. Carsana, A., Confalone, E., Palmieri, M., Libonati, M., and Furia, A. (1988). Structure of the bovine pancreatic ribonuclease gene: The unique intervening sequence in the 5' untranslated region contains a promoter-like element. Nucleic Acids Res. 16, 54915502. Chusid, M. J., Dale, D. C., West, B. C., and Wolff, S. M. (1975). The hypereosinophilic syndrome: Analysis of fourteen cases with review of the literature. Medicine 54, 1-27. Crestfield, A. M., Stein, W. H., and Moore, S. (1963). Alkylation and identification of the histidine residues at the active site of ribonuclease. J. Biol. Chem. 238, 2413-2420. Durack, D. T., Sumi, S. M., and Klebanoff, S. J. (1979). Neurotoxicity of human eosinophils. Proc. Natl. Acad. Sci. U.S.A. 76, 1443-1447. Durack, D. T., Ackerman, S. J., Loegering, D. A., and Gleich, G. J. (1981). Purification of human eosinophil-derived neurotoxin. Proc. Natl. Acad. Sci. U.S.A. 78, 5165-5169. Fredens, K., Dahl, R., and Venge, P. (1982). The Gordon phenomenon induced by the eosinophil cationic protein and eosinophil protein X. J. Allergy Clin. Immunol. 70, 361-366. Gleich, G. J., and Adolphson, C. R. (1986). The eosinophilic leukocyte: Structure and function. Adv. Immunol. 39, 177-253. Gleich, G. J., Loegering, D. A., Bell, M. P., Checkel, J. L., Ackerman, S. J., and McKean, D. J. (1986). Biochemical and functional similarities between human eosinophilderived neurotoxin and eosinophil cationic protein: Homology with ribonuclease. Proc. Natl. Acad. Sci. U.S.A. 83, 3146-3150. Gleich, G.J., Kita, H., and Adolphson, C. R. (1995). Eosinophils. In "Samter's Immunologic Diseases" (M. M. Frank, K. F. Austen, H. N. Claman, and E. R. Unanue, eds.), 5th Ed., Vol. 1, Chapter 14, pp. 205-245. Little, Brown, Boston, Massachusetts. Gordon, M. H. (1933). Remarks on Hodgkin's disease. A pathogenic agent in the glands, and its application in diagnosis. Br. Med. J. 1, 641-644. Gullberg, U., Widegren, B., Arnason, U., Egesten, A., and Olsson, I. (1986). The cytotoxic eosinophil cationic protein (ECP) has ribonuclease activity. Biochem. Biophys. Res. Commun. 139, 1239-1242. Hallgren, R., Terent, A., and Venge, P. (1983). Eosinophil cationic protein (ECP) in the cerebrospinal fluid. J. Neurol. Sci. 58, 57-71. Hamann, K. J., Barker, R. L., Loegering, D. A., and Gleich, G. J. (1987). Comparative toxicity of purified human eosinophil granule proteins for newborn larvae of Trichinella spiralis. J. Parasitol. 73, 523-529. Hamann, K. J., Barker, R. L., Loegering, D. A., Pease, L. R., and Gleich, G. J. (1989). Sequence of human eosinophil-derived neurotoxin cDNA: Identity of deduced amino acid sequence with human nonsecretory ribonucleases. Gene 83, 161-167. Hamann, K. J., Gleich, G. J., Checkel, J. L., Loegering, D. A., McCall, J. W., and Barker, R. L. (1990a). In vitro killing of microfilariae of Brugia pahangi and Brugia malayi by eosinophil granule proteins. J. Immunol. 144, 3166-3173. Hamann, K. J., Ten, R. M., Loegering, D. A., Jenkins, R. B., Heise, M. T., Schad, C. R., Pease, L. R., Gleich, G. J., and Barker, R. L. (1990b). Structure and chromosome
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localization of the human eosinophil-derived neurotoxic and eosinophil cationic protein genes: Evidence for intronless coding sequences in the ribonuclease gene superfamily. Genomics 7, 535-546. Hamann, K. J., Barker, R. L., Ten, R. M., and Gleich, G. J. (1991). The molecular biology of eosinophil granule proteins. Int. Arch. Allergy Appl. Immunol. 94, 202-209. Hertzman, P. A., Blevins, W. L., Mayer, J., Greenfield, B., Ting, M., and Gleich, G. (1990). Association of the eosinophilia-myalgia syndrome with the ingestion of tryptophan. N. Engl. J. Med. 322, 869-873. Hofsteenge, J., Muller, D. R., de Beer, T., Loftier, A., Richter, W. J., and Vliegenthart, J. F. G. (1994). New type of linkage between a carbohydrate and a protein: CGlycosylation of a specific tryptophan residue in human RNase Us. Biochemistry 33, 13524-13530. Kita, H., Weiler, D. A., Abu-Ghazaleh, R., Sanderson, C. J., and Gleich, G. J. (1992). Release of granule proteins from eosinophils cultured with IL-5. J. Immunol. 149, 629-635. Kita, H., Mayeno, A. N., Weyand, C. M., Goronzy, J. J., Weiler, D. A., Lundy, S. K., Abrams, J. S., and Gleich, G. J. (1995). Eosinophil-active cytokine from mononuclear cells cultured with L-tryptophan products: An unexpected consequence of endotoxin contamination. J. Allergy Clin. Immunol. 95, 1261-1267. Kurachi, K., Davie, E. W., Strydom, D. J., Riordan, J. F., and Vallee, B. L. (1985). Sequence of the cDNA and gene for angiogenin, a human angiogenesis factor. Biochemistry 24, 5494-5499. Lehrer, R. I., Szklarek, D., Barton, A., Ganz, T., Hamann, K. J., and Gleich, G. J. (1989). Antibacterial properties of eosinophil major basic protein and eosinophil cationic protein. J. Immunol. 142, 4428-4434. Martin, R. W., Duffy, J., Engel, A. G., Lie, J. T., Bowles, C. A., Moyer, T. P., and Gleich, G. J. (1990). The clinical spectrum of the eosinophilia-myalgia syndrome associated with L-tryptophan ingestion. Clinical features in 20 patients and aspects of pathophysiology. Ann. Intern. Med. 113, 124-134. Mastrianni, D. M., Eddy, R. L., Rosenberg, H. F., Corrette, S. E., Shows, T. B., Tenen, D. G., and Ackerman, S. J. (1992). Localization of the human eosinophil CharcotLeyden crystal protein (lysophospholipase) gene (CLC) to chromosome 19 and the human ribonuclease 2 (eosinophil-derived neurotoxin) and ribonuclease 3 (eosinophil cationic protein) genes (RNS2 and RNS3) to chromosome 14. Genomics 13, 240-242. Mayeno, A. N., Lin, F., Foote, C. S., Loegering, D. A., Ames, M. M., Hedberg, C. W., and Gleich, G. J. (1990). Characterization of "Peak E," a novel amino acid associated with eosinophilia-myalgia syndrome. Science 250, 1707-1708. Mayeno, A. N., Belongia, E. A., Lin, F., Lundy, S. K., and Gleich, G. J. (1992). 3(Phenylamino)alanine, a novel aniline-derived amino acid associated with the eosinophilia-myalgia syndrome: A link to the Toxic Oil Syndrome? Mayo Clin. Proc. 67, 1134-1139. Molina, H. A., Kierszenbaum, F., Hamann, K. J., and Gleich, G. J. (1988). Toxic effects produced or mediated by human eosinophil granule components on Trypanosoma cruzi. Am. J. Trop. Med. Hyg. 38, 327-334. Moore, P. M., Harley, J. B., and Fauci, A. S. (1985). Neurologic dysfunction in the idiopathic hypereosinophilic syndrome. Ann. Intern. Med. 102, 109-114. Motojima, S., Frigas, E., Loegering, D. A., and Gleich, G. J. (1989). Toxicity of eosinophil cationic proteins for guinea pig tracheal epithelium in vitro. Am. Rev. Respir. Dis. 139, 801-805.
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Newton, D. L., Nicholls, P. J., Rybak, S. M., and Youle, R. J. (1994a). Expression and characterization of recombinant human eosinophil-derived neurotoxin and eosinophilderived neurotoxin anti-transferrin receptor sFv. J. Biol. Chem. 269, 26739-26745. Newton, D. L., Walbridge, S., Mikulski, S. M., Ardelt, W., Shogen, K., Ackerman, S. J., Rybak, S. M., and Youle, R. J. (1994b). Toxicity of an antitumor ribonuclease to Purkinje neurons. J. Neurol. 14, 538-544. Palmieri, M., Carsana, A., Furia, A., and Libonati, M. (1985). Sequence analysis of a cloned cDNA coding for bovine seminal ribonuclease. Eur. J. Biochem. 152, 275-277. Peters, M. S., Rodriguez, M., and Gleich, G. J. (1986). Localization of human eosinophil granule major basic protein, eosinophil cationic protein, and eosinophil-derived neurotoxin by immunoelectron microscopy. Lab. Invest. 54, 656-662. Peterson, C. G. B., Jornvall, H., and Venge, P. (1988). Purification and characterization of eosinophil cationic protein from normal human eosinophils. Eur. J. Haematol. 40, 415-423. Plapp, B. V. (1973). Mechanisms of carboxymethylation of bovine pancreatic nucleases by haloacetates and tosylglycolate. J. Biol. Chem. 248, 4896-4900. Prick, J. J. W., Gabreels-Festen, A. A. W. M., Korten, J. J., and van der Wiel, T. W. M. (1988). Neurological manifestations of the hypereosinophilic syndrome (HES). Clin. Neurol. Neurosurg. 90, 269-273. Ribo, M., Beintema, J. J., Osset, M., Fernandez, E., Bravo, J., De Llorens, R., and Cuchillo, C. M. (1994). Heterogeneity in the glycosylation pattern of human pancreatic ribonuclease. Biol. Chem. Hoppe-Seyler 375, 357-363. Rosenberg, H. F. (1995). Recombinant human eosinophil cationic protein. Ribonuclease activity is not essential for cytotoxicity. J. Biol. Chem. 270, 7876-7881. Rosenberg, H. F., Ackerman, S. J., and Tenen, D. G. (1989a). Human eosinophil cationic protein: Molecular cloning of a cytotoxin and helminthotoxin with ribonuclease activity. J. Exp. Med. 170, 163-176. Rosenberg, H. F., Tenen, D. G., and Ackerman, S. J. (1989b). Molecular cloning of the human eosinophil-derived neurotoxin: A member of the ribonuclease gene family. Proc. Natl. Acad. Sci. U.S.A. 86, 4460-4464. Rybak, S. M., Saxena, S. K., Ackerman, E. J., and Youle, R. J. (1991). Cytotoxic potential of ribonuclease and ribonuclease hybrid proteins. J. Biol. Chem. 266, 21202-21207. Saxena, S. K., Rybak, S. M., Davey, R. T., Youle, R. J., and Ackerman, E. J. (1992). Angiogenin is a cytotoxic, tRNA-specific ribonuclease in the RNase A superfamily. J. Biol. Chem. 267, 21982-21986. Silver, R. M. (1993). Eosinophilia-myalgia syndrome, toxic-oil syndrome, and diffuse fasciitis with eosinophilia. Curr. Opin. Rheum. 5, 802-808. Slifman, N. R., Loegering, D. A., McKean, D. J., and Gleich, G. J. (1986). Ribonuclease activity associated with human eosinophil-derived neurotoxin and eosinophil cationic protein. J. Immunol. 137, 2913-2917. Sorrentino, S., and Glitz, D. G. (1991). Ribonuclease activity and substrate preference of human eosinophil cationic protein (ECP). FEBS Lett. 288, 23-26. Sorrentino, S., Tucker, G. K., and Glitz, D. G. (1988). Purification and characterization of a ribonuclease from human liver. J. Biol. Chem. 263, 16125-16131. Sorrentino, S., Glitz, D. G., Hamann, K. J., Loegering, D. A., Checkel, J. L., and Gleich, G. J. (1992). Eosinophil-derived neurotoxin and human liver ribonuclease: Identity of structure and linkage of neurotoxicity to nuclease activity. J. Biol. Chem. 267,1485914865.
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Spry, C. J. F. (1993). The idiopathic hypereosinophilic syndrome. In "Eosinophils: Biological and Clinial Aspects" (S. Makino and T. Fukuda, eds.), pp. 403-419. CRC Press, Boca Raton, Florida. Strydom, D. J., Fett, J. W., Lobb, R. R., Alderman, E. M., Bethune, J. L., Riordan, J. F., and Vallee, B. L. (1985). Amino acid sequence of human tumor derived angiogenin. Biochemistry 24, 5486-5494. Varga, J., Uitto, J., and Jimenez, S. A. (1992). The cause and pathogenesis of the eosinophilia-myalgia syndrome. Ann. Intern. Med. 116, 140-147. Venge, P. (1993). Serum measurements of eosinophil cationic protein (ECP) in bronchial asthma. Clin. Exper. Allergy 23, (Suppl), 3-7. Venge, P. (1994). Eosinophil activity in bronchial asthma. Allergy Proc. 15, 139-141. Weickmann, J. L., Elson, M., and Glitz, D. G. (1981). Purification and characterization of human pancreatic ribonuclease. Biochemistry 20, 1272-1278. Woolley, K. L., Adelroth, E., Woolley, M. J., Ellis, R., Jordana, M., and O'Byrne, P. M. (1995). Effects of allergen challenge on eosinophils, eosinophil cationic protein, and granulocyte-macrophage colony-stimulating factor in mild asthma. Am. J. Respir. Crit. Care Med. 151, 1915-1924. Wu, Y., Mikulski, S. M., Ardelt, W., Rybak, S. M., and Youle, R. J. (1993). A cytotoxic ribonuclease: Study of the mechanism of onconase cytotoxicity. J. Biol. Chem. 268, 10686-10693. Wu, Y., Saxena, S. K., Ardelt, W., Gadina, M., Mikulski, S. M., De Lorenzo, C., D'Alessio, G., and Youle, R. J. (1995). A study of the intracellular routing of cytotoxic ribonucleases. J. Biol. Chem. 270, 17476-17481. Yamaoka, K. A., Miyasaka, N., and Kashiwazaki, S. (1991). L-Tryptophan contaminant "Peak E" and interleukin-5 production from T cells (letter). Lancet 338, 1468. Yamaoka, K. A., Miyasaka, N., lnuo, G., Saito, I., Kolb, J. P., Fujita, K., and Kashiwazaki, S. (1994). 1,1'-Ethylidencbis(tryptophan) (Peak E) induces functional activation of human eosinophils and interleukin 5 production from T lymphocytes: Association of eosinophilia-myalgia syndrome with a t.-tryptophan contaminant. J. Clin. Immunol. 14, 50-60. Yasuda, T., Mizuta, K., Sato, W., and Kishi, K. (1990). Purification and characterization of a ribonuclease from human spleen: Immunological and enzymological comparison with nonsecretory ribonuclease from human urine. Eur. J. Biochem. 191, 523-529. Yazdanbakhsh, M., Tai, P., Spry, C. J. F., Gleich, G. J., and Roos, D. (1987). Synergism between eosinophil cationic protein and oxygen metabolites in killing of schistosomula of Schistosoma mansoni. J. Immunol. 138, 3443-3447. Young, J. D., Peterson, C. G. B., Venge, P., and Cohn, Z. A. (1986). Mechanism of membrane damage mediated by human eosinophil cationic protein. Nature (London) 321, 613-616.
14 Structure and Function of Angiogenin J A M E S F. R I O R D A N Center for Biochemical and Biophysical Sciences and Medicine Harvard Medical School Boston, Massachusetts 02115
I. Introduction II. Angiogenesis A. Other Angiogenic Molecules B. Biological Assays III. Isolation of Angiogenin A. Human Angiogenin B. Bovine Angiogenin C. Other Species IV. Characterization as Member of the Ribonuclease Family A. Primary Structure B. Tertiary Structure C. Enzymatic Activity D. Substrate Specificity E. Inhibition by RNase Inhibitor F. Functional Residues V. Relationship of RNase Activity and Angiogenic Activity VI. Interaction with Endothelial Cells A. Binding to Endothelial Cells B. Induction of Second Messengers C. Effect on Endothelial Cell Growth D. Angiogenin-Binding Proteins E. Nuclear Translocation of Angiogenin VII. Angiogenin Enhancement of Actin Acceleration of Plasminogen Activation VIII. Angiogenin Support of Endothelial Cell Adhesion IX. Mechanism of Action X. Biology of Angiogenin A. Angiogenin and Wound Healing B. Antiangiogenin Antibody Suppression of Tumor Growth XI. Epilogue References
445 RIBONUCLEASES: STRUCTURES AND FUNCTIONS
Copyright 9 1997 by Academic Press, Inc. All rights of reproduction in any form reserved.
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I. INTRODUCTION Angiogenin is, in many ways, the most unusual member of the ribonuclease superfamily. It shares 33% sequence identity with bovine pancreatic RNase A and has structurally equivalent counterparts for the two histidines and one lysine that comprise the catalytic residues for ribonucleolytic activity (Strydom et al., 1985; Kurachi et al., 1985). Nevertheless, early studies indicated that it was virtually inactive toward R N A substrates. With time it was discerned that angiogenin could, in fact, catalyze the cleavage of ribosomal RNA, albeit very weakly, and it seemed like a unique property of the protein inasmuch as there was no apparent activity with more conventional RNase A substrates (Shapiro et al., 1986, 1987a). Eventually angiogenin was established as a true ribonuclease with the same kind of specificity as RNase A (Shapiro et al., 1987b; Rybak and Vallee, 1988). It cleaves preferentially on the 3' side of pyrimidines to generate a cyclic phosphate product that is subsequently hydrolyzed, but it is 4 to 6 orders of magnitude less active in routine assays. Despite such seemingly minuscule potency, this ribonucleolytic activity is absolutely critical to the biological function of angiogenin (Fett et al., 1985; Shapiro and Vallee, 1987, 1989; Shapiro et al., 1989), which is, supposedly, to induce the growth of new blood vessels under pathological and perhaps normal circumstances. Indeed, it is the only ribonuclease that can stimulate blood vessel formation in standard assays for angiogenesis (Fettet al., 1985). It should be noted that angiogenin is expressed in a wide range of different human cells, which suggests that it may have other biological functions as well (Rybak et al., 1987; Moenner et al., 1994). One such function may be to inhibit the degranulation of polymorphonuclear leukocytes (Tschesche et al., 1994). Angiogenin is one of at least eight polypeptides that have been shown to possess angiogenic activity (Folkman and Klagsbrun, 1987; Schott and Marrow, 1993), and is the only one thus far to have been identified and purified based solely on its ability to stimulate such neovascularization. The others were first characterized in terms of some other property and were only later shown to be angiogenic. Much has been learned about the features of the angiogenin structure that are important both for its ribonucleolytic as well as its angiogenic activity. In contrast, there are wide gaps in our knowledge of its biological f u n c t i o n and mechanism of action. In this regard, it resembles the other angiogenic polypeptides for which mechanistic information is also sparse. Of course, at this juncture
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it may be understandable that so little is understood, given the complexity of the angiogenic process. This is a rapidly developing field, however, and it offers considerable promise of fundamental breakthroughs in the foreseeable future.
II. A N G I O G E N E S I S
Embryonic development, wound healing, endometrial proliferation, and several other physiological processes are all dependent on angiogenesis, as are a variety of pathological conditions such as diabetic retinopathy, psoriasis, arthritis, and especially neoplasia (Folkman and Cotran, 1976). Blood vessels have long been known to proliferate in the vicinity of solid tumors. Based on this and other evidence, Folkman (1971) proposed that these tumors would be unable to grow beyond a certain limiting volume or to metastasize unless they elicited a blood supply from the host that would provide nutrients, remove wastes, and allow malignant cells to migrate to distant sites. Messenger molecules released from the tumor cells were the presumptive initiators of the new blood vessel growth. Molecular details of the mechanism of tumor-induced angiogenesis are only beginning to emerge. It is believed that one or more angiogenic substances are released either from the tumor, or from its extracellular matrix by the action of the tumor-secreted molecules, or both. This substance(s) diffuses radially until it encounters an existing microvessel, activates its endothelial cells, and induces capillary sprouting. One of the early consequences of endothelial cell activation is the production of proteases such as plasminogen activator, plasmin, and collagenase, which are needed to degrade the basement membrane surrounding the microvessel and to allow the cells to migrate, proliferate, and form microtubules. The tubules are drawn toward the tumor either by selective adhesion or chemotaxis, and in the process anastomose with other tubules to form capillary loops through which blood flows. Eventually a bed of capillaries is constituted, complete with basement membrane and even pericytes, and this supplies the tumor with its nutritional needs and supports proliferation (Blood and Zetter, 1990). It would seem improbable that a single molecule, especially a ribonuclease, would be directly responsible for all of these steps and hence it is not too surprising that so many angiogenic factors have been identified.
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Some cooperativity between them might be expected, but such interactions are unknown. Not all angiogenic molecules have the ability to elicit the same range of biological responses in vitro. Some are pluripotent and can stimulate endothelial cell proliferation and migration, as well as induce secretion and production of proteases, whereas others may not act on endothelial cells at all (Blood and Zetter, 1990).
A. Other Angiogenic Molecules The biological and biochemical properties of the various polypeptides that have been shown to possess angiogenic activity differ considerably (Klagsbrun and D'Amore, 1991; Zagzag, 1995). Among them, angiogenin is the only ribonuclease. Members of the fibroblast growth factor (FGF) family, comprising at least nine related mitogens of which acidic and basic FGF are the best known, stimulate endothelial cells to produce most of the features of new capillary growth. They also stimulate growth of smooth muscle, fibroblast, and epithelial cells and have a strong affinity for heparin. Epidermal growth factor (EGF) is similarly pleiotropic and acts on a range of cell types. Transforming growth factor a (TGF-c~) has potent angiogenic activity and competes with EGF for binding to its receptors. TGF-/3, which is angiogenic in vivo, was first identified on the basis of its ability to induce a transformed state in cultured nonneoplastic rat kidney fibroblasts. It also induces mitogenesis of osteoblasts and Schwann cells, and morphogenesis of fibroblasts, but it inhibits proliferation of epithelial and, paradoxically, endothelial cells. Similarly tumor necrosis factor ct (TNF-c~) inhibits endothelial cell growth in vitro but is angiogenic in vivo and, hence, is considered an indirect angiogenic factor. Perhaps the most specific angiogenic protein is vascular endothelial growth factor (VEGF), actually a family of secreted homodimeric glycoproteins that are the only known selective mitogens for endothelial cells (Ferrara et al., 1992). They also promote vascular permeability, stimulate monocyte migration, and induce the synthesis of plasminogen activator and collagenase. Platelet-derived growth factor (PDGF) is angiogenic in vivo and stimulates the growth of capillary but not macrovascular endothelial cells. It is also an effective mitogen for fibroblasts and smooth muscle cells. Platelets produce another angiogenic protein that, like FGF, lacks a secretory signal peptide and stimulates chemotaxis of endothelial cells. Although it is referred to as platelet-derived endothelial cell growth
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factor (PD-ECGF), it is not a classic mitogen. It resembles angiogenin in this regard, and also in having enzymatic activity. In fact, P D - E C G F has been identified as a thymidine phosphorylase (Moghaddam and Bicknell, 1992), and its enzymatic activity is essential for its angiogenic function (Finnis et al., 1993). The mechanism by which it stimulates thymidine uptake in endothelial cells is thought to be by modulating the concentration of thymidine in the culture medium and in the cell. However, it may also lead to the production of 2-deoxy-o-ribose, which can be generated from thymidine and has been shown to be both angiogenic and chemotactic (Haraguchi et al., 1994). With the exception of angiogenin, all of these angiogenic proteins were first isolated based on some other biological property and were only subsequently recognized to also have angiogenic activity. Cellular receptors for these proteins, other than angiogenin and PD-ECGF, have been identified and characterized, and their signal transduction pathways have been the subject of extensive investigation (Klagsbrun and D'Amore, 1991).
B. Biological Assays
Several different types of angiogenesis assays have been described, all of which attempt to measure new capillary growth. Perhaps the most useful methods have been variations of the chick chorioallantoic membrane (CAM) assay (Knighton et al., 1977). Access to this membrane, which covers the developing chick embryo, is gained either by creating a window in the shell or by cracking the egg and transferring the embryo to a petri dish. A test sample is incorporated into a slow-release polymer pellet such as methylcellulose, or dried onto a plastic coverslip, which is then placed on the membrane. After 2 or 3 days blood vessel growth is recorded as either positive or negative. It is important to use multiple eggs (10-25 per set) along with a positive and negative (water) control. At least three sets are required to evaluate statistical significance (Fett et al., 1985; Shapiro et al., 1987a). A more complicated assay involves implantation of the sample, again in a slow-release pellet, into a pocket created in the normally avascular cornea of a rabbit eye at a fixed distance from the peripheral blood vessels (Langer and Folkman, 1976). Capillary growth extending through the cornea toward the sample can be seen after several days. Other assays measure the infiltration of blood vessels
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into sample-impregnated polyvinyl sponges implanted subcutaneously into mice. The hamster cheek pouch, the rabbit ear, and several other animal models have also been employed. A so-called in vitro assay uses collagen gels and measures endothelial tube formation in culture in either two or three dimensions (Jimi et al., 1995). It is important to emphasize that although the subject of angiogenesis has become increasingly popular, no single assay method has been adopted by the scientific community as the standard of reference. Moreover, because artifacts abound, a single positive response can never be accepted as evidence for true angiogenesis. Multiple assays over a range of concentrations are essential. There is no agreement on the type of endothelial cell that should be employed in so-called in vitro assays, no specification for the nature of the substratum on which cells should be grown, and no good way to recognize artifacts. Despite these handicaps, progress has been achieved often by monitoring some other biological property of an angiogenic agent, such as mitogenesis, induction of cell migration, or enzymatic activity.
!I!. I S O L A T I O N O F A N G I O G E N I N
A. Human Angiogenin Early work on angiogenesis factors was motivated, in part, by their relationship to cancer. The finding by Folkman that the growth of experimental tumors required a blood supply, together with the observation of tumor-associated blood vessel proliferation, led him to suggest that tumor cells release a diffusible mediator of vessel growth (Folkman, 1971). Such a substance should therefore be present in tumorconditioned medium, and this has been established: conditioned media from different types of tumors are indeed angiogenic, though not to the same degree (Olson et al., 1994). The colon carcinoma cell line HT-29 proved to be particularly advantageous for isolation purposes. These cells can be kept viable for many weeks in the absence of added serum and they continue to secrete protein (Alderman et al., 1985). Angiogenin was first purified from this conditioned medium by a series of chromatographic steps. A typical preparation begins with culture medium obtained from large-scale cell factories. After acidification, clarification, and buf-
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fer exchange the sample is applied to carboxymethyl (CM)-cellulose, and the bound fraction is eluted with salt and further fractionated by reversed-phase high-performance liquid chromatography (HPLC). Homogeneous active product is obtained in a yield of about 0.5/~g/liter of medium (Fett et al., 1985). At this point in the history of angiogenin its relationship to pancreatic RNase was unknown, of course, and hence isolation had to be based solely on CAM assays. These are time-consuming and tedious, and call for rather large quantities of material because multiple sets of eggs are required for each fractionation step. Despite these difficulties the purified protein finally obtained displayed activity with as little as 0.5 ng per egg. It was a very basic (pl > 9.5), single-chain protein of molecular weight ---14,000, with a blocked N-terminal residue. The presence of angiogenin in medium conditioned by a human tumor cell line raised the question of whether it could be found in normal tissues or body fluids, or if it is a tumor-specific protein. Normal human plasma was therefore examined for its content of angiogenin owing to its ready availability and the potential diagnostic or prognostic relevance of any positive finding. A modification of the procedure developed for isolating angiogenin from HT-29 cell medium produced a plasma protein that was physically and functionally indistinguishable from angiogenin obtained previously (Shapiro et al., 1987a). The yield was 60-150 p~g/ liter of plasma, more than 100-fold greater than from conditioned medium. More importantly, the studies demonstrated that angiogenin is not a protein synthesized specifically by tumor cells but one that is present in normal plasma. They also opened the way to obtain angiogenin from other species. Recombinant angiogenin was first expressed in transformed BHK cells under the transcriptional control of a metallothionein promoter and this system increased the yield to 400 p,g/liter (Kurachi et al., 1988). Subsequently a system was developed for expression of angiogenin as inclusion bodies in Escherichia coli (Den6fle et al., 1987; Shapiro et al., 1988b). The solubilized product, Met-(-1) angiogenin, is purified by cation exchange chromatography and reversed-phase HPLC, and is obtained in amounts of 1-2 mg/liter. It can be converted to the native
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angiogenin. This generates a soluble protein that can be isolated from the cell lysate as pyroGlu-angiogenin also in yields of a few milligrams per liter (Shapiro and Vallee, 1992), and is now used for routine production.
B. Bovine Angiogenin Bovine sera and plasma were examined both as potential sources of a different species of angiogenin and for the implications their angiogenin content might have with respect to studies of cells in culture (Bond and Vallee, 1988). The protein can be obtained from adult bovine plasma in yield of 30-80 ~g/liter and from fetal serum at about 30 ~g/liter. Because fetal bovine serum is typically diluted to 10% or less in cell culture medium, it will contribute at least 3 ng/ml of angiogenin, which could under some circumstances exert a biological effect. Isolation was greatly facilitated by a novel assay (Bond, 1988) based on the capacity of angiogenin to bind to placental RNase inhibitor (see Section IV,E), which thereby prevents it from inhibiting a standard amount of RNase A. The remaining amount of RNase activity is directly proportional to the amount of angiogenin in the test sample. Bovine angiogenin does not have an N-terminal pyroGlu but otherwise is closely similar to the human protein both in terms of structure and function. The protein was also found to be present in bovine milk (Maes et al., 1988), remarkably at 10 times higher concentration than in serum, from which it can be isolated readily by the usual three-step chromatographic procedure (Bond and Vallee, 1988). Human milk, on the other hand, contains extremely small amounts of angiogenin (R. Shapiro, 1996, personal communication).
C. Other Species Angiogenins have also been purified from rabbit, pig, and mouse serum (Bond et al., 1993). All are homologous to the pancreatic ribonucleases, are highly basic, contain the essential catalytic residues, and display weak activity toward RNase A substrates. However, like the human and bovine proteins they are potent inhibitors of in vitro protein synthesis (see Section IV,D). Preliminary studies have indicated the presence of an angiogeninlike protein in sheep plasma, but not horseshoe crab hemolymph.
14
453
Structure and Function of Angiogenin
IV. CHARACTERIZATION AS MEMBER OF THE RIBONUCLEASE FAMILY A. Primary Structure The primary structures of human, bovine, rabbit, pig, and mouse angiogenins, deduced either from direct protein sequencing or from c D N A or both (Bond and Strydom, 1989; Bond et al., 1993), are shown in Fig. 1. The human protein contains 123 amino acids whose sequence is 33% identical to that of bovine pancreatic RNase A. Sequence identity between mouse, rabbit, pig, and bovine angiogenins, relative to human, is
1
Human
Mouse
Pig
Bovine
AQ
Human
R
Mouse
R
Pig
Q
Rabbit Bovine
D S
Rabbit
R N
DS
K
DY
Y DAKP
R P C K D R N T F I H G NK
R
E N L
70
I S K S S
V
Y R E N L
M
S K S P
V
Y N
R S K S P
I
- N
F F
D L
i00
R
T A G
Rabbit
R
T S G
G
T E D S
S A G
T A G
I S K S S
F
N VVVA
S
N I V
F F
E
I S KS
H V V T
V
ERM
D GR
E S I
K
N D E
80
T
V
N D I K A
I
G
Q P C Q
L H G G S'P W P P C
T
H K
G G ~ NRP
H K G G
PWP S RP
P C R
P C
G
P C R
120
C E N G L P V H
L
Q
I A C E N G L P V H
F
E
V F Q Q K V H
E
F I T P R H
I A C E N G L P V H
I A VA
C E N G L P V H
I V V G C E N G L P V H
F F
F
E
E
G G
E D R 90
H V G G
K
NN G
E D K
G G ~ P RP
ii0
K
N D K N G
H T
I
F NM
GA-
I
R
K
K
E N K'N
T T
I
D I K A
30
ETM
60
i K A I
I K A I F V H G N K G S I K D V
R P C K E V N T F I H G T RN
S P C K D T N T
E S I
D DR
N D R
F I H G N K S N
G
D
D D R
F
50
S P C K D V N T
Human
R
F L T Q
S
Y RG
Bovine
Y DAKP
40
Bovine
Pig
K
Q
20
S P C K D I N T F I H G N K R S
G
Y G K N
R
Q H Y DAKP
F LT
F LT
IH
Rabbit
Mouse
K
TH
H R
Pig
Q
T K F L T Q H H DAKP
KH
ED
Human
Mouse
i0
T H F L T Q H Y DAKP
N S
I F R R P
FFSL
F I I T S Q
Fig. 1. Alignment of mammalian angiogenin sequences. Numbering is based on the human protein; residues conserved in all five angiogenins are boxed. From Bond et al. (1993), with permission of Elsevier Science B.V.
454
James F. Riordan
75, 73, 66, and 64%, respectively. Among all five, 62 of the residues are invariant and another 15 are substituted conservatively. None undergo posttranslational modification apart from pyroGlu formation. All contain a much higher percentage of basic residues than any of the other RNases except for eosinophil cationic protein (ECP). This highly basic character is reminiscent of other physiologically active proteins such as plateletderived growth factor, basic FGF, and y-interferon. Of the total invariant plus conservative residues, 52 also occur at the same position in at least some of the pancreatic RNases and are likely involved in maintaining the overall RNase structure (Bond et al., 1993). The remainder are unique to the angiogenins and they tend to cluster in certain regions of the tertiary structure (discussed in detail in Section IV,B). In particular, residues 54, 62, 64, 69, 70, 109, and 110 are all spatially close, with residues 62-69 occupying a large, convoluted, exposed loop (Acharya et al., 1994). This area contains four deletions with respect to RNase A and differs significantly from the same area in other RNases. In all known mammalian RNases there is a disulfide bond between Cys-65 and Cys-72 (Beintema et al., 1988). Both of these Cys residues have been replaced in all of the angiogenins, and, in addition, two intervening residues are deleted. As will be shown, this region of the protein appears to play a special role in angiogenic function. Other regions with groups of conserved residues that differ from the RNases include residues 5-11 and 16-21, which suggests that the N-terminal region of angiogenin might have a unique functional role. Mutagenesis of angiogenin by replacing residues 8-22 with 7-21 of RNase A had no effect on its enzymatic activity but increased its angiogenic potency and decreased its capacity to inhibit cell-free protein synthesis (Bond and Vallee, 1990). The angiogenins differ from the RNases in not having a positively charged residue at either position 65 or 117, and this has been thought to contribute to their very low ribonucleolytic activity (Beintema et al., 1988). This may be a partial explanation, but the occlusion of the substrate B~ binding pocket by the side chain of residue 117, as observed by X-ray crystallography, might well be a more significant negative factor for in vitro catalysis.
B Tertiary Structure A three-dimensional structure of angiogenin was computed, based on its sequence homology to pancreatic RNase A, by minimization of its
14
Structure and Function of Angiogenin
455
conformational energy, assuming that the structures of the two proteins would be quite similar (Palmer et al., 1986). This structure served as a useful model for several years until the crystal structure of the human protein was determined by X-ray analysis (Acharya et al., 1994). Actually, many of the structural predictions were in reasonable agreement with the results found by crystallography (Allen et al., 1994). The overall folds of angiogenin and RNase A are indeed similar (Fig. 2) as has also been determined by 1H N M R spectroscopy (Reisdorf et al., 1994). The only major differences are in the surface loop regions and, especially, in the C-terminal segment starting at residue 116 (the calculated structure omitted residues 120-123). Residues comprising the active center, i.e., His-13, Lys-40, and His-114, occupy essentially the same positions in angiogenin as in RNase A. These residues constitute what in RNase A has been called the P1 site, where catalysis of phosphodiester bond cleavage occurs (Arus et al., 1982). In addition, angiogenin has a specificity-defining, B1 site--which includes Thr-44mwhere the pyrimidine moiety that contributes the scissile 3'-oxygen binds, and a B2 site where the nucleoside that contributes the scissile 5'-oxygen binds. These substrate-binding pockets are not identical to those in RNase, however. The most startling difference is seen at the B1 site. In RNase it is an open pocket with Thr-45 at one side and Ser-123 at the other. In angiogenin there is no "pocket." It is occupied by the side chain of Gin-117, a consequence of the markedly different orientation of the seven Cterminal residues of angiogenin. When the RNase inhibitor, uridine vanadate, is modeled into the putative B1 site of angiogenin, it is seen that the Gln-117 side chain passes through the uridine ring (Fig. 3). Because angiogenin has detectable ribonucleolytic activity, even with C2',3'-P, and its substrate specificity is sensitive to mutations involving Thr-44, it seems likely that substrate must bind to the B~ site. For this to occur the protein would have to undergo a conformational change. To date, a crystal structure of an angiogenin/B~-ligand complex has not been obtained that would confirm such a change, but mutagenesis of Gln-117 is consistent with this view: replacing it by Ala or Gly increases activity 11- to 18-fold and 21- to 30-fold, respectively, depending on substrate, but without any change in specificity (Russo et al., 1994) (see later, Table I). Presumably, these mutations facilitate the conformational change. In the native protein, and in the absence of an appropriate conformational change trigger, only a very small fraction of the molecules would exist in the active conformation, thus accounting for the low ribonucleolytic activity. These findings have implications for the mechanism of action in vivo, as will be discussed later.
Fig. 2. The polypeptide fold for angiogenin (left) and RNase A (right), drawn with the program MOLSCRIPT. From Acharya et al. (1994), with permission of National Academy of Science.
Fig. 3. (Left) Crystal structure of the active site of human angiogenin with uridine vanadate (UV, in black) added by modeling. (Right) Crystal structure of the RNase A-UV complex. Selected side chains are shown. From Acharya et al. (1995), with permission of National Academy of Science.
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James F. Riordan
Another region of angiogenin that differs significantly from RNase is composed of two adjacent loops involving residues 59-68 and 108-110. The former, which replaces the fourth disulfide loop of RNase that is missing from angiogenin, has little structural relationship to its RNase counterpart in terms of either backbone or side chain positions. In RNase this region constitutes one side of the B2 binding site, but in angiogenin it seems to be associated with binding to cell surface receptors and is therefore critical to angiogenesis but not to ribonucleolytic activity (Hallahan et al., 1991). Indeed, the B2 site is not well conserved in angiogenin. Of the three RNase A residues thought to hydrogen bond to the base (Gln-69, Asn-71, and G l u - l l l ) (Zegers et al., 1994; Nogu6s et al., 1995), only two have counterparts in angiogenin: Asn-71 corresponds to Asn-68 and G l u - l l l corresponds to Glu-108. Mutagenesis studies indicate that only Asn-71 has a significant role in the activity of RNase (Tarragona-Fiol et al., 1993) and that Glu-108 has minimal importance in angiogenin (Curran et al., 1993a). There does not appear to be Gln-69 equivalent in angiogenin, but Gln-69 has no significant B2 binding function in RNase A (Tarragona-Fiol et al., 1993). The crystal structure of bovine angiogenin has also been determined and is closely similar to that of the human protein, including a B~ site that again is blocked in this case by Glu-ll8, the counterpart of Gin117 (Acharya et al., 1995). Of interest is the fact that the bovine protein has a putative Arg-Gly-Asp (RGD) recognition element not present in human angiogenin. However, its conformation differs considerably from those found in proteins that have integrin-binding activity.
C. Enzymatic Activity It might be informative to recount the steps leading to the present understanding of the enzymatic activity of angiogenin. From its primary structure it seemed likely that angiogenin should have ribonucleolytic activity. Not only did it share a high degree of sequence identity with all other RNases, but it contained all the important catalytic residues, His-13, His-114, and Lys-40. It was therefore surprising that initial studies, prompted by this homology failed to detect any enzymatic activity. The first indication that it might be ribonucleolytic emerged only after an intensive effort was made to find a possible RNA substrate (Shapiro et al., 1986). A large assortment of prospective candidates was examined from which 28S and 18S ribosomal RNA were the only ones found to
14
Structure and Function of Angiogenin
459
undergo cleavage through the action of angiogenin. Compared to RNase A, this activity was extremely low and it was necessary to employ elaborate HPLC purification procedures to be sure that the activity was not due to a contaminating RNase. A characteristic feature of the activity, which also helped to rule out a contaminant, was the size of the degradation products--from 100-500 nucleotides, which is much larger than those generated by RNase A. Subsequent studies confirmed the ribonucleolytic activity of angiogenin. The protein was shown to abolish cell-free protein synthesis apparently by specifically inactivating ribosomal RNA (St. Clair et aL, 1987, 1988). It was even more effective than RNase A in this system. A more precise assay for the ribonucleolytic activity of angiogenin was then developed based on hydrolysis of tRNA (Shapiro et al., 1987b). Conditions were optimized to give detectable results with as little as 0.1/xg of angiogenin. Later, a sensitive HPLC method for quantitation of substrate and reaction products was developed that made it possible to measure kcat/Km values for dinucleoside phosphates (Harper and Vallee, 1988b; Shapiro et al., 1988b). Values obtained for recombinant human Met-(-1) angiogenin and various mutants with dinucleotides are listed in Table I. They are 5 to 6 orders of magnitude lower than those for RNase A. This assay has been modified to allow determination of individual kinetic parameters (Russo et al., 1994). Previous efforts in this regard has been unsuccessful owing to anomalies in the plots of kinetic data (Curran et al., 1993a). The modified assay conditions, however, gave linear Lineweaver-Burk plots over a range of substrate concentrations at least 3-fold above and below Km. For angiogenin acting on CpA, gm is 62 mM and kcat is 0.69 sec -~. Both values are quite different from those for RNase A: The former is about 120-fold greater and the latter is more than 6000-fold less. This assay was also employed to demonstrate that angiogenin catalyzes the hydrolysis of both cytidine and uridine cyclic phosphates. It is interesting that the kcat/Km values for these substrates are only about 25-fold lower than those for the best CpN and UpN substrates, whereas for RNase A the difference is about a factor of 650.
D. Substrate Specificity The overall ribonucleolytic specificity of angiogenin was first established by examination of the partial digests generated from its action on
460
James F. Riordan
TABLE I Activity of Angiogenin Mutants toward Dinucleoside 3',5'-Phosphates and t R N A ~
kcat/Km (M -l sec -1) Mutant
CpA
Met-(-1) angiogenin Qll7G Qll7A Dll6N Dll6A Dll6H Dll6P Dll6S S118D Sll8R T44H T44A T44D ARH-I (58-70) ARH-II (38-41) ARH-IV (I + II) ARH-III (8-22) Angiogenin K Angiogenin E E108Q E108K N61D N109D RNase A
CpG
12.0
1.0
4.0
310 172 7 20 44 0.6 5.7 7.3 3.2 2.3 0.5 0.7 2800 220
15 7.9 0.5 0.9 3.0 0.2 1.9 0.6 0.6 1.1 0.4 2.1 220 75
116 56 -~ -0.3 9.5 2.1 0.7 0.2 0.1 0.1 50 50
----~ 0.07 4.6 0.2 0.1 0.1 0.07 0.5 11 2
30 18 8 15 18 0.1 16 0.3 0.4 0.02 0.04 0.9 300 20
7200
450
120
18
942
13
0.8
17.4 16.8 16 14 10.9 13.9 9.7 x 106
0.8 4.4 1.6 1.5 0.6 0.7 3.9 • 106
UpG
tRNA (rel.)
UpA
0.4
3.4
0.4
4.1 5.1 2.4 3.5 3.6 4.4 5.6 • 105
0.2 0.2 0.2 0.3 ~ ~ 2.4 • 105
1.0
1.0 1.0 0.4 1.4 0.7 1.0 1.0 5.5 • 104
a Activities toward dinucleotides were measured in 33 mM MES and 33 mM NaCI, pH 5.9 at 37~ (except for Q l l 7 G and Q l l 7 A , which were in 0.2 M MES, pH 5.9 at 25~ as described by Hallahan et al. (1991). Activities toward tRNA were determined by a precipitation assay (Shapiro et al., 1987b) and are relative to angiogenin.
RNA
molecules
E. c o l i ( R y b a k pyrimidines,
of known and
sequence,
Vallee,
as d o e s R N a s e
i.e., t h e 5S R N A s
1 9 8 8 ) . It c l e a v e s
from yeast and
almost
exclusively after
A, and most often when
t h e n e x t b a s e is
14
Structure and Function of Angiogenin
461
adenine. It is interesting that in this case the products of hydrolysis are 5-12 nucleotides in length, much shorter than with 18S and 28S rRNA. More quantitative studies with dinucleoside phosphates indicated that angiogenin favors C over U by a factor of about 10 in the N site of NpN' substrates, and A over G by a factor of about 3 in the N' site (Curran et al., 1993b). In contrast, RNase A prefers C over U by only 2-fold in the N site and A over G by 17-fold in N'. The order of base preference in the N' site is, nevertheless, the same for angiogenin and RNase A, A>G>C>U. The evidence to date strongly suggests that the physiological substrate of angiogenin is R N A - - e i t h e r a specific R N A or a specific class of RNAs, but in any case RNA. As noted, it exhibits a characteristic activity toward isolated 28S and 18S rRNA wherein the major products are 100-500 nucleotides in length, in contrast to the much smaller products generated by RNase A. Moreover, relative to RNase A it is more active toward R N A substrates than toward dinucleotides. Most notable is its ability to inhibit protein synthesis by an mRNA-dependent rabbit reticulocyte lysate, which in this case appears to be due to cleavage of rRNA (St. Clair et al., 1987, 1988). Addition of 40 nM angiogenin to the lysate completely destroys its capacity to support protein synthesis after m R N A supplementation. In contrast, under the same conditions, 40 nM RNase A diminishes activity by only 50%. Addition of intact ribosomes to an angiogenin-treated lysate restores protein synthesis activity, but angiogenin-treated ribosomes do not do so when added to a ribosomedependent translation system. The actual site or sites of cleavage of rRNA are unknown and, in fact, it is not even possible to detect any degradation by formaldehyde/agarose gel electrophoresis even though protein synthesis was inhibited completely. This may not be surprising because inhibition apparently occurs at the elongation or termination step of protein synthesis, and inactivation of only a few ribosomes on a polysome could block further synthesis by all the others. The effect of angiogenin on 18S rRNA has been thought to be responsible for inhibition of protein synthesis (St. Clair et aL, 1988). This was shown by treating isolated 40S and 60S ribosomal subunits with angiogenin. Translation was abolished on treatment of the former but not the later. Prolonged exposure of the 40S subunit to angiogenin generated a faster migrating species of RNA, only slightly smaller than 18S rRNA and indicative of limited cleavage at either the 3' or 5' end, or both. Remarkably, naked 18S rRNA is extensively degraded to form characteristic products that are 100-500 nucleotides in length. In contrast, 28S
462
James F. Riordan
rRNA, either naked or in the intact ribosome (or 60S subunit), is always cleaved into the 100- to 500-nucleotide-long fragments. Neither 5.8S nor 5S rRNA is degraded by angiogenin when present in the ribosome, but purified 5S rRNA from yeast is cleaved to fragments of 5-12 nucleotides (Rybak and Vallee, 1988). These results do not identify rRNA as the natural target of angiogenin in vivo. Indeed, it has been shown that injection of angiogenin into frog oocytes abolishes protein synthesis by complete hydrolysis of tRNA, not rRNA (Saxena et al., 1992). The basis for the difference from the above results has not been established. Nevertheless, all of these data indicate that angiogenin acts on RNA substrates and, in addition, they show that RNA structure contributes importantly to its cleavage specificity.
E. Inhibition by RNase Inhibitor Potent inhibitors of RNase have been shown to be present in the cytosol of cells from essentially all mammalian tissues. A detailed discussion of the properties of these inhibitors is presented in Chapter 19 in this volume and, hence, only those aspects pertinent to angiogenin are given here. One inhibitor that has received particular attention in this regard is that isolated from human placentaman acid-labile, sulfhydryldependent protein with a molecular weight of 51,000. It is a single polypeptide chain composed of tandem alternating 28- and 29-residue leucine-rich repeats, a motif often found in proteins that bind tightly to other proteins (Lee and Vallee, 1993). Indeed, it bind~ extremely tightly to RNase A with a 1:1 stoichiometry and a K~ of (4-7) x 10-14 M (Lee et al., 1989). It binds even more tightly to angiogenin and to eosinophilderived neurotoxin with Ki values of (7-9) x 10 -16 M (Lee et al., 1989; Shapiro and Vallee, 1991). This value is one of the lowest ever reported for the binding of two proteins and is even less than that for avidin and biotin. The X-ray structure of the RNase inhibitor from porcine liver has been determined as has that of its complex with RNase A (Kobe and Deisenhofer, 1993, 1995). Some 28 residues of the inhibitor are in contact with the enzyme and all but 4 of these are conserved in the inhibitor from human placenta (and 3 of the 4 are conservative replacements). On the other hand, 24 residues from RNase are involved in contacts with the inhibitor, but less than half of these are conserved in the sequence of
14 Structure and Function of Angiogenin
463
angiogenin (Shapiro et al., 1995). In fact, many of those that are conserved occupy structurally different positions. Mutagenesis, chemical modification, and fluorescence have identified several residues in angiogenin that appear to be involved in its interaction with the inhibitor. These are Lys-40, Arg-5, Arg-32, His-114, and Trp-89, in decreasing order of importance. Some but not all of these have equivalent counterparts in RNase A. Thus, it would seem that the inhibitor is able to interact differently with different members of the RNase superfamily, likely by exercising some degree of flexibility that allows it to accommodate all of them with remarkable affinity. The tandem repeat structure of RNase inhibitor suggested that part of the molecule might not be necessary for interaction with angiogenin. This led to studies that showed that 6 of the 14 central repeats could be deleted without major detriment to binding (Lee and Vallee, 1990a,b; 1993). Removal of repeats 5 to 8 reduced binding to both RNase A and angiogenin by 2500-fold. Removal of repeats 11 and 12 reduced binding to RNase by 630-fold and to angiogenin by 75,000-fold. Nevertheless, the mutant inhibitors still had subnanomolar Ki values. Deletion of residues 1-90 of the porcine liver inhibitor had almost no effect on its binding to RNase A.
A. Funtional Residues
1. C h e m i c a l M o d i f i c a t i o n
Recognition of the homology between RNase and angiogenin, and the subsequent identification of its enzymatic activity, opened the way to employ experimental approaches developed over many years at the hands of RNase to discern the mechanism of action of angiogenin. Chemical reagents specific for lysine, histidine, and arginine all decrease the ribonucleolytic activity of angiogenin, much as they do with RNase A (Shapiro et al., 1987b). One major difference is that with angiogenin, the rate of inactivation for lysine and histidine modification is up to 10fold slower, whereas for arginine modification it is significantly faster than with RNase A. Thus, for example, bromoacetate carboxymethylates N-3 of His-13 and N-1 of His-114, as it does with the comparable residues His-12 and His-119 in RNase A (Shapiro et al., 1988a). However, the reactivity of His-114 is significantly less than that of His-119, perhaps
464
James F. Riordan
because it does not interact with Asp-116 in the same way that His-119 interacts with Asp-121. In addition, with 1-fluoro-2,4-dinitrobenzene, RNase loses 75% of its activity on modification of 1.4 of its 10 lysines (primarily Lys-41). In contrast, 3.8 of the 7 lysines of angiogenin must be modified for a comparable loss of activity, none of which is Lys-40, although this residue is, nevertheless, essential for catalysis as shown by site-specific mutagenesis (Shapiro et al., 1989). No specific arginine residue has been found to be responsible for the enhanced susceptibility of angiogenin to phenylglyoxal (Shapiro and Vallee, 1992). Mutation of Arg-5 or Arg-33 to Ala decreases ribonucleolytic activity by 4- and 7-fold, respectively, but both mutants are still further inactivated by arginine reagents. 2. Site-Directed Mutagenesis
Because chemical approaches had not helped to define the function of Lys-40 in angiogenin, mutagenesis was employed to replace it with glutamine or arginine (Shapiro et al., 1989). In the former case ribonucleolytic activity was decreased below the limit of detection, i.e., at least 2000-fold, and in the latter it was reduced 45-fold. Thus, it seems apparent that Lys-40 fulfills the same functional role in angiogenin as Lys-41 does in RNase A. Similarly, site-directed mutagenesis has shown His-13 and His-ll4 to be equivalent to His-12 and His-ll9 (Shapiro and Vallee, 1989). Replacement of either one by alanine essentially abolishes enzymatic activity toward tRNA. Replacing His-13 by glutamine only reduced activity 300-fold and replacing His-ll4 by asparagine reduced it at least 3300-fold. These last two mutations are considered conservative substitutions; the former would place an NH function in a position analogous to N-3 of His-13, whereas the latter does the same for the N1 of His-ll4. The virtual inactivity of H13A, H l l 4 A , and H l l 4 N is consistent with the widely accepted mechanism of RNase A in which these histidines act as general base and general acid catalysts, respectively. The low activity of H13Q, if real, would suggest that it is still able to assist in either the deprotonation of the ribose 2'-hydroxyl group or stabilization of the transition state, assuming that it has one or the other of these roles in angiogenin. Targets for mutagenesis also have included residues thought to be involved in other functions related to catalysis. One such residue in RNase is Asp-121, which is conserved in all members of the pancreatic RNase family. Its carboxylate side chain is believed to hydrogen bond
14
Structure and Function of Angiogenin
465
to the imidazole N-3 of His-119 and thereby direct N-1 for participation in catalysis (and also activate it for alkylation by bromoacetate). It also hydrogen bonds to Lys-66 and thereby helps fix the 65-72 disulfide loop that forms part of the B2 binding site. Semisynthetic analogs of RNase in which Asp-121 is replaced by asparagine or alanine have only 5 and 48% activity toward yeast RNA, respectively (Lin et al., 1970). X-ray crystallography has revealed that in the former case, the orientation of His-ll9 is changed, perhaps accounting for the diminished activity. Although the precise role of Asp-121 in the catalytic mechanism of RNase A is unknown, it is not surprising that replacement would reduce activity (DeMel et al., 1992). It is not a crucial role, however, because the alanine analog still has appreciable activity. Mutagenesis of Asp-ll6 in angiogenin, the counterpart of Asp-121 in RNase, gave surprising results (Harper and Vallee, 1988b) (see Table I). Instead of the enzymatic activity being decreased, it was enhanced substantially. The Dll6N, D l l 6 A , and D l l 6 H mutants had 8-, 15-, and 18-fold greater activity toward tRNA than the native protein. The increase in activity toward dinucleotide substrates was much less and for D116N activity actually decreased. Six more mutants were examined to explore the basis for this phenomenon and their activities toward tRNA ranged from 10-fold less (Dll6P) to 16-fold greater (Dll6S) (Curran et al., 1993b). Subsequent structural information obtained by X-ray crystallography of angiogenin provided the needed insight (Acharya et al., 1994, 1995). The orientation of the side chain of Asp116 in angiogenin differs from that of Asp-121 in RNase. It hydrogen bonds directly to the - - O H of Ser-ll8, and probably not to His-ll4. Although the structures of the Asp-116 mutants have not yet been determined, it seems likely that any such mutation would disrupt the interaction with Ser-ll8. This could have major consequences in terms of Gln117, whose side chain occupies the B1 bdining pocket and interferes with substrate binding. It is notable, however, that mutation of Ser-118 either to Arg or to Asp diminishes activity severalfold without altering specificity (Curran et al., 1993a), especially because its RNase A conterpart, Ser-123, is thought to play a key role in accommodating both pyrimidines in the B1 site (Gilliland et al., 1994; Zegers et al., 1994). Other residues have also been the targets of mutagenesis primarily because of their correspondence to residues in RNase A that by Xray crystallography have been assigned some role related to catalysis. Mutations of Thr-44, which is thought to be a component of the B1 binding site in RNase A, are particularly interesting in that they alter
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James F. Riordan
both activity and specificity dramatically (Curran et al., 1993a) (see Table I). Moreover, the nature of the replacement is critical. Ala and His decrease activity toward tRNA by 25- and 40-fold, respectively, and reduce cleavage of cytidylyl more than uridylyl dinucleotides. Asp, on the other hand, does not alter activity toward tRNA, decreases that with CpN dinucleotides up to 40-fold, and doubles that toward UpA. These changes are most remarkable in view of the fact that the B1 site in angiogenin is occluded by Gln-ll7. Such responses to mutagenesis of Thr-44 would be understandable if, in fact, substrate does bind to the B1 site during catalysis, which can only happen if some kind of structural reorganization of the C-terminal region of the protein takes place.
3. Regional Mutagenesis: RNase/Angiogenin Chimeras An alternative approach to exploring the structure-function relationships of angiogenin has been to identify and mutate those regions of its primary structure that distinguish it from other RNases and, hence, may account for its low ribonucleolytic activity and/or be critical to its unique angiogenic activity. One such region encompasses residues 58-70. Most notable in this sequence is the absence of the two cysteine residues that form a disulfide loop in RNase. A mutant angiogenin, ARH-I, was prepared in which residues 59-73 of RNase A were substituted for residues 58-70 of angiogenin (Harper and Vallee, 1989). It exhibited a markedly increased ribonucleolytic activity toward RNA substrates (300to 600-fold) and dinucleoside phosphates (20- to 200-fold) (Table I), but a substantially decreased or abolished angiogenic activity. This was the first indication that this region of angiogenin might be involved in its angiogenic activity, and showed that ribonucleolytic activity, although necessary, is not sufficient for angiogenic activity. The converse experiment has also been carried out in which residues 62-71 of angiogenin were inserted into RNase A in place of residues 63-74 (Allemann et aL, 1991; Raines et al., 1995). The resultant hybrid had less than 2% activity toward UpA relative to RNase, but this is still far more than that for angiogenin. Remarkably, its potency as an inhibitor of protein translation was somewhat better than that of RNase but less than that of angiogenin. Another region of angiogenin that differs substantially from RNase A includes the residues surrounding the active site lysine. Residues 38-42 of RNase A were substituted for residues 38-41 of angiogenin to give the hybrid ARH-II (Bond and Vallee, 1990) (Table I). As with ARH-
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I, it has greatly increased enzymatic activity (5- to 75-fold) and no significant angiogenic activity. What accounts for these changes in activity has not been established. Yet another mutant, ARH-IV, was prepared that incorporated the changes of both ARH-I and ARH-II (Harper et al., 1990). Not surprisingly, the enzymatic activity of this mutant was greater than that of either ARH-I or ARH-II. One further regional mutant was prepared in which the highly conserved segment of angiogenin-containing residues 8-21 was replaced by residues 7-21 of RNase A (Bond and Vallee, 1990). Its angiogenic potency was actually increased as was its interaction with RNase inhibitor. However, its enzymatic activities were essentially unchanged (Table I) except for its ability to inhibit cell-free protein synthesis, which, compared to angiogenin, was decreased about 20- to 30-fold. The structural basis for these changes is unclear, but the implication is that the unique Nterminal region of angiogenin plays some role in its biological activities.
V. R E L A T I O N S H I P OF RNase ACTIVITY A N D A N G I O G E N I C ACTIVITY
It required persistent effort to discern that angiogenin was a catalytically functional ribonuclease, but once this became apparent it was quickly shown that destruction of ribonucleolytic activity either by chemical modification or by treatment with RNase inhibitor also destroyed angiogenic activity (Shapiro et al., 1986; Shapiro and Vallee, 1987). Later, mutagenesis of Asp-ll6 was found to increase activity toward tRNA almost 20 times and to increase angiogenic activity by 1 to 2 orders of magnitude (Harper and Vallee, 1988b). Conversely, replacement of active site residues abolished both activities. It seemed clear that the angiogenic activity of angiogenin was critically dependent on its ribonucleolytic activity. But such activity per se was not sufficient for angiogenesis because no other member of the RNase superfamily had been found to induce blood vessel formation. Harper and Vallee (1989) reported that ARH-I has dramatically increased enzymatic activity (up to 600-fold) but is essentially devoid of CAM activity. This unique region of angiogenin obviously contributes importantly to the process of angiogenesis. Specifically, it appears to be part of a cell-surface receptor binding site that is distinct from the enzy-
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matic active site of angiogenin. Several pieces of evidence support this conclusion. Limited proteolysis, which played such an important role in the elucidation of the structure/function relationships of RNase A, showed that angiogenin was resistant to most proteases--most notably to subtilisin-but was quite sensitive to endoprotease Lys-C (Harper and Vallee, 1988a). This enzyme did not change its activity toward tRNA, however, despite cleavage of the Lys-60/Asn-61 bond. Later this product, called angiogenin K, was found to be inactive on the chick CAM (Hallahan et al., 1991). Another proteolytic product generated from angiogenin by an unidentified protease from baby hamster kidney cells had similar properties--essentially unchanged enzymatic activitity (Table I) and no CAM activity. In this case the bond cleaved was between Glu-67 and Asn-68 and the product was called angiogenin E. Even if both bonds were cleaved and residues 61-67 were removed the product was still enzymatically active. Clearly this region, which in RNase A constitutes part of the B2 binding site, is not essential to the ribonucleolytic activity of angiogenin but it appears to be very critical for induction of blood vessel formation. Mutagenesis has demonstrated that the active site residues His-13, Lys-40, and His-ll4 are all critical for the enzymatic activity of angiogenin, which, in turn, is essential for its angiogenic activity. Importantly, both H13A and H l l 4 A inhibit the angiogenic activity of native angiogenin, presumably by competing for its cellular receptor (Shapiro and Vallee, 1989). Although the catalytic sites are no longer functional, their cell-binding sites remain intact. In the case of angiogenin K or E, no such inhibitory effect was observed. Similarly, when this region was replaced by the corresponding region of RNase A, the product, ARHI, was devoid of angiogenic activity and did not compete on the CAM with the native protein (Hallahan et aL, 1991). It therefore seems likely that the role of this region in the mechanism of angiogenesis is to serve as part of the endothelial cell-binding site. The evolutionary process that converted this region into a cell-binding site may have simultaneously diminished its effectiveness as a B2 substrate-binding site. The specific residues constituting a B2 site in angiogenin have not been identified and, in fact, they are also not very well characterized for RNase A. Asn-71 is apparently an important component for all NpN' substrates of RNase A and Glu-111 is important when N' is guanosine, but Gin-69, previously suggested as important, is not (Tarragona-Fiol et al., 1993). Replacing Glu-111 by Gin reduces the
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activity of RNase A toward CpG by a factor of 8, whereas replacing the corresponding residue, Glu-108, in angiogenin by either Gln or Lys has a very much smaller effect (Curran et al., 1993a) (Table I). Whatever the BE site residues in angiogenin may be, the site is less selective than that in RNase. A comparison of angiogenin sequences from five different species identifies three residues in the region from 60 to 68 that are strictly conserved: Asn-61, Gly-62, and Pro-64. The Asn-Gly sequence is particularly susceptible to nucleophilic attack on the side chain amide carbonyl group, which results in deamidation. The desamido-61 product, obtained by incubating human angiogenin at pH 10, retains activity as a ribonuclease but is essentially inactive on the CAM and does not inhibit the CAM activity of native angiogenin (Hallahan et al., 1992). Alkaline treatment also deamidates Asn-109, which is in the only other conserved Asn-Gly sequence. The desamido-109 product can be separated from desamido-61 by HPLC and, surprisingly, it too is enzymatically active but angiogenically inactive. From the crystal structure of angiogenin, Ash-61 and Asn-109 are close to each other on adjacent loops that bear little structural similarity to their counterparts in RNase A (Acharya et al., 1994), and, hence, the cell-binding site may involve both loops. Deamidation gives rise to both aspartic and isoaspartic acids, with the latter form predominant. Mutagenesis was therefore employed to obtain the specific Asp-61 and Asp-109 angiogenins (Hallahan et al., 1992). Their activities were the same as those of the deamidated products (Table I) with the striking exception that they were able to inhibit the activity of native angiogenin on the CAM. This suggests that the ability to bind to the putative cell surface receptor plus having ribonucleolytic activity is not suffiicient and that Asn-61 and Asn-109 have some additional function, as yet undefined, that is required for angiogenesis. In this regard, Raines and co-workers (1995) have reported an extremely interesting result obtained by regional mutagenesis of RNase A. They substituted residues 58-70 of angiogenin for the analogous 15 residues of RNase, the converse of Harper and Vallee's studies of ARHI. This resulted in a 20-fold decrease in kcat/Km for cleavage of UpA, a small decrease in thermal stability (loss of one disulfide bond?), and, remarkably, the induction of angiogenic activity based on penetration of blood vessels into a special disk implanted into an adult mouse. Apparently, insertion of this recognition site into RNase A was sufficient to convert it into an angiogenic molecule (even though it lacked the counterpart of Asn-109). Moreover, they found that the peptide corre-
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sponding to angiogenin residues 58-70 inhibited neovascularization. It should be noted that the effects observed are smallm36.4% neovascularization in the control versus 45.2% in the hybrid, 18% inhibition by the peptide--but are nevertheless consistent with the role assigned to this region of angiogenin in the process of angiogenesis. On the other hand, additional properties of angiogenin that might also be important for blood vessel growth have not yet been examined with the hybrid.
Vl. INTERACTION WITH ENDOTHELIAL CELLS A. Binding to Endothelial Cells
Iodinated angiogenin binds to CPAE cells in a time- and concentrationdependent manner (Badet et al., 1989). Binding is reversible, saturable, and specific, is enhanced severalfold by copper, and is blocked by the RNase inhibitor from human placenta. Two types of binding sites have been identified. Approximately 200,000 molecules of angiogenin bind per cell to the first type with an apparent dissociation constant of 5 nM, and several million bind with a lower affinity of 200 nM to the second type. The latter are thought to be pericellular components. The highaffinity sites are characteristic of receptors in that binding triggers a series of intracellular events. No specific binding was seen with lung fibroblasts. Heath et al. (1989) showed that angiogenin binds to a variety of cell types, including fibroblasts, myocytes, mast cells, macrophages, and peripheral blood lymphocytes. Under the conditions used by Heath and co-workers, binding was not typical of classical ligand-receptor systems, because only relatively small amounts bound per cell. Moreover, experiments at 37 ~ and 4~ gave no evidence of internalization. These experiments were performed with confluent cells, whereas Badet et al. (1989) had examined cells during the first half of exponential growth and found that specific binding diminished markedly as cell density approached confluency. Chamoux et al. (1991) also studied the binding of angiogenin to endothelial cells and, under their conditons, binding was specific for bovine brain capillary endothelial cells. It did not occur with bovine aortic arch endothelial cells, smooth muscle cells, or fibroblasts; it was enhanced, but only slightly, by copper, it was diminished by RNase inhibitor, and it
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was abolished by treating the cells with trypsin. An apparent dissociation constant of 0.5 nM was reported and the number of molecules bound per cell was only 11,000. Again, this diminished markedly with increasing cell density. It should be noted that many of the results reported in this preliminary communication differ from those reported by others and are in need of additional clarification and confirmation.
B. Induction of Second Messengers
The interaction of angiogenin with CPAE cells stimulates a rapid but transient burst of diacylglycerol (DG) production, which reaches a maximum in 2.5 min and returns to base line by 10 min (Bicknell and Vallee, 1988). The peak of the dose response occurs at 1 ng/ml angiogenin, but this varies with cell type (0.1 ng/ml for BACE cells and 1/~g/ ml for H U V E cells). The appearance of DG is accompanied by a small and similarly transient increase in IP3 and IP2 followed by a more substantial and sustained increase in IP. There are also small temporary decreases in PIP2 and PIP, and a more significant decrease in PI. A small increase in lyso-PI was observed as well. These changes were interpreted to indicate activation of an inositol-specific phospholipase C that acts briefly on PIP2 and then on PI. The amount of IP3 produced is not sufficient to induce a burst of cystosolic calcium. The appearance of lysoPI suggests that phospholipase A2 is also activated, likely giving rise to arachidonic acid. In a subsequent study, Bicknell and Vallee (1989) found that angiogenin indeed stimulates endothelial cells to secrete prostacyclin, a product of arachidonate metabolism. The time course and concentration dependence were similar to those observed for DG but, in contrast, the effect was not seen with CPAE cells, only with H U V E and B A C E cells. In addition, the effect was inhibited by pertussis toxin whereas the D G response was not. A DG lipase inhibitor, indomethacin, quinacrine [a phospholipase A2 (PLA2) inhibitor], H7 (a protein kinase inhibitor), and an inhibitor of prostacyclin synthesis all blocked the stimulated secretion, as did down-regulation of C kinase by phorbol ester. The sensitivity to pertussis toxin suggests the involvement of a G protein in signal transduction to PLA2. The authors speculated that DG, released in response to angiogenin, activates C kinase, which phosphorylates lipocortins and thus abolishes their ability to inhibit PLA2. The inhibition
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of prostacyclin secretion by H7 and by phorbol myristic acid (PMA) argues for a role for C kinase in the angiogenin stimulatory effect. Smooth muscle cells respond to angiogenin much like endothelial cells (Moore and Riordan, 1990). There is a rapid activation of a phosphoinositide-specific phospholipase C (PLC), which generates DG and, in this case, IP3, but again no cytosolic calcium. After peaking at 30 sec, the D G level drops below that of the control, where it remains for at least 20 min, likely the consequence of hydrolysis by D G lipase. There is also rapid esterification of cholesterol by the fatty acids derived from phosphoinositide hydrolysis, but the mechanism underlying this increase is unknown. Angiogenin also appears to activate a G protein in smooth muscle cells that inhibits adenylate cyclase (Xiao et al., 1989). Thus, it induces a transient, concentration-dependent decrease in cyclic AMP that is prevented by pertussis toxin. The fact that angiogenin induces these responses points to one or more as yet unidentified cellsurface receptors, but the interrelationship between the various secondmessenger signals elicited is complex and thus far a connection to angiogenesis has not been established.
C. Effect on Endothelial Cell Growth
There is one report in the literature that describes a mitogenic effect of angiogenin on endothelial cells (Chamoux et al., 1991). Both bovine and human angiogenin were said to stimulate the proliferation of brain capillary but not aortic arch endothelial cells. A concentration of 100 ng/ml was about 80% as effective as 1 ng/ml bFGF. This same concentration of angiogenin also stimulated capillary endothelial cells arrested in Go to enter G~, as measured by the uptake of labeled thymidine. Although not yet confirmed, these results were believed to suggest that angiogenin is involved in endothelial homeostasis. Heath et al. (1989) examined the effect of angiogenin on quiescent bovine adrenal capillary endothelial cells and found that it did not induce D N A synthesis. This observation has often been cited as suggesting that angiogenin must be an indirect stimulator of angiogenesis, perhaps affecting endothelial cell functions that are important to blood vessel development. Indeed, Heath et al. found that although angiogenin was not a direct mitogen, it could modulate the response of endothelial, smooth muscle, and fibroblast cells to other mitogens. Surprisingly, it
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actually inhibits the stimulation of D N A synthesis in B A C E cells produced by conditioned medium from Im-9 lymphoblasts or 3T3-L1 adipocytes. In contrast, it enhances the proliferation of smooth muscle and fibroblast cells stimulated by conditioned medium from rat liver B R L 3A cells or RPMI 1788 cells, respectively. Modulation correlates with the in vitro synthesis of PIP2, but how this pertains to blood vessel formation is still unclear.
D. Angiogenin-Binding Proteins The induction of second messengers, the results of regional mutagenesis and limited proteolysis, competition by inactive mutants, and its specific binding to endothelial cells all point to an interaction between angiogenin and a cell-surface recept0r/binding protein. Initial attempts to identify such a molecule, like those in search of ribonucleolytic activity, were unsuccessful. Part of the problem may have had to do with the use of confluent cells and the particular cross-linking agents employed, but more important was the tendency of the binding protein to dissociate from the cell surface. Heparin sulfate was found to release from the plasma membrane of CPAE and GM 7373 cells a substance that bound to an angiogenin-Sepharose column and, when cross-linked to angiogenin with a water-soluble carbodiimide, gave a 58-kDa band on S D S PAGE (Hu et al., 1991). The binding protein was subsequently identified as a smooth muscle type of cz-actin (Hu et al., 1993) and its location on the cell surface was established by immunofluorescent staining (Moroianu et al., 1993). However, it has not yet been established whether actin is a functional receptor that is responsible for the observed cellular responses to angiogenin, nor is its known how actin binds to or dissociates from the cell surface. It is significant in this regard that antiactin antibodies inhibit the angiogenic activity of angiogenin (Hu et al., 1993). Actin, of course, binds to many proteins, and the biological significance of these interactions is not always understood. The effect of various inhibitors on the induction of second messengers implies that angiogenin may interact with a G-protein-coupled receptor but thus far direct evidence for any type of classical receptor has not been observed. Two novel ligands to the FGF receptor have been identified and cloned from a X e n o p u s laevis cDNA library based on their ability to increase tyrosine phosphorylation in cotransformed yeast cells
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(Kinoshita et al., 1995). Remarkably, one of these ligands is distantly related to angiogenin. Whether this points to the FGF receptor as an angiogenin-binding, signal-transducing protein in endothelial cells remains to be investigated. It may be pertinent that an 8-fold molar excess of bFGF completely inhibited the uptake of angiogenin by endothelial cells (Moroianu and Riordan, 1994a).
E. Nuclear Translocation of Angiogenin What is the fate of angiogenin once it binds to the endothelial cell surface? The dissociability of cell surface actin and its disappearance on incubating the cells with angiogenin suggested that some or all of the angiogenin-actin complex is released into the extracellular environment. Another possibility was that angiogenin blocks the antiactin antibody recognition site of the cell-bound complex or that the complex might be internalized into the cell, or both. Earlier studies with ~25I-labeled angiogenin at 4 ~ and 37~ had indicated that it was not internalized (Heath et al., 1989). However, immunofluorescence with an antihuman angiogenin monoclonal antibody identified an intracellular pathway for human angiogenin in growing CPAE cells (Moroianu and Riordan, 1994a). It undergoes endocytosis and is translocated from the cell surface to the nucleus, where it accumulates in the nucleoli. Whether all of the angiogenin that binds to the cell is internalized or whether some dissociates as an actin complex has not been established, although there is circumstantial evidence in support of the latter possibility. In addition, it is not known if the receptor to which angiogenin binds prior to internalization is indeed actin. A different receptor, as yet unidentified, may be required for this process. In this regard, internalization is not observed with confluent, nonproliferating cells even though actin can be detected on their surface. Of course, there may be a specific nuclear translocation mechanism that only operates in proliferating cells. This is especially significant in view of the fact that angiogenin is a circulating plasma protein and the vast majority of the vascular endothelial cells are in a nonproliferating, confluent state. Only when perturbed by trauma, disease, or some other change in local environment would these cells become responsive to angiogenin and serve as initiation sites for neovascularization.
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One implication of the nuclear localization of angiogenin is that the substrate for its ribonucleolytic activity may be within the nucleolus. In eukaryotic cells, the nucleolus is a specialized subcompartment wherein ribosome biogenesis occurs. Pre-rRNA is first transcribed from rDNA genes located in the nucleolus and then undergoes a series of processing modifications. At the same time the primary transcript undergoes sequential cleavages to generate mature forms of rRNA. The presence of angiogenin might enhance this pre-rRNA processing. Exogenous actin, an antiactin antibody, heparin, and heparinase treatment all inhibit the internalization of angiogenin, suggesting the involvement of cell surface actin and heparan sulfate proteoglycans in this process. Moreover, excess basic FGF, which is known to bind to surface heparan sulfates, completely inhibits angiogenin endocytosis. It has been established that angiogenin has a cell-surface-binding site that is essential for its angiogenin activity, but which can be modified without significant effect on its ribonucleolytic activity. CPAE cells do not internalize four enzymatically active angiogenin derivatives whose cell-binding site is modified, but they do internalize two enzymatically inactive mutants whose cell-binding site is intact. None of the mutants is angiogenically active. Proteins that are translocated to the nucleus of eukaryotic cells are targeted by specific nuclear localization signals (NLSs) that typically contain a high proportion of basic amino acids. The prototype NLS is the sequence PKKKRKY of the SV40 large-T antigen. Angiogenin contains a sequence, R3~RRGL, that has this function (Moroianu and Riordan, 1994b). Thus, the R33A mutant of angiogenin, in which Arg-33 is changed to Ala, is not translocated to the nucleus and lacks angiogenic activity. Moreover, a synthetic peptide, C G G R R R G L , when coupled to fluorescein isothiocyanate (FITC)-albumin, to an antihistone monoclonal antibody (Mab), to an antihuman nucleolus Mab, and to FITCR33A angiogenin by cross-linking with maleimidobenzoyl-N-hydroxysuccinimide, endowed all of them with the ability to be translocated to the nucleus of permeabilized cells (Moroianu and Riordan, 1994b). It is important to point out that other angiogenic factors also undergo nuclear translocation. Exogenous basic and acidic FGF, epidermal growth factor, and platelet-derived growth factor are endocytosed by target cells and translocated from the cell surface to the nucleus, where they accumulate in the nucleolus. This appears to be a general nuclear transport pathway that is a critical step in the process of angiogenesis (Moroianu and Riordan, 1994c). This view is supported by the observa-
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tion that endocytosis and nuclear translocation of angiogenin, bFGF, and aFGF are inhibited by two potent inhibitors of angiogenesis, protamine and platelet factor 4.
VII. A N G I O G E N I N E N H A N C E M E N T OF ACTIN A C C E L E R A T I O N OF PLASMINOGEN ACTIVATION
Actin accelerates the tPA-catalyzed activation of plasminogen (Lind and Smith, 1991a) but it is a noncompetitive inhibitor of plasmin (Lind and Smith, 1991b). Plasmin and tPA are involved in a series of normal and pathological processes of cell migration and invasion associated with wound healing, inflammation, and tumor cell metastasis. Hence it has been proposed that extracellular actin may play an important role in the angiogenic process in addition to its direct effects on fibrinolysis and thrombolysis (Lee and Galbraith, 1992). Angiogenin binds to actin and the complex is more effective than actin alone in stimulating tPA to produce plasmin (Hu and Riordan, 1993). Importantly, the angiogenin-actin complex does not inhibit plasmin and hence the overall proteolytic activity of a tPA-plasminogen mixture is enhanced ll-fold by the complex, whereas actin or angiogenin alone increase plasmin activity only about 2-fold. Binding of angiogenin to cell surface actin could therefore lead to activation of several protease cascades, such as the matrix metalloprotease and plasminogen activator/ plasmin serine protease systems. This could result in degradation of basement membrane and the extracellular matrix, thereby allowing endothelial cells to penetrate and migrate into the perivascular tissue, an essential feature of the angiogenesis process. Angiogenin indeed stimulates the cell-associated proteolytic activity of endothelial cells and promotes their invasion of fibrin gel and Matrigel basement membrane substrata (Hu et al., 1994). A transformed bovine aortic cell line GM 7373 is five times more invasive when grown in the presence of angiogenin than in its absence. The effect is inhibited by an antiangiogenin antibody and by c~2-antiplasmin. Angiogenin also induces a 14-fold increase in cell-associated proteolytic activity; as a consequence, the cells display an enhanced ability to digest an underlying fibrin gel matrix. In a separate study, bovine angiogenin was found to stimulate both urokinase mRNA and urokinase activity in bovine aortic endothelial cells (Jimi et al., 1995). Thus, angiogenin promotes several aspects
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of cell function that are important for endothelial migration and angiogenesis.
VIII. A N G I O G E N I N S U P P O R T OF E N D O T H E L I A L CELL A D H E S I O N
Trypsinized, washed CPAE cells are unable to adhere to an uncoated bateriological plastic surface but do adhere and spread when the surface is coated with angiogenin (Soncin, 1992). The effect seems to be specific because RNase A is much less effective in supporting such adhesion. Angiogenin has to be adsorbed to the plastic for the cells to adhere: cells added to uncoated plastic in an incubation solution containing angiogenin fail to do so. Human angiogenin is not as effective as bovine angiogenin for binding CPAE cells but is more effective for hamster fibroblasts. Because these fibroblasts lack high-affinity sites for angiogenin but do have low-affinity/high-capacity sites, it seems likely that the latter are responsible for cell adhesion. Binding is not mediated through integrins that can bind multiple ligands because fibronectin, laminin, and collagen, which interact with cell surfaces through such integrins, do not affect cell binding to angiogenin. However, Ca 2+ and/ or Mg 2+ are important for the CPAE cell-angiogenin interaction, much as they are in other cell-matrix molecule interactions. In addition, a synthetic peptide, RGDS, corresponding to the sequence essential for the interaction of extracellular matrix molecules with their cell receptors, inhibits CPAE cell adhesion to bovine angiogenin, which also contains this sequence. Other angiogenins, including the human type, do not have this sequence and yet RGDS still inhibits CPAE cell adhesion onto human angiogenin. Human colon adenocarcinoma HT-29 cells also adhere to angiogenincoated plastic and, indeed, they adhere faster than they do to extracellular matrix molecules (Soncin et al., 1994). In contrast to CPAE cells, binding does not require Ca 2+ and/or Mg 2+ and is not inhibited by RGDS. It is inhibited by heparin, however, and by treating the cells with heparinase or heparitinase, suggesting that human angiogenin recognizes a heparan sulfate type of proteoglycan on the surface of HT-29 cells. Inhibition of proteoglycan synthesis significantly reduces HT-29 cell binding to angiogenin, consistent with this view. It is worth emphasizing that angiogenin binds tightly to heparin-Sepharose and requires 0.8 M salt
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for elution. This affinity exceeds that of several growth and/or angiogenic factors and suggests that the activity of angiogenin is, in some way, regulated by heparin (Soncin et aL, 1994). Remarkably, heparin has no effect on the binding of CPAE cells to human angiogenin. Apparently, different classes of receptors are involved in binding, depending on cell type and, perhaps, species of angiogenin. In the case of endothelial cells, angiogenin serves as a substratum for cell adhesion by inducing an interaction similar to but separate from that of other extracellular molecules, which may be critical to angiogenesis. For tumor cells, it may provide a basis for cell adhesion during metastasis.
IX. MECHANISM OF ACTION
Angiogenin is an extracellular molecule that likely is synthesized mainly in liver (Weiner et al., 1987) and is released into the circulation. Because it is a potent inducer of neovascularization, details of its mode of action have been examined primarily with endothelial cells. For simplicity these details can be considered from three separate points of view, although as they are all interrelated. First, angiogenin is a ribonuclease and its enzymatic activity is essential for angiogenesis. It seems reasonable to conclude that the activity is operative within the cell, i.e., in the nucleolus, most likely in conjunction with an RNA substrate that is related to ribosome biogenesis. It could act enzymatically outside the cell, perhaps, if there was an extracellular RNA molecule available that could somehow transmit information. But blood plasma contains several other RNases with the same substrate specificity as angiogenin and much greater enzymatic activity, making it unlikely that a given RNA molecule would survive (at least in plasma) to serve as the target of angiogenin. It has been hypothesized (Benner, 1988) that extracellular RNA (presumably extravascular) in conjunction with RNases and RNase inhibitors assists in the development of tissues in higher organisms, but this remains a hypothesis. Second, the so-called cell-binding site of angiogenin is also critical for angiogenesis and this, together with the observed second-messenger responses, suggests that it (rather than some RNA hydrolysis product) must interact with its target cell in order to elicit its biological response. The competition observed with catalytic site-modified, binding-siteintact variants of angiogenin is in accord with this view. We have been
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unable to detect any cell surface R N A and, hence, this leaves intracellular R N A as the most probable substrate. In order to come into contact with this substrate the protein must get inside the cell. Heath et al. (1989) were unable to find evidence for internalization of 125I-labeled angiogenin into confluent cells, but it has since been shown by immunofluorescence that the protein is endocytosed specifically by subconfluent, proliferating CPAE cells (Moroianu and Riordan, 1994a). Most likely it enters the cell in an endocytotic vesicle, possibly as a receptor-ligand complex. There is no indication that angiogenin is released from the endocytotic vesicle into the cytosol. In fact, when it is directly microinjected into the cytosol it is cytotoxic by virtue of its inhibitory effect on protein synthesis (Saxena et al., 1992). This experiment was done with X e n o p u s laevis oocytes, which contain a cytosolic protein that inhibits frog RNase but is ineffective against mammalian RNases (Beintema et al., 1988). Most mammalian cells, presumably including endothelial cells, contain a cytosolic RNase inhibitor that is exceedingly potent toward mammalian RNases, including angiogenin. If endocytosed angiogenin were to enter the cytosol it would be either inhibited or cytotoxic. This suggests that the intravesicular angiogenin is targeted to a cellular organelle and, as summarized above, it is indeed translocated to the nucleus and accumulates in the nucleolus. The mechanism of nuclear translocation, the cellular machinery involved in vesicular transport to the nuclear membrane, the mode of entry through the nuclear pore, and the molecules required for nucleolar localization are as yet unknown. The fact that angiogenin has an essential NLS strongly supports the classic pore route. If so, it must be released from the endocytotic vesicle in the perinuclear region so that its NLS can be recognized by a specific NLS-binding protein. Targeting of the vesicle directly to the nuclear pores is possible, but would require a targeting signal typically contributed by the cytosolic domain of a putative membrane receptor, and this would not explain the need for a NLS in angiogenin. Vesicular fusion with the outer nuclear membrane is another possible route of entry, but there is no evidence, thus far, pointing to such a mechanism. Not enough information is available to allow speculation as to what angiogenin does within the nucleus that might ultimately result in angiogenesis. As noted, the enzymatic activity of angiogenin is orders of magnitude less than that of RNase A and yet it is essential for the protein to exercise its biological function. Moreover, even to have such low activity in vitro, angiogenin must undergo a conformational change to remove Gln-ll7
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from obstructing the B1 site. We do not know how much such a change would affect the catalytic activity of angiogenin, but if only Gln-ll7 were affected it seems unlikely that it would ever be as active as RNase A. The regional mutagenesis studies of Harper and Vallee (1989) and Raines et al. (1995) demonstrate that the replacement of the fourth disulfide loop/ B2 binding site sequence of RNase by a cell-binding site sequence substantially suppresses the enzymatic activity of angiogenin. These two regions have markedly different structures, however (Acharya et al., 1995), and it is possible that a conformational change in angiogenin could affect this part of the protein as well. Whether angiogenin requires full RNase-like activity is, of course, unknown, but one can imagine a scenario that keeps angiogenin in its inactive conformation until it reaches its functional target within the nucleolus. There ligand binding to its NLS or some other recognition site could trigger the appropriate structural changes and only then generate whatever activity is needed. There is also an extracellular role possible for angiogenin that involves binding to cell surface actin and heparan sulfate proteoglycans, but may not have anything to do with its ribonucleolytic activity. The angiogenin/ actin complex that is released from the cell surface may participate in the activation of tissue plasminogen activator. The subsequent generation of plasmin activity may be related to cell migration and capillary formation by activation of matrix-degrading enzymes. Angiogenin released from tumor cells may serve as a substratum for the adhesion of migrating endothelial cells and direct their growth toward the tumor. Conceivably it could also be a substratum for the tumor cells and help to promote metastasis. The overall mechanism of action of angiogenin as it relates to angiogenesis and as outlined here is summarized in Fig. 4.
X. BIOLOGY OF ANGIOGENIN A. Angiogenin and Wound Healing
The rabbit cornea, the chick CAM, and the hamster cheek pouch, among others, have all been used as loci to test for induction of angiogenesis, largely because they are reasonably accessible and allow ready assessment of new blood vessel formation. If new vessels could be induced to grow into and around an area of injury, this might enhance
14
481
Structure and Function of Angiogenin
ANGIOGENIN
(~)_
ANG-AngBP
_
AngBP
SECONDMESSENGER~ ~ SIGNALTRANSDUCTION ~) ~ ('~ ~:~ ~ ~ - ~ /
NUCLEUS I
ENDOCYTOSIS (~)/ / NUCLEAR TRANSLOCATION
(~
N ~
Fig. 4. Hypothetical scheme for the mechanism of action of angiogenin. (1) Angiogenin binds to its cellular receptor, indicated here as the angiogenin-binding protein (AngBP; actin), but which may involve heparan sulfate proteoglycans as well as a more conventional receptor. Binding occurs through the cell-binding region as indicated. This results in (2) activation of phospholipases C and A2, among others, with consequent secondmessenger signal transduction, the details of which have not been elucidated. In addition, angiogenin is endocytosed (3) and undergoes translocation to the nucleus (4). It then is taken up by the nucleus (5) and localized to the nucleolus (6), where in conjunction with nuclear "activation" signals initiated by the second messengers (7) it potentiates the endothelial cell for proliferation. Some of the angiogenin forms an extracellular complex with the angiogenin-binding protein (8), which assists in activating proteolytic cascades. Extracellular angiogenin also serves as a substratum that promotes cell adhesion and migration (9). The overall outcome of these processes is angiogenesis (10).
the process of healing. The blood supply to the menisci of the knee is quite sparse and it is well known that injuries in this location heal poorly. To test the potential of biochemically induced neovascularization as an aid to wound healing, a method was developed to examine the effect of angiogenin in the avascular cartilage of the rabbit knee (King and Vallee, 1991). The protein was implanted into experimentally injured menisci in the form of a methyl cellulose disk. Knees were then exposed and examined microscopically at different periods of time. Neovascularization was found in 3 of 8 knees at <4 weeks, in 30 of 53 at 4-10 weeks, and in 6 of 14 at >10 weeks for an overall 52% positive response. Only
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James F. Riordan
2 of 22 controls showed evidence of angiogenesis (9%). These initial studies show that, because vascularization is considered part of the normal wound healing response, angiogenin may provide a sustained stimulus and could prove useful to enhance healing of meniscal or similar injuries.
B. Antiangiogenin Antibody Suppression of Tumor Growth
The hypothesis that angiogenesis is essential for tumor growth (Folkman, 1971) led to the isolation of angiogenin from the conditioned medium of tumor cells grown in culture (Fett et al., 1985). Clearly, inhibition of angiogenesis would be an attractive approach to the treatment of both primary and metastatic cancer. Olson et al. (1994) showed that the growth of human colon adenocarcinoma (HT-29) cells in athymic mice is dependent on their production of angiogenin. A specific neutralizing monoclonal antibody to human angiogenin (Fett et al., 1994), in microgram doses, delays or prevents the appearance of HT-29 tumors in a statistically significant manner. By using a more sensitive model, Olson et al. (1995) were able to demonstrate a dramatic increase in the efficacy of this antibody. Up to 65% of treated animals remained tumor free for at least 30 days. A second neutralizing monoclonal antibody with a different epitope specificity was also effective in inhibiting tumor growth. Two other cell lines, a lung adenocarcinoma and a fibrosarcomamboth of which secrete angiogenin when grown in culture--were also sensitive to antibody treatment. In addition, actin, which also binds to angiogenin and inhibits its CAM activity, resembles the monoclonal antibodies by preventing the growth of tumors in 63% of treated animals. Antiangiogenin therapy most likely works by specific extracellular inactivation of tumor-derived angiogenin and inhibition of angiogenininduced neovascularization. Once tumor cells that escape therapy grow to palpable size, they are no longer susceptible to antibody administration. Nevertheless, the results clearly demonstrate a critical role for angiogenin in the early stages of tumor development, and indicate that inhibition of its function could be an effective means of therapy for malignant disease.
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XI. E P I L O G U E
It is now some ten years since the discovery of angiogenin and its recognition as a member of the ribonuclease superfamily. From the beginning it was evident that all the effort dedicated to the study of the pancreatic RNases would provide an exceptionally advantageous resource for expediting the elucidation of the structure-function relationships of angiogenin, and this has been borne out. As detailed above, much is now known about its structure, particularly in regard to its ribonucleolytic activity. But RNase A is not angiogenic and, as far as is known, has no other function than to degrade dietary RNA. Hence, it has been less useful in providing insight into the biological properties of angiogenin, which, as a consequence, has been slower to emerge. It remains one of the more potent inducers of neovascularization, and the inhibitory effect of antiangiogenin monoclonal antibodies on tumor growth clearly demonstrates that it plays a critical role in the early stages of this process, where it is known that blood vessel formation is essential. However, it is a normal constituent of the circulation which is contained in a vasculature that rarely undergoes proliferation. This implies a tightly regulated mechanism for its interaction with endothelial cells, perhaps via receptor modulation, which may, in fact, underlie the difficulties encountered thus far in identifying an angiogenin receptor. It also suggests that angiogenin may have some other (additional) nonpathological function and, indeed, recent reports provide initial evidence in this regard (Moenner et al., 1994; Tschesche et al., 1994; Matousek et al., 1995). A number of studies now underway, including targeted gene disruption and active site modeling, offer promise of adding important new information about the mechanism of action of angiogenin.
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Hu, G.-f., Chang, S.-I., Riordan, J. F., and Vallee, B. L. (1991). An angiogenin-binding protein from endothelial cells. Proc. Natl. Acad. Sci. U.S.A. 88, 2227-2231. Hu, G.-f., Strydom, D. J., Fett, J. W., Riordan, J. F., and Vallee, B. L., (1993). Actin is a binding protein for angiogenin. Proc. Natl. Acad. Sci. U.S.A. 90, 1217-1221. Hu, G.-f., Riordan, J. F., and Vallee, B. L. (1994). Angiogenin promotes invasiveness of cultured endothelial cells by stimulation of cell-associated proteolytic activities. Proc. Natl. Acad. Sci. U.S.A. 91, 12096-12100. Jimi, S.-i., Ito, K.-i., Kohno, K., Ono, M., Kuwano, M., Itagaki, Y., and Ishikawa, H. (1995). Modulation by bovine angiogenin of tubular morphogenesis and expression of plasminogen activator in bovine endothelial cells. Biochem. Biophys. Res. Commun. 211, 476-483. King, T. V., and Vallee, B. L. (1991). Neovascularization of the meniscus with angiogenin. J. Bone Joint Surg. 73-B, 587-590. Kinoshita, N., Minshull, J., and Kirschner, M. W. (1995). The identification of two novel ligands of the FGF receptor by a yeast screening method and their activity in Xenopus development. Cell (Cambridge, Mass.) 83, 621-630. Klagsbrun, M., and D'Amore, P. A. (1991). Regulators of angiogenesis. Annu. Rev. Physiol. 53, 217-239. Knighton, D., Ausprunk, D., Tapper, D., and Folkman, J. (1977). Avascular and vascular phases of tumor growth in the chick embryo. Br. J. Cancer 35, 347-356. Kobe, B., and Deisenhofer, J. (1993). Crystal structure of porcine ribonuclease inhibitor, a protein with leucine-rich repeats. Nature (London) 366, 751-756. Kobe, B., and Deisenhofer, J. (1995). A structural basis of the interactions between leucinerich repeats and protein ligands. Nature (London) 374, 183-186. Kurachi, K., Davie, E. W., Strydom, D. J., Riordan, J. F., and Vallee, B. L. (1985). Sequence of the cDNA and gene for angiogenin, a human angiogenesis factor. Biochemistry 24, 5494-5499. Kurachi, K., Rybak, S. M., Fett, J. W., Shapiro, R., Strydom, D. J., Olson, K. A., Riordan, J. F., Davie, E. W., and Vallee, B. L. (1988). Expression of human angiogenin in cultured baby hamster kidney cells. Biochemistry 27, 6557-6562. Langer, R., and Folkman, J. (1976). Polymers for the sustained release of proteins and other macromolecules. Nature (London) 263, 797-800. Lee, W. W., and Galbraith, R. M. (1992). The extracellular actin-scavenger system and actin toxicity. N. Engl. J. Med. 326, 1335-1341. Lee, F. S., and Vallee, B. L. (1990a). Modular mutagenesis of human placental ribonuclease inhibitor, a protein with leucine-rich repeats. Proc. Natl., Acad. Sci. U.S.A. 87, 18791883. Lee, F. S., and Vallee, B. L. (1990b). Kinetic characterization of two active mutants of placental ribonuclease inhibitor that lack internal repeats. Biochemistry 29, 6633-6638. Lee, F. S., and Vallee, B. L. (1993). Structure and action of mammalian ribonuclease (angiogenin) inhibitor. Prog. Nucleic Acid. Res. Mol. Biol. 44, 1-30. Lee, F. S., Shapiro, R. S., and Vallee, B. L. (1989). Tight-binding inhibition of angiogenin and ribonuclease A by placental ribonuclease inhibitor. Biochemistry 28, 225-230. Lin, M. C., Gutte, B., Moore, S., and Merrifield, R. B. (1970). Regeneration of activity by mixture of ribonuclease enzymatically degraded from the COOH terminal and a synthetic COOH-terminal tetradecapeptide. J. Biol. Chem. 245, 5169-5170. Lind, S. E., and Smith, C. J. (1991a). Actin accelerates plasmin generation by tissue plasminogen activator. J. Biol. Chem. 266, 17673-17678.
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15 Antitumor RNases R I C H A R D J. YOULE* A N D G I U S E P P E D ' A L E S S I O t *Biochemistry Section, Surgical Neurology Branch National Institute of Neurological Disorders and Stroke National Institutes of Health Bethesda, Maryland 20892 tDipartimento di Chimica Organica e Biologica Universith Degli Studi di Napoli Federico II 80134 Naples, Italy
I. Introduction II. Bovine Seminal RNase A. In Vitro and in Vivo Studies B. Antimetastatic Effect C. Structural and Functional Determinants of BS-RNase Antitumor Action D. Mechanism of BS-RNase Antitumor Action Ill. Amphibian RNases A. Onconase from the Leopard Frog Rana pipiens B. Lectins from the Bullfrog Rana catesbeiana and the Rice Paddy Frog Rana japonica IV. Concluding Remarks References
I. I N T R O D U C T I O N
Almost half a century ago, during the first great impetus in cancer research, the ready availability, "off the shelf," of a homogeneous enzyme protein, RNase A, already a useful model in many fields, induced RIBONUCLEASES: STRUCTURES AND FUNCTIONS
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numerous investigators to test the protein for anticancer activity. The concept of studying RNase A, the ribonuclease from bovine pancreas, for its potential as an antitumor agent was not without foundation, considering the importance that cellular RNAs were gaining during the same years as molecules essential for cell growth. However, although some of the results of these investigations appeared very promising, not all data unequivocally pointed in the same direction, as a review by J. S. Roth (1963) concluded years later. Slowly, RNase A ceased to be tested as an antitumor drug (after all, it is not an antitumor drug), and another RNase from fungi, a-sarcin, so named for its potent antisarcoma action (Olson et al., 1965), also disappointed the experimental oncologist, because it was found to be toxic to all animal cells, tumoral and nontumoral [see Wool (1984) for a review]. Somewhat coincidentally, the idea that an RNase could be an antitumor drug was regarded as dubious, although a strong correlation was found between RNase activation in tumor tissue and tumor regression (Daoust and DeLamirande, 1975). Today, the realization that some RNases have antitumor activity may not be so surprising. Furthermore, current knowledge of the antitumor RNases may encourage speculation on what might have been the basis for the early peculiar findings on the antitumor action of RNase A. One possibility is that some preparations may have contained a substantial dimeric fraction of the protein, possibly produced through repeated lyophilizations, we know now that natural or engineered dimeric RNases do have antitumor action, as reviewed herein. Furthermore, as is discussed later, once artificially inoculated into a cell, RNase A becomes a powerful cytotoxin; hence it may be surmised that in some earlier investigations, in some cell culture conditions, the RNase was internalized in such a way that affected the survival of tumor cells.
II. B O V I N E S E M I N A L RNase
A. In Vitro a n d in Vivo S t u d i e s
The first observation of the antitumor action of bovine seminal RNase (BS-RNase) was reported by J. Matousek (1973). He found that an
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aspermatogenic protein with RNase activity, isolated from bull semen, induced degenerative effects on Crocker tumors implanted in mice, reducing the weight of the tumors. A high mortality was observed in the treated animals, a finding confirmed in a successive report (Stanek and Matousek, 1976) describing the BS-RNase treatment of rats bearing Walker carcinosarcoma. In this case adverse effects were not detected if the treatment was started on animals bearing small tumors. It should be mentioned that the enzyme preparations used in these studies were not homogeneous. After these first in vivo studies, all the investigations that followed, for a long period, were on tumor cells in vitro (Matousek and Stanek, 1977; Cinatl et al., 1977; Matousek et al., 1979; Vescia and Tramontano, 1981; Vescia et al., 1980). Table I summarizes the malignant cell types used in these and other more recent investigations: in all cases, tumor cells have been found to be susceptible to the toxic action of the protein. Nonmalignant cells are affected by the protein only slightly or are totally
TABLE ! Malignant Human and Rodent Cells Susceptible to Selective Cytotoxic Action by BS-RNase
Cell line BP-8 EL-4 HeLa BHKPY BHKT6Tr5 3T3SV 3T3Py 41 A3 C6 TK-6 MPTK-6 3LL CHO SVT2 9L
Origin
Animal source
Leukemic cells Leukemic cells Malignant cells Polyoma-transformed fibroblasts Polyoma-transformed fibroblasts SV40-transformed fibroblasts Polyoma-transformed fibroblasts Neuroblastoma-derived cells Glioma-derived cells Oncogene-transformed thyroid follicular cells Lung metastasis-derived cells (from TK-6 tumor) Lung carcinoma Malignant cells SV40-transformed fibroblasts Glioma cells
Mouse Mouse Human Hamster Hamster Mouse Mouse Mouse Rat Rat Rat Mouse Hamster Mouse Rat
Ref."
a K e y to r e f e r e n c e s : 1, Matousek and Stanek (1977); 2, Cinatl et al. (1977); 3, Vescia et al. (1980); 4, Vescia and Tramontano (1981); 5, Laccetti et al. (1992); 6, Laccetti et al. (1994); 7, Mastronicola et al. (1995); 8, Wu et al. (1995).
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resistant, especially when they do not derive from stabilized lines in culture, but from primary cultures at early passages (Vescia et al., 1980). The experimental dual system made up of a stabilized cell line and of its malignant counterpart (Vescia et al., 1980), generally obtained by viral transformation, has been especially useful in defining the antitumor action of BS-RNase, because it contained a built-in control of the selectivity of the toxic action of the protein toward malignant cells, hence of its actual antitumor action. The in vitro effect of BS-RNase is cytotoxic rather than cytostatic, and is exerted whether the protein is added to substrate-attached, proliferating cells or to cells prior to their adhesion to the substrate (Laccetti et aL, 1992; Vescia et al., 1980). Further in vivo studies have been reported. These were carried out on carcinomas induced in rats by subcutaneous injection of cells derived from a thyroid follicular carcinoma (Laccetti et al., 1992), and on tumors induced in mice inoculated intramuscularly with Lewis lung carcinoma cells (Laccetti et al., 1994). In both series of experiments a dramatic inhibition of tumor growth was observed: in the former series, the protein was administered locally in the peritumoral area, whereas in the latter, administration was by intraperitoneal injections, thus the protein reached the tumor through a systemic route. Interestingly, in neither case, no adverse effects of the protein on animal survival were detected. In the experiments with rats inoculated with carcinoma cells, the animals were treated during 14 days with a total dosage of 14 mg of protein per animal, and after 30 days no deaths were registered. In both sets of experiments, no significant toxic effects were detected in the treated animals; in fact, the tumor-bearing mice inoculated with Lewis lung carcinoma cells were cured of the characteristic anemia induced by the tumor condition (Laccetti et al., 1994).
B. Antimetastatic Effect
When the in vitro cytotoxic effect of BS-RNase on cells derived from a primary tumor was compared to the effect on cells of the same histotype, but derived from a metastatic tumor, it appeared that the antitumor protein was much more effective on the latter, more malignant, cell type (Laccetti et al., 1992, 1994). These in vitro data were confirmed by the in vivo experiments carried out by measuring the number of metastases
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produced in the lungs of mice inoculated intramuscularly with Lewis lung carcinoma cells, and then treated intraperitoneally with BS-RNase (Laccetti et al., 1994). Very few and very small metastatic foci were detected in the lungs of the treated animals (see Fig. 1), and an apparent preferential action of the protein on metastatic cells, rather than on the primary tumor, was observed. When the treatment was started 24 hr after implantation of the tumor, and consisted of the administration of 20/zg of protein per gram of animal weight, tumor growth was inhibited by about 66%, whereas the reduction of the number of metastases in the lungs was more than 90%. This strong antimetastatic effect may be considered of particular interest for potential therapeutic applications of BS-RNase as an antitumor drug, considering that for some cell systems conventional antineoplatic drugs have greater cytotoxic effects on nonmalignant or on less malignant cells (Spalletti-Cernia et al., 1995).
C. Structural and Functional Determinants of BS-RNase Antitumor Action
As has been found for other ribonucleases endowed with special (noncatalytic) bioactions (RISBases; see Chapter 12) (D'Alessio, 1993), and in particular for onconase, another antitumor RNase (see Section Ill,A), BS-RNase loses its antitumor activity when deprived of its catalytic activity. This has been determined by testing the antitumor action of the enzyme following inactivation either by chemical modification (Vescia et aL, 1980) or by replacement via site-directed mutagenesis (Kim et al., 1995), of one of the catalytically essential His residues. The data collected on the degradation of cellular RNA, which occurs in BS-RNase-treated tumor cells (see Section II,D), are in line with these findings. However, it cannot be excluded that elimination of an active site His residue might not only alter the chemistry of the active site, but might also affect the global structure of the protein, which might thus lose its capacity of interaction with other cellular structures, on which its antitumor action depends. As for the structural determinants underlying the antitumor action of BS-RNase, the early finding that the dimeric structure of the protein is an essential prerequisite for its antitumor action (Vescia et al., 1980) has been confirmed (Kim et al., 1995): monomerized BS-RNase has no antitumor activity. This has been correlated to the finding of Bartholeyns
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and Baudhuin (1976), that dimers of RNase A obtained by cross-linking the monomeric protein with bifunctional reagents are antitumor agents. Although there is a striking correlation between the loss of antitumor activity on monomerization of naturally dimeric BS-RNase, and the production of this activity in RNase A on dimerization, there are significant differences between the biological actions as displayed by RNase A dimers and by BS-RNase. The effect of the RNase A dimers is cytostatic rather than cytotoxic, and is not lost on inactivation of the dimerized enzyme by chemical modification of the active site His residues. Furthermore, dimeric RNase A was found to provoke labilization of the lysosomes (Bartholeyns and Baudhuin, 1976), whereas BS-RNase does not have this effect (A. Russo and G. D'Allesio, unpublished). It should be stressed that preparations of dimeric RNase A used in these investigations were heterogeneous mixtures of dimers with cross-links across different lysine residues, with some monosubstituted lysines. A homogeneous preparation of dimeric RNase A with antitumor action has been obtained through site-directed mutagenesis of the cDNA coding for the protein, followed by expression in E s c h e r i c h i a coli (Di Donato et al., 1994). The follow-up on this observation has confirmed the lack of an actual relationship between the antitumor action of crosslinked dimers of RNase A, obtained by reaction with bifunctional reagents, and that of BS-RNase. Dimeric mutants of RNase A, stabilized through disulfide cross-links mimicking the intersubunit disulfides of BS-RNase, display only a modest antitumor effect, but this is strongly enhanced when, by insertion of other mutations into the RNase A sequence, the dimeric mutants acquire the property of exchanging the Nterminal segments between subunits (Di Donato et al., 1995). In fact, the intensity of the antitumor action in a series of dimeric RNase A mutants satisfactorily correlates with the extent of exchange of the Nterminal domains between protomers. Likewise, isolated preparations of the quaternary form of BS-RNase characterized by N-terminal exchange have a greater antitumor action than do preparations of the form with no exchange (Cafaro et al., 1995). Furthermore, BS-RNase mutants with minimal N-terminal exchange exhibit a low-level antitumoral action (Kim et al., 1995). Hence, the dimeric structure of BS-RNase as such, although essential, is not a sufficient structural basis for its antitumor action: it is important, and possibly essential, that in the dimer the two subunits exchange their N-terminal domains.
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Under conditions that affect the integrity of the Golgi apparatus, BSRNase monomers also exhibit antitumor properties (Wu et al., 1995). This surprising result is discussed in Section II,D. Recombinant BS-RNase expressed in heterologous prokaryotic systems that do not provide for the removal of the N-terminal Met exhibits less antitumor action than does the mature protein. Full activity can be restored on removal of the extra Met residue (Adinolfi et al., 1995).
D. Mechanism of BS-RNase Antitumor Action
The selective toxicity of BS-RNase toward malignant cells can be due (1) to a selective recognition and uptake of the protein by malignant cells and/or (2) to a differential fate of the internalized protein in the tumor cells. Early indications that BS-RNase is internalized by both normal and malignant cells, and to about the same extent (Vescia et al., 1980), were later confirmed (Mastronicola et al., 1995): labeled BS-RNase binds with the same KD (~1 • 10 -7 M) to both cell types. However, the protein-binding sites are on the extracellular matrix surrounding the cells, and no membrane-bound receptor-like sites have been detected. The matrix binding can be correlated with the antitumor action, because (1) homologous RNases and BS-RNase monomers, which are inactive as antitumor agents, do not displace bound BS-RNase, and (2) Chinese hamster ovary (CHO) cells grown in suspension, with little pericellular matrix, bind very little BS-RNase and are resistant to the toxic action of BS-RNase, whereas when grown as monolayers the cells bind the protein and are sensitive to its toxic action. In the same study (Mastronicola et al., 1995), internalization of the protein by nonmalignant cells was found to have no apparent effects on the cell machinery, whereas in malignant cells BS-RNase was found to degrade ribosomal RNA and inhibit protein synthesis. These data suggest that the basis for the selective toxicity of BS-RNase for tumor cells must be explored inside the cell. A realistic hypothesis is that in normal cells BS-RNase is routed through a "nontoxic" pathway that does not lead to the cytosol, whereas in tumor cells a "toxic" pathway is open, conveying the protein to the cytosol, where it degrades ribosomal RNA. The loss of rRNA would in turn undermine protein biosynthesis and provoke the death of the cell.
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This hypothesis is supported also by data collected from two distinct experimental approaches. The first, based on the notion that tumor cells have a higher net negative charge compared to normal cells (Becker, 1975), led to the finding (Mancheno et al., 1994) that BS-RNase can alter and permeate artificial membranes built with negatively charged phospholipids. This effect is not shared by monomeric BS-RNase, which is inactive as an antitumor agent. The absence of a direct interaction of BS-RNase with the plasma membrane suggests that intracellular membranes may be affected by the protein. The other approach was based on the use of agents that disrupt the Golgi complex, such as retinoic acid and monensin. In their presence, the cytotoxic effect of BS-RNase was greatly enhanced, with an increased degradation of ribosomal RNA (Wu et al., 1995). Furthermore, in the presence of retinoic acid, human pancreatic RNase, angiogenin, and monomeric BSoRNase became cytotoxic for tumor cells. These data suggest that retinoic acid disrupts the nontoxic intracellular pathway that is ordinarily used by nontoxic RNases, giving these proteins access to the cytosol, where they degrade ribosomal RNA and affect cell survival. The increased cytotoxicity of BS-RNase may be explained by allowing that, even in tumor cells, an intact Golgi apparatus routes a fraction of the BS-RNase through the nontoxic pathway; once the Golgi is disordered, however, even that fraction of protein reaches the cytosol, thus increasing the level of cytosolic RNase activity. However, the cytotoxicity of nontoxic RNases in the cytosol in the presence of retinoic acid is still surprising, as is their toxicity when they are artificially inoculated into a cell (Rybak et al., 1991; Saxena et al., 1991). It would be expected that when these RNases reach the cytosol, they are complexed and inactivated by the cytosolic protein RNase inhibitor (RI). One possibility is that tumor cells have low levels of RI. Although many studies have been carried out to investigate possible differential levels of RI in tumor versus nontumor cells, at present it is difficult to reach a conclusion, because both low and increased levels of inhibitor have been reported for tumor cells (see Lee and Vallee, 1993). In its native dimeric state BS-RNase is resistant to RNase inhibitor (Murthy and Sirdeshmukh, 1992), whereas in its monomeric form it is readily bound and inactivated. However, BS-RNase resistance to RI may not explain the RNase activity in the cytosol of internalized tumor cells. In the reducing cytosolic environment, on cleavage of the readily reduced intersubunit disulfides, about one-third of the RNase would be monomerized, hence easily inactivated by RI. The remaining two-thirds,
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derived from the protein quaternary form in which the subunits exchange their N-terminal segments, would still be dimeric, as noncovalent dimers, and would remain resistant to RI (see Chapter 19). Recent data (Murthy et al., 1996) have shown that under mild reducing conditions free dissociated monomers and noncovalent dimers are in equilibrium, and that in the presence of RI the noncovalent dimers of BS-RNase monomerize and are removed from the equilibrium. This should render all the BSRNase that reaches the cytosol susceptible to the neutralizing action of the inhibitor, hence unable to exert antitumor action. Because this does not happen, one may surmise that there is a low level of inhibitor in the tumor cells. Alternatively, in both normal and tumor cells, cytosolic RI levels might be inadequate to neutralize an influx of external RNase molecules. This would once again stress the importance of the existence of a toxic pathway in tumor cells, to explain the selectivity of BS-RNase antitumor action. Either conclusion would support results of an experiment (Saxena, 1991, unpublished) in which noncytotoxic mammalian RNases and an amphibian RNase, such as onconase, were inoculated into Xenopus oocytes. The Xenopus cells contain the native amphibian RNase inhibitor, which is inactive on mammalian RNases but presumably active on amphibian RNases (Kraft and Shortman, 1970; Roth, 1962). The experiment showed that onconase was only moderately active as a cytotoxin, but the mammalian RNases were over 100 times more toxic than onconase. One interpretation is that the amphibian inhibitor did act on onconase, but its levels were inadequate to neutralize the enzyme completely; on the other hand, the inhibitor was totally inert toward the mammalian enzymes.
III. A M P H I B I A N R N a s e s
Two independent approaches led to the discovery that frog RNases have antitumor activity. In one case an RNase was identified from studies of differentiation factors and in another case through research on lectins, both with a goal of developing cancer therapeutics. Testing the idea that, during development, factors and forces that control cell growth can be used to differentiate tumor cells and thus block cancer development revealed that extracts from frog embryos inhibit the growth of a frog tumor (K. Shogen, 1993, personal communication). Proceeding quickly into human trials, a trademarked extract of
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Richard J. Youle and Giuseppe D'Alessio
R a n a p i p i e n s embryos, Pannon, was administered to cancer patients. Encouraging results from these clinical trials led to the purification of an active component from the frog embryo extract (Ardelt et al., 1991); it was initially named P-30 and is now called onconase. Sequencing of onconase showed it to be homologous to RNase A. Simultaneously, lectins from a variety of sources were found to agglutinate cells via an activity based on specific sugar recognition sites. In a few cases, cancer cells were selectively agglutinated, and the lectins in these cases became a tool to study changes in surface glycoprotein and glycolipid composition occurring in malignancy. This work suggested that the differences in surface carbohydrate could be used for therapy by using lectins to bind cells and inhibit cell growth or to target toxic proteins to these cells. Lectins are found widely in nature in both plants and animals and in exotic species ranging from mushrooms to sponges. Several frog tissues have been found to contain lectins that selectively agglutinate tumor cells (Sakakibara et al., 1976). These lectins had anticancer activity in animal models and on purification and sequencing were found to be members of the RNase A superfamily (Kawauchi et al., 1975; Nitta et al., 1993).
A. Onconase from the Leopard Frog Rana pipiens 1. In Vitro Studies Onconase has been studied most thoroughly in tissue culture to probe its mechanism of action. When onconase is incubated with tumor cells at concentrations up to 1 ~g/ml, cell growth is slowed markedly. Cells stop dividing in the G~ phase of mitosis (Darzynkiewicz et al., 1988). Onconase inhibits cell proliferation at low doses (Darzynkiewicz et al., 1988) and actually kills cells at higher doses (10-100/xg/ml, or around 10-6-10 -5 M) (Wu et al., 1993). However, different cell lines have different levels of sensitivity to the protein. Interestingly, onconase toxicity for cells in culture synergizes with a number of other cytotoxic and chemotherapeutic drugs, including tamoxifen, an antiestrogen; trifluoroperazine, a calmodulin inhibitor; lovostatin, a cholesterol synthesis inhibitor; and retinoic acid (Mikulski et al., 1990a, 1992; Wu et al., 1995). How does this frog ribonuclease inhibit cell growth? A series of steps have been proposed modeled on how protein toxins such as ricin and
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diphtheria toxin kill cells (Youle et al., 1993). First, there is evidence that onconase binds the cell surface. Direct binding analysis showed that indeed onconase binds cells with affinities of 6 • 10 -8 to 2.5 • 10 -7 M. This saturable binding of onconase to cells appears to be the first step in the cytotoxic pathway and, as determined by preincubation and washing experiments, appears to be a mandatory step in the toxicity pathway (Wu et al., 1993). The receptor is unknown. Sialic acid and other simple sugars do not competitively block toxicity and neuraminidase treatment of cells also failed to block toxicity (Y. N. Wu, 1993, unpublished data), in contrast to results with sialic acid-binding lectins discussed in Section III,B (Nitta et al., 1994a). After binding the cell surface, onconase is thought to reach the cytosol, where RNA, the likely substrate, is found. Endocytosis of onconase may be a necessary step because ATP is required for toxicity (Wu et al., 1993). Internalization into endosomes does not deliver proteins to the cytosol, and it is not clear how the hydrophilic RNase crosses the lipid membrane surrounding the cytosol. However, disruption of the Golgi apparatus with monensin and retinoic acid increases the toxicity of onconase (Wu et aL, 1995), likely by allowing more of the onconase into the cytosol (Wu et al., 1994). Ammonium chloride, a lysosomotropic drug with some properties similar to those of monensin, had no effect on onconase (Wu et aL, 1993) yet blocked the toxicity of the sialic acidbinding lectin (Nitta et aL, 1994b). Onconase expresses ribonuclease activity, albeit orders of magnitude less than is found for RNase A. With yeast RNA as a substrate, the relative kcat of onconase is 0.1% that of bovine RNase A (Boix et al., 1996). The substrate specificity, optimal pH, and optimal salt concentration of onconase all differ from those of RNase A. The optimal dinucleotide substrate for onconase is UpG, reflecting the uridine preference for the B~ site and the guanosine specificity for the B2 site (Ardelt et aL, 1994; Boix et al., 1996). Kinetic and structural studies on RNase A identified Thr-45, Phe-120, and Ser-123 as the main residues of the B~ site and loop 65-72 for the B2 site (Richards and Wyckoff, 1971). Thr45 and Ser-123, by interactions with two conserved water molecules, provide a dynamic mechanism for uridine or cytidine accommodation in B~ (Gilliland et aL, 1994). Thr-45 is conserved in all members of the RNase family identified to date, but Ser-123 is deleted in the frog members of this family. The 65-72 loop in RNase A identified for B2 site specificity is deleted in the frog RNases (Fig. 2). A computer model of onconase with d(UpG) (S. Mosimann, 1995, unpublished data) implicates
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Richard J. Youle and Giuseppe D'Alessio
Fig. 2. Sequence homology among frog RNases and bovine RNase A. The amino acid numbering is for bovine RNase A.
Lys-33 in uridine B1 specificity, and Ser-54, Asn-56, and Glu-91 in B2 binding. As discussed later, the sialic acid-binding lectins have the same preference for UpG and have similar amino acids at these positions. RNase enzyme activity is required for onconase toxicity in tissue culture (Wu et al., 1993) and in animals (Newton et al., 1994a), as is seen for BS-RNase (see Section II,C). Direct examination of cellular R N A shows that rRNA within intact cells treated with onconase can be degraded in certain situations. However, tRNA (Boix et al., 1996; Lin et al., 1994) and viral RNA (Saxena et al., 1996) are more sensitive to onconase than are several mRNAs or rRNA. 2. In Vivo Studies
In an animal model of cancer, M109 carcinoma cells were injected intraperitoneally and onconase was given to the mice intraperitoneally by various schedules, starting 1 day after the tumor was injected (Mikulski et al., 1990b). The more frequently the onconase was administered, up to
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daily injections of 12 tzg/mouse, the more toxic onconase was to the mice. Weekly injections of 40/xg/mouse gave the best therapy, with 30% long-term survivors, whereas saline-treated control animals all died from the tumor. Results of phase 1 clinical trials have been published (Mikulski et al., 1993). Dose-limiting toxicity appears to be kidney damage manifested as proteinuria and edema. Tumor regression was seen infrequently, but tumor cell growth is reported to be slowed and patient survival may be enhanced. A clinical trial of onconase combined with tamoxifen has been performed. A phase 3 clinical trial planned to study pancreatic cancer may ascertain the degree of efficacy onconase exhibits. The schedule of drug administration appears important for animal toxicity (Mikulski et al., 1990b) and likely for human toxicity and tumor cell killing as well. Onconase is rapidly cleared from mouse plasma and accumulates in the kidney (Vasandani et al., 1996), consistent with the type of toxicity seen in the clinic. 3. Structural Determinants
Onconase has a molecular mass of 11,816 kDa and is highly basic (Ardelt et al., 1991). The protein is homologous to members of the RNase A superfamily (Ardelt et al., 1991), with 28% identity to bovine RNase A (Fig. 2). The crystal structure of onconase has been solved (Mosimann et al., 1994). Most of the residues involved in the active center of RNase A (Gin-11, His-12, His-119, Lys-41, Phe-120, and Thr45) have homologous counterparts in onconase: Lys-9, His-10, His-97, Lys-31, Phe-98, and Thr-35. One major difference between the active sites of RNase A and onconase is the N-terminal residue. In RNase A Lys-1 extends far out of the active site whereas the pyroglutamyl residue found at the N terminus of onconase folds back against the N-terminal a helix and forms a hydrogen bond with Val-96 in the C-terminal/3 sheet. Pyr-1 of onconase also hydrogen bonds to Lys-9, which simultaneously interacts with the main phosphate of the substrate (Mosimann et al., 1994). Studies of recombinant onconase show that the N-terminal pyroglutamyl residue is one key to both catalysis and toxicity. Expression of an onconase gene in bacteria with an additional methionine at the N terminus was accomplished, as for eosinophil-derived neutrotoxin (EDN) (Newton et al., 1994b) and human pancreatic RNase (Wu et al., 1995). Whereas the latter two RNases were fully active, onconase was not.
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Therefore another recombinant form of onconase was produced that allowed cleavage of the N-terminal methionine with cyanogen bromide. On removal of the N-terminal methionine and cyclization of the Gln-1 to pyroglutamic acid, the enzyme activity and toxicity of onconase were reconstituted (Boix et al., 1996). It is interesting that all four known forms of frog RNases have a conserved Pyr-1 in the same position at the N terminus (Fig. 2). Another feature proposed to account for the toxicity of onconase is its resistance to the mammalian ribonuclease inhibitor (Wu et al., 1993). The Ki between RI and onconase is 107 times or more higher than that between RI and RNase A (Boix et al., 1996). This brings up an intriguing connection with BS-RNase. As discussed previously, BS-RNase in its dimeric form is RI insensitive (Murthy and Sirdeshmukh, 1992). One additional mystery is how these antitumor RNases differ functionally from RNase A to account for the dramatic difference in toxicity. For example, injection of RNase A into the CSF of animals showed it to be more than 5000 times less toxic than either onconase or E D N (Newton et al., 1994a; Sorrentino et al., 1992). However, there are no obvious determinants common to E D N and onconase in either the amino acid sequence (Youle et al., 1993) or in the crystal structure (Mosimann, 1996), thus no accounting for their common toxicity. In tissue culture, RNase A and E D N were not toxic to cells up to 10 -5 M, whereas onconase becomes toxic above 10 -7 M (Newton et al., 1994b).
B. Lectins from the Bullfrog Rana catesbeiana and the Rice Paddy Frog Rana japonica 1. Rana catesbeiana Lectin A sialic acid-binding lectin has been purified from the eggs of the bullfrog, R a n a catesbeiana (Titani et al., 1987). It is highly basic and has a molecular mass around 13,000 Da. It may bind to negatively charged molecules generally and it remains unclear as to how specifically it binds sialic acid, because sialic acid alone does not block cell agglutination. The sialic acid specificity of the lectin was deduced based on the observation that complex glycolipids and glycoproteins blocked agglutination and sialidase treatment of the glycoproteins inactivated them.
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The sialic acid-binding lectin from R. catesbeiana was sequenced (Titani et al., 1987) and later found to be homologous to members of the RNase A superfamily (Lewis et al., 1989). It has three of the four disulfide bonds conserved with RNase A (Fig. 2). There is an additional disulfide bond at the extreme C terminus, homologous to a disulfide bond in onconase (Fig. 2). Active site residues involved in catalysis of RNase A, His-12, His-ll9, and Lys-41, are conserved and the sialic acid lectin expresses RNase activity (Nitta et al., 1993) as predicted (Lewis et al., 1989). Independently, a very similar, if not identical, RNase was isolated from R. catesbeiana eggs (Liao, 1992). Studying transcription by microinjection into X e n o p u s laevis and R. catesbeiana eggs, Liao found that transcription was inhibited only in the Rana eggs due to the presence of large amounts of a ribonuclease. Partial sequence analysis of the purified RNase showed 20 of 20 amino acids sequenced identical to residues 59-70 of the sialic acid-binding lectin from the same tissue source and species. Studying the substrate specificity of this RNase showed differences from RNase A and similarities among many of the frog RNases. R a n a catesbeiana egg RNase preferred CpG and UpG substrates and was very similar, but not identical in dinucleotide specificity, to egg RNases of R. catesbeiana reported by Okabe et al. (1991). The lack of an abundant RNase in X e n o p u s eggs may explain why it is such a good system for studying transcription and a relevant model to study mammalian RNase cytotoxicity (Rybak et al., 1991; Saxena et al., 1992). Interestingly, an inhibitor of the R. catesbeiana RNase found in the frog egg extract was sensitive to sulfhydryl modification. The R. catesbeiana egg RNase (Liao, 1992), the sialic acid-binding lectin (Nitta et al., 1993), and onconase (Wu et al., 1993) are also insensitive to the mammalian RI, as noted long ago by Roth (1962). Having now determined the crystal structure of mammalian RI (Kobe and Deisenhofer, 1993) and of the RNase A - R I complex (Kobe and Deisenhofer, 1995), it will be interesting to characterize the differences between mammalian RNase-RI interactions and the frog R N a s e - R I interactions. To help understand the physiological function of such a highabundance protein in eggs, the localization of the RNase within R. catesbeiana was investigated (Liao and Wang, 1994). Examining many adult frog tissues revealed high amounts of the egg RNase only in the ovary, the site of the eggs. The only other detectable RNase was found in liver and kidney but appeared not to be identical to the egg form; possibly this was accounted for by the known R. catesbeiana liver enzyme (Nagano
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et aL, 1976). Looking subcellularly within the R. catesbeiana egg revealed
that the RNase was sequestered in protein-rich yolk granules. The organelles store the protein needed by the growing embryo and contain a crystalline core of protein surrounded by a matrix of soluble protein. The RNase resides within this soluble matrix. Thus the protein is not cytosolic and inhibition of the obvious deleterious activity of such a highabundance RNase is not such a problem. In analogy to the plant toxin ricin, also sequestered in seed protein-rich storage granules surrounding a protein crystalline core, the RNase may serve as a defense toxin to protect the eggs. 2. R a n a japonica Lectin
A sialic acid-binding lectin was also isolated from eggs of Rana japonica, the Japanese rice paddy frog (Kawauchi et aL, 1975). This lectin agglutinates Ehrlich ascites tumor cells (Kawauchi et aL, 1975) and a number of other tumor cell lines (Sakakibara et al., 1979), but not erythrocytes. The lectin specificity of this protein is similar to that of the sialic acid-binding lectin from R. catesbeiana (Nitta et aL, 1987). Glycoproteins and gangliosides inhibit tumor cell agglutination but no monosaccharides or disaccharides assayed were able to prevent agglutination. Sialic acids on a glycoprotein isolated as a lectin inhibitor appear important for agglutination inhibition (Sakakibara et aL, 1979). Sequencing of this lectin showed close homology to the R. catesbeiana sialic acid-binding lectin (Kamiya et aL, 1990) (Fig. 2). Most of the biological properties of the R. japonica lectin are very similar to those of the R. catesbeiana sialic acid-binding lectin, as discussed below. 3. R a n a catesbeiana Liver RNase
Another frog RNase, isolated from R. catesbeiana liver (Nagano et al., 1976), was sequenced (Nitta et al., 1989). It is 70% identical to the R. catesbeiana sialic acid-binding lectin and 65% identical to the R. japonica sialic acid-binding lectin (Fig. 2). Although it is very similar to the Rana lectins in sequence and enzymatic properties, it is almost 100 times less effective than the lectins in tumor cell agglutination (Okabe et al., 1991).
4. Comparison of in Vitro Properties The sialic acid-binding lectins selectively agglutinate tumor cells (Kawauchi et aL, 1975). Certain mixtures of gangliosides and a glycoprotein
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Antitumor RNases
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blocked agglutination by the R. japonica lectin and neuraminidase treatment of the glycoprotein prevented the inhibition of agglutination, indicating that sialic acid residues were involved in the binding specificities (Nitta et al., 1987). Heparin, mucin, and spermine all inhibit agglutination of sialic acid-binding lectins of R. japonica and R. catesbeiana, whereas asialomucin was less active (Nitta et aL, 1994b). Further evidence for a sialic acid-containing receptor mediating the toxicity of the lectins has been obtained (Nitta et al., 1994a). Sialidase treatment of cells decreased toxicity. A cell population was selected for resistance to the toxicity of the R. catesbeiana sialic acid-binding lectin by growing P388 leukemia cells in the presence of the lectin. However, the cells remained a mixture of survivors and were apparently not picked clonally as individual cell lines. The selected cells were 100-fold resistant to lectin cytotoxicity, yet the surface binding of the lectin was similar to that of the parental cell line. The authors suggest that decreased internalization of the RNase confers resistance. Comparing the enzymatic activity of the three frog RNases, R. catesbeiana sialic acid-binding lectin, R. catesbeiana liver RNase, and R. japonica sialic acid-binding lectin showed distinct differences with RNase A. At the B~ nucleotide binding site of the Rana enzymes uridine is preferred over cytidine and at the B2 subsite guanosine is preferred over uridine, which is preferred over adenosine or cytidine (Okabe et al., 1991). The dinucleotide specificity of these Rana ribonucleases was quite similar to that of the R. pipiens onconase discussed in detail previously (Boix et al., 1996). Thus it is interesting that the R. catesbeiana liver RNase does not agglutinate tumor cells, even at concentrations 100 times higher than those of the other two lectins that agglutinate cells (Okabe et al., 1991). It remains to be seen whether the liver RNase has antitumor activity and it would be an interesting experiment to determine if tumor cell agglutination correlates with antitumor activity among RNases with such similar sequence and enzyme activity.
5. Antitumor Activity Examining the potential for Rana proteins in cancer models, both the R. catesbeiana and R. japonica sialic acid-binding lectins were added to tumor cells in vitro. The lectins inhibited cell growth at < 1 - 1 0 ~g/ml, depending on the cell line assayed (Nitta et al., 1994b). As an in vivo model, tumor cells were injected intraperitoneally followed by intraperitoneal RNase 24 hr later. Significant extension in survival was seen after
508
Richard J. Youle and Giuseppe D'Alessio
administering doses of R. catesbeiana sialic acid-binding lectins ranging from 0.01 to 1 mg/mouse. Animal toxicity was indicated by transient weight loss in the highest dose range, around 1 mg/mouse. Fractionated doses, to approach a continuous infusion, resulted in greater antitumor activity (Nitta et aL, 1994b), similar to the results found with onconase (Mikulski et aL, 1990b), as previously discussed.
IV. C O N C L U D I N G R E M A R K S
Although many of the structural features that mediate the toxicity of certain members of the RNase family are being deciphered, it remains unclear as to why tumor cells are selectively sensitive to these RNases and why some RNases can extend survival in animal models (Laccetti et al., 1992; Mikulski et al., 1990b; Nitta et al., 1994b). Perhaps the ribonucleases selectively bind, enter, or destroy tumor cells. On the other hand, perhaps normal cells are affected similarly by the RNases, but tumor cells respond differently--due to a more rapid growth rate, a deregulated mitotic pathway, or other metabolic defect. Discerning the common structural and functional features of the antitumor RNases that distinguish them from other members of the RNase A family should help elucidate their mechanism. For example, the resistance of all the antitumor RNases to RI, as discussed in this review, suggests that this is an important feature for cytotoxicity. Another intriguing feature is the dimer structure of BS-RNase. Studies on the bovine seminal RNase show that dimer formation is crucial for cytotoxicity. Although the frog RNases do not form covalent dimers, to date there is no information about the possibility that they form noncovalent dimers. However, for a protein to agglutinate cells it has to bind to two cells simultaneously. For example, ricin is a weak agglutinin and it has one high-affinity galactose-binding site. A related R i c i n u s agglutinin is a dimer of ricins with two galactose-binding sites and more potently agglutinates cells. With the small size and single active site in the RNases, it would be easier to conceive of linking cells together if the RNases formed dimers. Nucleotides can partially block agglutination activity, indicating that the enzyme active site may participate in cell binding (Okabe et al., 1991) or, more speculatively, in RNase dimer formation. RNase dimers have a qualitatively different substrate specificity compared to monomers. RNase dimers can cleave double-stranded R N A (Libonati
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and Floridi, 1969). d s R N A cleavage may itself impart some tumorselective killing because free d s R N A in a cell can activate the 2',5'oligoadenylate pathway. This pathway is composed of a protein kinase that phosphorylates eucaryotic initiation factor-2 to block protein synthesis, and a 2',5'-oligoadenylate synthetase that produces unusual 2',5'linked oligoadenylates, which in turn activate RNase L, which can cleave ribosomal RNA. Both the protein kinase and oligoadenylate synthetase are activated by d s R N A and on activation inhibit cell division. This same pathway is activated by interferon, a cytokine with clinical antitumor activity. Thus, production of d s R N A breaks in a cell by a dimeric RNase could activate an endogenous cascade that may selectively kill tumor cells. One other feature is that the frog egg lectins are more stable to heat, pH, and guanidine hydrochloride denaturation than is RNase A (Liao, 1992; Okabe et al., 1991), possibly due to the C-terminal disulfide bond. Onconase is also more stable in vivo than are a number of other RNases (Vasandani, et al., 1996). Exceptional stability within cells and tissues may contribute to the exceptional in vivo properties of these proteins. With the structural relationship among members of the RNase superfamily becoming well understood, disparate research efforts are converging to some degree and similar assays, such as RNase enzyme activity, protein synthesis inhibition, and ascites tumor diminution, are utilized with many of these antitumor RNases, including BS-RNase, onconase, and R a n a lectins. Further solving the mysteries as to the mechanism of the antitumor activity of these proteins may well allow the development of new cancer therapeutics with entirely new mechanisms of action.
ACKNOWLEDGMENTS
We thank Drs. Ester Boix, Shailendra Saxena, Ravi Sirdeshmukh, and Veena Vasandani for many thoughtful discussions and for reading the manuscript. The support of the Associazione ltaliana per la Ricerca sul Cancro is gratefully acknowledged.
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Ardelt, W., Mikulski, S. M., and Shogen, K. (1991). Amino acid sequence of an anti-tumor protein from Rana pipiens oocytes and early embryos. J. Biol. Chem. 266, 245-251. Ardelt, W., Lee, H.-S., Randolph, G., Viera, A., Mikulski, S. M., and Shogen, K. (1994). Enzymatic characterization of onconase, a novel ribonuclease with antitumor activity. Protein Sci. 3(Suppl. 1), 137 (Abstract 486). Bartholeyns, J., and Baudhuin, P. (1976). Inhibition of tumor cell proliferation by dimerized ribonuclease. Proc. Natl. Acad. Sci. U.S.A. 73, 573-576. Becker, F. F. (1975). "Cancer," Vol. 4. Plenum, New York. Boix, E., Wu, Y. N., Vasandani, V. M., Saxena, S. K., Ardelt, W., Ladner, J., and Youle, R. J. (1996). Role of the N-terminus in RNase A homologues: Differences in catalytic activity, ribonuclease inhibitor interaction and cytotoxicity. J. Mol. Biol. 257, 992-1007. Cafaro, V., DeLorenzo, C., Piccoli, R., Bracale, A., Mastronicola, M. R., DiDonato, A., and D'Alessio, G. (1995). The antitumor action of seminal ribonuclease and its quaternary conformation. FEBS Lett. 354, 31-34. Cinatl, J., Matousek, J., and Stanek, R. (1977). Action of RNase A and AS RNase on growth of cells in vitro. Folia Biol. (Prague) 23, 235-244. D'Alessio, G. (1993). New and cryptic biological messages from RNases. Trends Cell Biol. 3, 106-109. Daoust, R., and DeLamirande, G. (1975). Ribonucleases and neoplasia. Sub-Cell. Biochem. 4, 185-211. Darzynkiewicz, Z., Carter, S. P., Mikulski, S. M., Ardelt, W. J., and Shogen, K. (1988). Cytostatic and cytotoxic effects of Pannon (P-30 protein), a novel anticancer agent. Cell Tissue Kinet. 21, 169-182. Di Donato, A., Cafaro, V., and D'Alessio, G. (1994). Ribonuclease A can be transformed into a dimeric ribonuclease with antitumor activity. J. Biol. Chem. 269, 17394-17396. Di Donato, A., Cafaro, V., Romeo, I., and D'Alessio, G. (1995). Hints on the evolutionary design of a dimeric RNase with special bioactions. Protein Sci. 4, 1470-1477. Gilliland, G. L., Dill, J., Pechik, I., Svensson, L. A., and Sjolin, L. (1994). The active site of bovine pancreatic ribonuclease: An example of solvent modulated specificity. Protein Pept. Lett. 1, 60-65. Kamiya, Y., Oyama, F., Oyama, R., Sakakibara, F., Nitta, K., Kawauchi, H., Takayanagi, Y., and Titani, K. (1990). Amino acid sequence of a lectin from Japanese frog (Rana japonica) eggs. J. Biochem. (Tokyo) 1tl8, 139-143. Kawauchi, H., Sakakibara, F., and Watanabe, K. (1975). Agglutinins of frog eggs: A new class of proteins causing preferential agglutination of tumor cells. Experientia 31, 364-365. Kim, J. S., Soucek, J., Matousek, J., and Raines, R. T. (1995). Catalytic activity of bovine seminal ribonuclease is essential for its immunosuppressive and other biological activities. Biochem. J. 308, 547-550. Kobe, B., and Deisenhofer, J. (1993). Crystal structure of porcine ribonuclease inhibitor, a protein with leucine-rich repeats. Nature (London) 366, 751-756. Kobe, B., and Deisenhofer, J. (1995). A structural basis of the interactions between leucinerich repeats and protein ligands. Nature (London) 374, 183-186. Kraft, N., and Shortman, K. (1970). The phylogeny of the ribonuclease-ribonuclease inhibitor system: Its distribution in tissues and its response during leukaemogenesis and aging. Aust. J. Biol. Sci. 23, 175-184. Laccetti, P., Portella, G., Mastronicola, M. R., Russo, A., Piccoli, R., and D'Alessio, G. (1992). In vivo and in vitro growth-inhibitory effect of bovine seminal ribonuclease
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16 2-5A-Dependent RNase L: A Regulated Endoribonuclease in the Interferon System R O B E R T H. S I L V E R M A N Department of Cancer Biology Research Institute, NN1 The Cleveland Clinic Foundation Cleveland, Ohio 44195
I. Perspectives on the 2-5A System II. Structure and Function of RNase L A. Physical Features B. Ankyrin Repeats C. 2-5A Binding Domain D. Protein Kinase and Cysteine-Rich Domains E. Carboxyl-Terminal Sequence and Ribonuclease Activity F. Homologies to Ribonucleases III. Biochemical Properties of RNase L A. Sequence Specificity B. Interactions with 2',5'-Linked Oligoadenylates C. Effects of ATP and Divalent Cations on Activity D. Dimerization in Response to 2-5A E. Binding to Heterologous Proteins IV. Distribution, Localization, and Regulation of RNase L and Its Gene A. Mammalian Cell-Type Distribution B. Intracellular Locations C. Localization of the RNase L Gene, R N S 4 D. Regulation of RNase L Levels E. Evolutionary Distribution V. Biological Activities of RNase L A. Antiviral Activity B. Antiproliferative Effects References
515 RIBONUCLEASES: STRUCTURES AND FUNCTIONS
Copyright 9 1997 by Academic Press, Inc. All rights of reproduction in any form reserved.
Robert H. Silverman
516 s3ATP 2-5A Synthetase ~dslNA ~2PPi pppA2'pS'A2'p5'A "2-5A"
RNase L inactive
2'PDE_.~I~ ATP + 2AMP P,tase'~~A2'p5'pA2'p 5'A + 3Pi
2-5A/RNaseL Activated
Fig. 1. The 2-5A system: an RNA degradation pathway.
I. P E R S P E C T I V E S O N T H E 2 - 5 A S Y S T E M
Eukaryotic cells maintain intricate and complex controls over virtually every step in the biology of RNA, including the synthesis, splicing/ processing, transport/localization, utilization, and destruction of R N A molecules. By catalyzing R N A decay, the cellular ribonucleases counterbalance transcription and are thus essential to the control of gene expression. Induction of RNA turnover can also provide an important host defense mechanism against certain viruses. The 2 - 5 A system (Fig. 1) (Kerr and Brown, 1978) is an R N A degradation pathway present in cells of higher vertebrates; it functions in the antiviral and anticellular activities of interferon and possibly in the more general control of R N A decay. The biological effects of the 2 - 5 A system are catalyzed by a unique endoribonuclease called 2-5A-dependent RNase (or RNase L).* RNase L is an unusual ribonuclease because it requires allosteric effectors to catalyze the hydrolysis of single-stranded RNA. Furthermore, the effectors are themselves very unusual, consisting of a type of oligoadenylate called 2-5A, with 2' --* 5' internucleotide linkages, in contrast to the typical 3'--> 5' linkages found in R N A and D N A (Fig. 2). Why the activators of RNase L possess this structure is unknown except that it provides an unambiguous signal for induction of R N A decay. This chapter provides the first extensive review of RNase L and will complement several previous reviews and volumes on the 2 - 5 A system and interferon * The name "RNase L," formerly 2-5A-dependent RNase, was recently recommended by the Nomenclature Committee of the International Society for Interferon and Cytokine Research [J. Interferon Cytokine Res. (1995) 15, 192].
16
517
2-5A-Dependent RNase L
NH2
O O--- P - - O - -
O
O
N
P--O--
P--O
I
I
O-
I
O-
O-
,,
,,
,,
O
V"
i/
NH2 N
O ,,
N
OH O - - P - - O ~ . O i O-
NH2 N
O II OH O -- O P--~ O . ~ o _ 1
N
OH OH Fig. 2.
Structure of the trimeric species of 2 - 5 A .
action (e.g., Lengyel, 1982; Johnston and Torrence, 1984; Williams and Silverman, 1985; Pestka et al., 1987; Sen and Lengyel, 1992; Kerr and Stark, 1992; Silverman, 1994). Events leading to the discovery of the 2-5A system and RNase L have been reviewed in Johnston and Torrence (1984) and are only mentioned here. Key advances in the field were (1) finding that protein synthesis in extracts of interferon-treated cells was exquisitely sensitive to inhibition by double-stranded RNA (dsRNA) (Kerr et al., 1974), leading to the discovery of 2-5A (Kerr and Brown, 1978); (2) finding that there was endoribonuclease activity in extracts of interferon-treated cells that required both dsRNA (Brown et al., 1976; Kerr et al., 1976) and ATP (Sen et aL, 1976); and (3) the direct demonstration of 2-5A-dependent RNase activity by Clemens and Williams (1978) (see also Baglioni et al., 1978; Ratner et al., 1978; Schmidt et al., 1978; Eppstein and Samuel, 1978; Williams et al., 1978). The 2-5A system functions in the molecular mechanism of interferon action against certain viruses. Interferons are cytokines that act, in part, as an early warning system that a virus infection has occurred. The type I interferons, mainly the a-interferons and/3-interferon, are produced and secreted by virus-infected cells (Vilcek and Sen, 1996). Interferons bind to specific cell surface receptors, activate Jak-STAT signal transduc-
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Robert H. Silverman
tion pathways, and induce selective patterns of gene expression (reviewed in Darnell et al., 1994). The proteins encoded by the interferonstimulated genes are responsible for the various biological activities of the interferons, including the antiviral state, cell growth inhibition, and a wide range of effects on the immune system. There are three wellestablished antiviral pathways of interferon action involving the dsRNAdependent protein kinase, PKR (reviewed in Williams, 1995), the Mx proteins (reviewed in Pavlovic and Staeheli, 1991), and the 2 - 5 A system (Fig. 1). Interferon treatment of cells induces transcription of both a set of genes encoding at least four different species of 2-5A synthetase (Chebath et al., 1987a) and a single gene encoding RNase L (Zhou et al., 1993); however, there are basal levels of these enzyme, especially RNase L, in most mammalian cells. The synthetases are activated by binding to dsRNA (Hovanessian et aL, 1977), a frequent by-product of virus infection. Once stimulated, the 2-5A synthetases convert ATP to PP~ and to a series of 2-5A molecules, such as the predominant trimeric species ppp5'(A2'p5')zA (Fig. 2). 2-5A is unstable in cells and in cellfree systems due to the action of 2' ,5 '-phosphodiesterase (Williams et al., 1978; Schmidt et al., 1979; Minks et al., 1979; Johnston and Hearl, 1987) and, probably, 5'-phosphatase activities (Fig. 1). At subnanomolar levels, 2-5A functions by binding to RNase L, converting it from its inactive to its catalytically active form.
II. S T R U C T U R E A N D F U N C T I O N OF RNase L
A . Physical Features
RNase L contains an array of structural and functional domains that are unique among known and sequenced ribonucleases (Fig. 3). The present analysis was made possible by the molecular cloning of murine and human RNase L (Zhou et al., 1993; Hassel et al., 1993). The initial cDNA to murine RNase L, lacking 89 3'-terminal codons, was obtained by a ligand screening protocol in which a 32p-labeled 2-5A analog was used to probe an expression library of interferon (a +/~) and cycloheximide-treated mouse L cell cDNA (Zhou et al., 1993). The partial murine RNase L cDNA was used as a probe in the cloning of the complete coding sequence of human RNase L. A preliminary view of
16
2-5A-Dependent RNase L
519
Fig. 3. Structural and functional domains of human and murine RNase L.
RNase L emerged from computer-assisted homology searches, deletion analysis, site-directed mutagenesis, and protein-protein interaction studies (Zhou et al., 1993; Hassel et al., 1993; Dong et al., 1995) (Fig. 3). The human form of RNase L is a 741-amino-acid polypeptide with a molecular mass of 83,543 Da (Fig. 4A). However, in the presence of 2-5A activators, RNase L dimerizes to form the catalytically active enzyme (Dong and Silverman, 1995) (Section III,D). Homogeneous, recombinant human RNase L is fully active in the presence of 2-5A and, therefore, RNase L does not require any other polypeptides (Dong et al., 1994). The charge density of human RNase L is relatively nonuniform (Fig. 4B). For instance, there is a group of negatively charge amino acids (EDVENEEDE) from positions 451-459 and a positively charged group at 677-684 (KHKKMKLK) (Zhou et al., 1993). The composition of human RNase L is greater than one-third charged amino acid residues (with 113 aspartic acids plus glutamic acids and 100 arginines plus lysines). However, the significance of the charge distribution is presently unknown (Fig. 4D). The potential functions of each of the domains, motifs, and homologies in RNase L will be individually considered in Sections II,B-II,D.
B. Ankyrin Repeats An intriguing feature of RNase L is the presence in the amino-terminal half of the enzyme of nine units of about 33 amino acids in length, each
A
Human RNase L 83543 m.w.
Molecular Weight
741
Length r amino acids
11.970 pMoles
1 microgram = Molar Extinction coefficient
64550+5%
1 A(280)=
1.29 m,q/ml
Charge Density of Human RNase L I
'
'
'
'
'
~
'
'
'
'
'
'
~
'
I
50 100 150 200 250 300 350 400 450 500 550 600 650 700
17
I
Average Charge
Positive Region
+
:',
9-13-1~
' - l~-IOQI-X3 ~ - - - F - 9 0 " ~ [ ] '."F ':9 - 1: 0 - , Negative Region .i i
,r,., lull
'[]C] .
' 9 '" fill
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'.
"
9
9
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9 i
9
'i
Composition of Human RNase L Amino Acid(s) Charged (RKHYCDE) Acidic (DE) Basic (KR) Polar (NCQSTY) Hydrophobic (AILFWV) A Ala C Cys D Asp E Glu F Phe G Gly H His I lie K Lys LLeu M Met N Asn P Pro Q Gln R Arg S Ser T Thr V Val W Trp Y Tyr
Number count 276 113 100 164 246 50 14 51 62 28 52 31 29 62 85 11 31 24 27 38 45 29 47 7 18
% by weiqht 43.56 16.61 16.62 21.82 31.77 4.26 1.73 7.03 9.58 4.93 3.55 509 3.93 9.51 11.52 1.73 4.23 2.79 4.14 7.10 4.69 3.51 5.58 1.56 3.52
% by frequency, 37.25 15.25 13.50 22.13 33.20 i 6.75 / 1.89 I 6.88 8.37 3.78 7.02 4.18 3.91 8.37 11.47 1.48 4.18 3.24 3.64 5.13 6.07 3.91 6.34 0.94 2.43
Fig. 4. Physical features of RNase L. (A) Data on human RNase L, (B) charge density of human RNase L, and (C) composition of human RNase L. The computer analysis was performed using the Lasergene program (DNASTAR Inc., Madison, Wi.).
16
2-5A-Dependent RNase L
521
containing an ankyrin-related repeat sequence (the nineth ankyrin repeat is incomplete) (Hassel et al., 1993) (Fig. 3). Ankyrin repeats, named after the ankyrin family of proteins, mediate interactions between and within many different proteins. These usually consist of 33 amino acid sequences of the general formula -G-(T/S)(P/A)LHhAA--GH--h(V/A)-LL--GA--(D/N) .... , where h is any hydrophobic amino acid (Breeden and Nasmyth, 1987; Lux et al., 1990; Blank et al., 1992; Nolan and Baltimore, 1992). Ankyrins binds integral membrane proteins and tubulin through their N-terminal domains consisting of 22 such elements (Lux et al., 1990). The central domain of erythrocyte ankyrin, which lacks ankyrin repeats, binds to spectrins and vimentin. Therefore, in a single protein, ankyrin repeats may bind some proteins whereas other proteins may attach through additional interaction domains. Interestingly, a number of proteins that contain ankyrin repeats function in transcriptional control, cell cycle regulation, and cell differentiation (Breeden and Nasmyth, 1987; Lux et al., 1990). In particular, the I-KB family of proteins contain six or seven ankyrin repeats required for the interaction with and inhibition of transcription factor NF-KB (reviewed in Blank et al., 1992; Nolan and Baltimore, 1992). Ankyrin repeat interactions can be either intermolecular (as in the case of I-KB and NF-KB) or intramolecular (as in p105, an inactive precursor of NF-KB subunit p50 and an ankyrin repeat inhibitory domain). However, the only function in common to all ankyrin repeat proteins may be the ability to contact and bind other protein sequences. Although the function of these repeats in RNase L is unknown, a controlling role in the dimerization and activation of the enzyme seems likely (Sections II,C and III,D). It is noteworthy that RNase L is the only nuclease known to contain ankyrin repeats, thus further underscoring its complexity.
C. 2 - 5 A Binding Domain
The ability to bind 2-5A with high affinity (gd = (4-11) X 10 -1~ M) (Silverman et al., 1988; Bayard et al., 1994) is a unique biochemical property of RNase L (Section Ill,B). In addition, the 2-5A binding domain is intrinsically interesting because it represents a new paradigm for protein/nucleic acid interactions. The path to localizing the 2-5A binding domain began with the isolation of a partial murine RNase L (lacking 89 C-terminal amino acids) by screening a cDNA library with
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Robert H. Siiverman
a radioactive 2-5A analog (Zhou et al., 1993) (Section II,A). Expression in vitro of nested 3'-terminal deletions of that cDNA clone showed that 2 - 5 A binding activity resides in the amino-terminal half of RNase L. Evidence to date favors a critical role of a conserved, duplicated phosphate-binding loop (P-loop) motif from residues 229-241 (GX6GXzGKT) and 253-275 [GX9(G/X)X9GKT] present in the human and murine forms of RNase L (Zhou et al., 1993). The P-loop motifs, formerly motif A (Walker et aL, 1982), are found in many adenine and guanine nucleotide-binding proteins in which the conserved lysine residues may interact with phosphate groups in the nucleotides (reviewed in Saraste et aL, 1990). These sequences are often present as a loop between a/3 strand and an a helix, as in adenylate kinases and the p21 ras protein. It is unknown if the P-loop motifs in RNase L form a similar structure. However, the essential role of the conserved lysines in these sequences was shown in site-directed mutagenesis studies (Zhou et al., 1993). Substitution of both lysines, at positions 240 and 274, with asparagines greatly reduced affinity for 2-5A (Zhou et aL, 1993). Interestingly, the GKT sequences are present at the same relative positions, amino acids 2 to 4, in the seventh and eighth ankyrin repeats (Hassel et al., 1993). This finding suggests that binding of 2-5A to RNase L could affect protein/protein interactions involving the ankyrin repeat domain. For instance, 2-5A binding could induce a conformation change in the enzyme that releases an internal clamp on the catalytic domain imposed by the ankyrin repeats. Alternatively, 2-5A binding could result in the unmasking of an interaction domain, permitting dimerization and activation of RNase L.
D. Protein Kinase and Cysteine-Rich Domains
RNase L has significant homology in its C-terminal half with protein kinase domains VI and VII and some additional homology to other protein kinase domains (in particular, domain II) (Zhou et al., 1993; Hanks et al., 1988). It is clear that the protein kinase homology region does not participate in 2-5A binding because that function is localized in the amino-terminal half of the protein (Section II,C). RNase L was proposed to be a "functional active protein kinase" with all 11 protein kinase domains, on the basis of a computer-assisted analysis (Bork and
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2-5A-Dependent RNase L
523
Sanders, 1993). However, some of the proposed protein kinase "domains" in RNase L are either incomplete or differ substantially from known protein kinase domains. For instance, "domain I" (IADTSEGGGIY) in human RNase L bears little resemblance to the canonical protein kinase domain I sequence (hGXGXXGXVh, where h is any hydrophobic residue), in motif VII an aspartate replaces a conserved glycine, in motif VIII a glutamine replaces an invariant glutamate, in motif IX an invariant aspartate is replaced with an arginine, and in motif XI a conserved arginine is present in the human but not in the murine form of RNase L. Clearly, computer modeling is no substitute for careful biochemical analysis. Although the functional significance of the protein kinase homology in RNase L remains unknown, it is one of the most interesting features of the enzyme. It would appear that RNase L evolved, at least in part, from a protein kinase. Alternatively, perhaps the similarity is due to evolutionary convergence to gain functions in common with protein kinases, such as ATP binding activity (Zhou et al., 1993). In protein kinases, the conserved lysine in domain II functions in binding to the a- and /3-phosphoryl groups of ATP and the conserved aspartate in domain VII chelates Mg 2§ complexed with ATP (reviewed in Taylor et al., 1992). Both of these amino acids are also present in the protein kinase homology region of RNase L. Although no other ribonucleases are known to have protein kinase domains, at least two serine/threonine protein kinases, DAP kinase (Deiss et al., 1995) and integrin-linked kinase (ILK) (G. Hannigan and S. Dedhar, 1995, personal communication), and two tyrosine protein kinases in Hydra vulgaris (Chan et al., 1994) and the Drosophila shark kinase (Ferrante et aL, 1995), contain ankyrin repeats. DAP kinase, a proposed mediator of y-interferon-induced cell death, has another feature in common with RNase L, two potential P-loop motifs. Within the protein kinase homology region of RNase L is a highcysteine-content region (Zhou et al., 1993). The spacing of these cysteines ( C X n C X 3 C X I 7 C X 3 C , residues 401-436 in human RNase L, and CXI~ CX25CX3CX6C, residues 395-444 in murine RNase L) bears resemblances to some zinc fingers, protein/nucleic acid-binding domains (Evans and Hollenberg, 1988), suggesting a role in RNA binding (Zhou et al., 1993). However, zinc at concentrations as low as 250 nM are inhibitory to the catalytic activity of RNase L and therefore the function of the cysteine-rich region remains unknown (Dong et al., 1994; B. Dong and R. Silverman, 1994, unpublished).
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Robert H. Siiverman
E. Carboxyl-Terminal Sequence and Ribonuclease Activity The catalytic sites in RNase L are currently unknown. However, studies on a truncated form of murine RNase L (ZB1) suggest that within the carboxy-terminal 89 residues are sequences that are required for ribonuclease activity. The ZB1 polypeptide, which functions as a dominant negative mutant, binds 2 - 5 A with full activity but lacks RNase activity. A C-terminal sequence of RNase L downstream of the protein kinase homology region, from amino acids 587 to 706, showed 29% sequence identity to the C-terminal sequence of the yeast IRE-1 protein, a putative protein kinase that is required for myo-inositol synthesis in Saccharomyces cerevisiae (Nikawa and Yamashita, 1992; Bork and Sanders, 1993). The significance of this similarity is presently unknown.
F. Homologies to Ribonucleases RNase L is not closely related to any previously reported nuclease sequence. However, some limited homology (24 to 32% identical plus conservative matches with some gaps) was observed between Escherichia coli RNase E residues 607 to 796 and RNase L residues 160 to 348. RNase E is an endoribonuclease whose sequence specificity ( G / A ) A U U ( A / U ) (Ehretsmann et al., 1992) bears some similarity to the RNase L consensus cleavage specificity of UU and UA (Section III,A). Characteristic ribonuclease domains, such as the pancreatic ribonuclease family signature sequence (C-K-Xz-N-T-F) (Prosite database, entry PS00127), are absent from RNase L. The lack of more substantial homology with other ribonucleases underscores the uniqueness of RNase L as a member of the ribonuclease family.
III. B I O C H E M I C A L P R O P E R T I E S O F R N a s e L
A. Sequence Specificity The general biochemical features of RNase L are summarized in Table I. RNase L is a general, single-stranded specific endoribonuclease
16
525
2-5A-Dependent RNase L
TABLE I Biochemical Properties of RNase L Property Size Functional definition Specificity Divalent cation effects ATP effect
Comments 83.5 kDa (forms homodimers when activated) A single-strand-specific endoribonuclease when activated by 2 - 5 A [e.g., p3A(2'p5'A)3]; lacks ribonuclease activity in the absence of 2-5A Most frequent cleavages at -UpNp/N, with preference for UU and UA Mg 2§ or Mn 2+ enhances 2-5A binding and ribonuclease activity Ribonuclease activity in the presence of 2-5A is moderately enhanced by ATP; ATP does not activate RNase L on its own.
with no intrinsic specificity for viral versus cellular RNA substrates. However, in the context of intact cells, it remains possible that the 2-5A system might discriminate between different potential RNA substrates. The RNA sequence preferences of RNase L were determined in parallel studies by the Kerr and Lengyel laboratories (Wreschner et al., 1981b; Floyd-Smith et al., 1981). In the former study, end-labeled viral or 18S rRNA substrates were partially digested by RNase L in crude extracts of rabbit, mouse, and human cells in response to the addition of 2-5A. In all three cell-free systems, RNase L showed a strong cleavage preference on the 3' side of UU and UA dimers, leaving 3'-phosphoryl and 5'-hydroxyl groups. Data obtained using partially purified, murine RNase L were in general agreement, with some exceptions (Floyd-Smith et al., 1981). Of the four ribohomopolymers, poly(rU), poly(rC), poly(rG), and poly(rA), only poly(rU) was degraded by RNase L in response to added 2-5A. A 5' end-labeled phage T7-specific RNA, partially digested by RNase L, yielded frequent cleavages after UA, UG, and UU and infrequent cuts after CA, AC, and CG. Recombinant human RNase L, produced in insect cells and purified to homogeneity with fast liquid protein chromatography, has been shown to cleave poly(rU) very efficiently in response to added 2-5A (Dong et al., 1994). In contrast to findings with murine RNase L by Floyd-Smith et al. (1981), poly(rA) was also
526
Robert H. Silverman
degraded, albeit much less efficiently than poly(rU), by recombinant human RNase L. Although many mRNAs encoding various cytokines, protooncoproteins, and lymphokines contain destabilizing, AU-rich sequences in their 3' untranslated regions, it is unknown if RNase L directly cleaves such sequences in intact cells (Shaw and Kamen, 1986; Wreschner and Rechavi, 1988). In contrast, poly(rG), poly(rC), and the single-stranded DNA molecules, poly(dA) and poly(dT), were not cleaved by RNase L (Dong et aL, 1994). Floyd-Smith et al. (1981) reported cleavages in only single-stranded loops of the 3' terminal region of phage R17 RNA by RNase L, indicating at least a preference of RNase L for single-stranded specific regions in RNA. Similarly, Nilsen et al. (1980) reported an absence of 2-5A-dependent cleavage of poly(rA):poly(rU), dsDNA, or ssDNA in crude cell extracts. An interesting suggestion was made that the RNase L cleavage preference for sites containing uridylate residues could be due to base-pairing with an adenylyl residue in 2-5A (Floyd-Smith et aL, 1981). Although the reason for the sequence specificity of RNase L is unknown, a prediction of this hypothesis is that redirecting binding of RNase L to different regions in an RNA substrate will alter its sequence specificity. This was directly demonstrated using 2-5A chemically attached through linkers to antisense deoxyribonucleotides (Torrence et al., 1993; Maitra et al., 1995). The 2-5A part of the chimeric oligonucleotide bound and activated RNase L while the antisense part caused the selective cleavage of "target" RNA molecules containing the complementary sequence. Interestingly, although the target RNA substrates were selectively degraded by RNase L, there was no apparent nucleotide preference in the RNA cleavage sites under these conditions. In one study involving crude cellfree systems, cleavages occurred mainly in a stretch of adenylyl residues in the RNA strands of the R N A : D N A hybrid (Torrence et al., 1993); in a separate study involving purified, recombinant RNase L the principal cleavage occurred 5' (upstream) of an oligonucleotide binding site in the mRNA for the protein kinase, PKR (Maitra et al., 1995). The catalytic and rapid (Kcat of about 7 sec -~) degradation of the PKR mRNA was shown. Apparently, RNase L will cleave RNA sequences to which it is directed regardless of the sequence. The 2-5A antisense approach has been adapted to intact cells, where the selective cleavage of PKR mRNA produced ablation of PKR activity and a defect in dsRNA-mediated activation of transcription factor NF-KB (Maran et al., 1994). By changing the sequence of the antisense cassette, 2-5A antisense has the potential to bring about the selective cleavage of different RNA molecule in vivo.
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2-SA-Dependent RNase L
527
Although RNase L activated by 2-5A in vitro cleaves RNA with, at best, moderate specificity, the selective cleavage of certain RNA species could occur in nature within the context of the intact cell. This could result from the intracellular attachment of 2-5A synthetase or RNase L to structural elements of the cell (Section IV, B), or from the occurrence of local pockets of 2-5A production within the cell. With regard to the latter, Nilsen and Baglioni (1979) proposed a mechanism by which single-stranded RNA sequences attached to double-stranded RNA, such as in the replicative intermediates of certain RNA viruses, can be preferentially degraded by the 2-5A system. In essence, their data show that single-stranded R N A s linked to dsRNA were degraded in extracts of interferon-treated cells more readily than single-stranded RNA lacking dsRNA. Heterogeneous nuclear RNA was also shown to promote synthesis of 2-5A, presumably due to duplex structures, and to be preferentially cleaved in extracts of interferon-treated cells (Nilsen et al., 1982b). Subsequently, Baglioni et al. (1984) showed that nascent reovirus RNA, synthesized in vitro from the dsRNA viral genome from purified virions, was preferentially degraded by RNase L in a cell-free system, apparently the result of localized production of 2-5A. A 2-5A analog inhibitor of RNase L, the 3'-methyl analog of 5'-monophosphorylated 2-5A, prevented reovirus RNA decay, thus providing evidence for the involvement of RNase L. The localized activation model is attractive because it provides a possible mechanism by which RNase L could discriminate between viral and cellular mRNA. However, there is little or no direct evidence that RNase L preferentially cleaves viral RNA, attached to replicative intermediates or otherwise, in intact, interferon-treated cells. To the contrary, in interferon-treated, encephalomyocarditis virus- or reovirus-infected cells there is significant decay of rRNA by RNase L (Wreschner et al., 1981a; Silverman et al., 1983; Nilsen et al., 1982a).
B. Interactions with 2',5'-Linked Oligoadenylates A combined chemical and enzymatic approach has produced a considerable amount of highly precise information about the atoms or groups of atoms in 2-5A that interact with RNase L [reviewed by Torrence et al. (1994)]. The goals of such studies are both basic and pharmacologic in nature. Chemically modified derivatives of 2-5A provide tools for probing the 2-5A binding/activation domain of RNase L. In addition,
528
Robert H. Silverman
2 - 5 A analog inhibitors of RNase L have been shown to be important reagents for studying the physiological role of the 2 - 5 A system (e.g., Torrence et aL, 1981, Baglioni et al., 1984; Watling et al., 1985). Moreover, because the catalytically active form of RNase L is a potent antiviral and anticellular protein, stable derivatives of 2 - 5 A able to penetrate cell membranes could provide novel therapeutic agents for viral infections, cancer, and other diseases (e.g., Torrence et al., 1993; Maran et al., 1994). The interactions of 2 - 5 A with RNase L have been measured at three levels: (1) association/dissociation (Knight et aL, 1980; Silverman and Krause, 1987), (2) activation of the catalytic function of RNase L (Clemens and Williams, 1978; Krause et aL, 1986), and (3) induction of RNase L homodimers (Dong and Silverman, 1995). The affinity of 2 - 5 A for RNase L is typically measured in a radiobinding assay using a 32P-labeled derivative of 2 - 5 A (Silverman et al., 1981; Nolan-Sorden et al., 1990). The bound, labeled 2 - 5 A derivative is measured either by a filter binding method (Knight et al., 1980) or after covalent cross-linking and gel electrophoresis (Wreschner et al., 1982; Floyd-Smith et al., 1982; NolanSorden et al., 1990). Analysis of Scatchard plots for the 2 - 5 A binding activity of the murine RNase L resulted in Kj values of (4-11) • 10 -11 M (Silverman et al., 1988; Bayard et al., 1994). The higher estimates (about 10 -l~ M) reported by Nilsen et aL (1981) could be due to differences in the method used to prepare the labeled 2 - 5 A probe. The affinity of human RNase L for 2 - 5 A was estimated to be about threefold lower than for the murine enzyme (Nilsen et al., 1981). In mouse liver, it was determined that there is 5.5 nmol of 2 - 5 A binding sites per kilogram of tissue (Silverman et al., 1988). Alternative 2-5A-binding polypeptides have been observed (Wreschner et al., 1982; St. Laurent et al., 1983); in at least some cases, these resulted from proteolysis of RNase L (Krause et al., 1985b). Early studies involving crude cell extracts revealed a requirement of at least two 5'-phosphoryl groups and at least three 2',5'-linked adenylyl residues for efficient activation of murine RNase L (Martin et al., 1979; Kerr and Brown 1978). Subsequently, it was shown that only a single 5'-phosphoryl group on the trimer 2',5'-linked oligoadenylate was required for efficient activation of human RNase L (Haugh et aL, 1983). The length of the 2 - 5 A species is not critical for activation of RNase L, provided there are at least three adenylyl residues present. However, a curious and unexplained finding is that RNase L from rabbit reticulocytes requires 2 - 5 A tetramer for its activity (Williams et al., 1979b), whereas
16 2-5A-Dependent RNase L
529
rabbit kidney RNase L responds to the trimer form of 2-5A (Krause and Silverman, 1993). At a concentration of 1 n M , the tetrameric to heptameric species, ppp5'(A2'p5')nA, n = 2-6, all activated murine RNase L, although the hexamer and heptamer were slightly less active (Haugh et al., 1983). The essential role of the adenylyl residues in 2 - 5 A was shown by the findings that RNase L has at least a 2000-fold lower affinity for the cytidine, uridine, or inosine analogs [p5'(C2'p5')2C, p5'(U2'p5')2U, and p5'(I2'p5')2I] compared to p5'(A2'p5')2A (Torrence et al., 1984); ppp5'A2'p5'A2'p5'N, where N is any of the common nucleotides other than adenylate, apparently failed to activate RNase L (Drocourt et al., 1982). These findings led Torrence et al. (1984) to suggest that RNase L interacts with N 1 nitrogen and/or the adenine 6-amino groups in 2-5A. In this regard, substitution of individual adenosine residues in the 2-5A trimer with inosines showed that the 5'-terminal adenine base is critical for both activation and binding to RNase L, the middle adenine is less important for these activities, and the 3'-terminal adenine is critical for activation but not binding to RNase L (Imai et al., 1985). Additional studies with individual 3'-deoxyadenosine-substituted derivatives of ppp5'(A2'p5')2A showed that only the 3'-OH of the central ribose is required for efficient activation of RNase L (Torrence et al., 1988). On the basis of these and other findings involving fluoro sugar analogs of 2-5A, it was suggested that the 3'-hydroxyl of the central adenosine of trimer 2-5A may hydrogen bond to an acceptor in RNase L (Torrence et al., 1994). Requirements for activation of human RNase L were directly demonstrated with homogeneous, recombinant enzyme (Dong et al., 1994). The dimeric species, pppA2'p5'A, was completely lacking in ability to activate RNase L, whereas ppp5'(A2'p5')2A, pp5'(A2'p5')3A, and p5'(A2'p5')3A, each 1 nM, maximally activated RNase L. Derivatives lacking 5'-phosphoryl groups showed greatly reduced ability to activate RNase L [e.g., A(2'p5')2A was 100-fold less active than the 5'-phosphorylated species], whereas the 3',5'-linked oligoadenylate, ppp5' (A3'p5')2A, completely failed to activate RNase L (thus confirming Lesiak et al., 1983). Another indication of the importance of the internucleotide linkages was provided by the observation that the stereochemistry of phosphorothioate linkage analogs of 2-5A affected the ability to activate a glutathione S-transferase/human RNase L (Sobol et al., 1995). To summarize, the current definition of 2-5A for activation of human RNase L is px5'(A2'p5')nA (x = 1-3; n > 2). As previously mentioned, the murine form of RNase L may require 2-5A containing a 5'-diphosphate.
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Robert H. Silverman
C. Effects of A T P and Divalent Cations on Activity
ATP stimulates or stabilizes RNase L activity in the presence of 2-5A, but has no effect in the absence of 2-5A. Wreschner et al. (1982) first reported that the presence of ATP could significantly stimulate or preserve the activity of RNase L during gel filtration of rabbit reticulocyte lysate. Addition of ATP or ADP, and to a lesser extent AMP, to a column buffer prevented an irreversible loss in RNase L activity that occurred during filtration through a Sepharose G-25 column. Subsequently, partial purification of murine RNase L and complete purification of the recombinant human RNase L showed about a twofold stimulatory effect of ATP on RNase L activity (Krause et aL, 1986; Dong et al., 1994). However, in these studies removal of ATP during purification of RNase L did not prevent subsequent activation of the enzyme by 2-5A. In addition, nonhydrolyzable ATP analogs were used to show that ATP hydrolysis was not required for the stimulation of RNase L activity (Krause et al., 1986). It was further shown that the ATP effect requires the presence of magnesium, manganese, or calcium (Dong et al., 1994). Nevertheless, RNase L that was extensively dialyzed and assayed in buffer lacking ATP and divalent cations and containing chelators (EGTA and E D T A ) retained a low, but clearly measurable, residual level of 2-5A-dependent ribonuclease activity. This residual activity was not unexpected because other ribonucleases that produce RNA cleavage products containing 3'-phosphates do not require divalent cations (Deutscher, 1993). Addition of divalent cations or ATP alone had no effect, but addition of both ATP and either magnesium or manganese greatly stimulated 2-5A-dependent RNase L activity (Dong et al., 1994). Apparently, divalent cations facilitate the binding of ATP and 2 - 5 A to RNase L. As mentioned previously, the ATP effect might be mediated through the protein kinase homology region in RNase L (Zhou et al., 1993). The presence of a reducing agent, 2-mercaptoethanol, was also reported to be necessary to maintain 2 - 5 A binding activity of RNase L (Bayard et al., 1994). Interestingly, the requirement for reducing agent was lessened by the addition of poly(rU).
D. Dimerization in Response to 2 - 5 A
Independent estimates of the size of native RNase L as it exists in mammalian cell extracts are usually, but not always, consistent with either a monomeric or dimeric form of the enzyme. For instance, monomeric RNase L was observed in mouse and rabbit reticulocytes (Revel et al.,
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1979; Wreschner et al., 1982), murine Ehrlich ascites tumor (EAT) cells (Floyd-Smith et aL, 1982; Floyd-Smith and Lengyel, 1986), and human HeLa cells and murine W7 cells (Nilsen et al., 1981). In contrast, extracts of murine E A T cells also produced a 185-kDa peak of RNase L activity on Sephadex G-200, especially at high protein concentrations (Slattery et al., 1979; Floyd-Smith et al., 1982; Wreschner et al., 1982). Bayard et al. (1994) also observed that mouse spleen RNase L cross-linked to a 2 - 5 A derivative occurs largely as a dimer of 160 kDa. Curiously, mouse spleen was reported to contain both 185- and 40-kDa species of RNase L (Bisbal et al., 1989). The 40-kDa protein may represent either a breakdown product of RNase L or a separate species of the enzyme. Also, these investigators observed a 160-kDa species of RNase L, reported to be a heterodimer, in extracts of human Daudi cells (Salehzada et al., 1993). These disparate findings show that RNase L exists as monomers or in association with itself or with other proteins (Section Ill,E). Studies show that purified, recombinant human RNase L is a monomer in the absence of 2-5A and a homodimer in the presence of 2-5A (Dong et al., 1994; Dong and Silverman, 1995). Dimerization was demonstrated by three independent methods, gel filtration, sedimentation in glycerol gradients, and chemical cross-linking. Nearly complete conversion to dimeric RNase L was observed at a 1 : 1 molar ratio of 2-5A to RNase L. A close correlation exists between the abilities of different oligoadenylates to induce dimer formation and induce RNase L catalytic activity. For instance, pppA2'p5'A and ppp5'(A3'p5')2A both failed to induce either dimer formation or ribonuclease activity. On the other hand, p5' (A2'p5')2A, pp5'(A2'p5')2A, and p5'(A2'p5')3A were all highly efficient at both inducing dimers and RNA cleavage. Core oligoadenylates lacking 5'-phosphate groups were very inefficient in both activities. Although these data certainly suggest that the catalytically active form of RNase L is a homodimer, additional studies will be necessary to determine if dimerization is actually required for catalysis of RNA decay. One possibility is that the catalytic domain of RNase L is shared between two RNase L monomers. If so, the principal function of 2-5A could be to induce a conformational change in RNase L that exposes the interaction domains (Dong and Silverman, 1995).
E. Binding to Heterologous Proteins
Interactions between RNase L and specific cellular proteins were investigated in two studies from the same group (Salehzada et al., 1993; Bisbal
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etal., 1995). A monoclonal antibody that recognized an 80-kDa R N A binding protein (RNABP) was found to immunoprecipitate both R N A B P and another 80-kDa protein (2-5ABP) from Daudi cell extracts (Salehzada et al., 1993). On the basis of these results, it was proposed that RNase L exists as two heterologous subunits, R N A B P (the RNA-binding and catalytic subunit) and 2 - 5 A B P (the 2 - 5 A binding and regulatory protein). However, when RNase L was cloned, expressed, and purified it was shown to consist of a single protein that formed homodimers in response to 2-5A (Zhou et al., 1993; Dong et al., 1994; Dong and Silverman, 1995). The identity of RNABP remains to be established, but if 2 - 5 A B P is RNase L (as it appears to be based on its size and affinity for 2-5A), then R N A B P is a candidate for a protein that interacts with RNase L. Subsequently Bisbal et al. (1995) reported cloning a 68-kDa protein (RLI, for RNase L inhibitor) by screening a Daudi cDNA expression library with a radiolabeled 2-5A molecule. Sequence comparison between RNase L and RLI did not show any significant homologies, except both proteins contained two Ploop motifs. Expression of a truncated form of RLI modestly reduced the antiencephalomyocarditis virus (EMCV) but not the antivesicular stomatitis virus (VSV) effect of interferon. The affinity of RLI for 2 - 5 A was shown to be poor, and yet RLI reversibly reduced the 2 - 5 A binding activity of RNase L in mammalian cell extracts but not in wheat germ extracts. In a rabbit reticulocyte translation system, immunoprecipitation with antibody against RNABP revealed a complex that included RNABP, RLI, and RNase L. It is presently unknown if either RLI or RNABP directly interacts with RNase L in vivo.
IV. DISTRIBUTION, LOCALIZATION, AND REGULATION OF RNase L A N D ITS GENE
A. Mammalian Cell-Type Distribution
RNase L is very widely distributed in mammalian tissues and cell lines in the absence of interferon treatment. For instance, RNase L was reported in rabbit liver, kidney, spleen, and reticulocytes (Williams et aL, 1979a; Nilsen et al., 1981; Krause and Silverman, 1993), in mouse liver, kidney, lung, intestine, spleen (Nilsen et al., 1981; Floyd-Smith and Denton, 1988a,b; Silverman et al., 1988; Bayard et al., 1994), brain, testis,
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thymus, intestine, and heart (S. Cates and R. Silverman, 1995, unpublished), and a large number of human cell types and in primary tissues such as lymphocytes (e.g., Nilsen et aL, 1981), colonic mucosae (Wang et al., 1995), and kidney (Bayard and Gabrion, 1993). In contrast, RNase L was not found in murine embryonal carcinoma cells prior to differentiation (Krause et al., 1985b) or in the murine neuroblastoma cell line, NIE 115 (Silverman et al., 1986). Spleen, liver, and lymphocytes contained relatively high levels of RNase L (Nilsen et al., 1981), and in another study RNase L amounts were highest in mouse spleen and lung and lowest in liver and intestine (Floyd-Smith and Denton, 1988a). Injection of mice with antibody to ~ +/3 interferon resulted in decreased levels of RNase L in spleen, with little or no effect in other organs, suggesting that endogenous interferon could induce RNase L synthesis in that tissue (Floyd-Smith and Denton, 1988a). In mice, RNase L levels were found to peak early in postnatal development in several mouse organs and then to decrease as the animals aged (Floyd-Smith and Denton, 1988b; Pfeifer et al., 1993). Because most of these studies were performed using 2-5A binding activity as a measure of RNase L, endogenous 2-5A, which competes with the labeled 2-5A probe, could lead to low estimations of RNase L amounts (Bayard and Gabrion, 1993). In this regard, determination of RNase L amounts using a monoclonal antibody to human RNase L showed the presence of RNase L in normal colonic mucosae with elevated levels in colorectal tumors and polyps (Wang et al., 1995). By using known amounts of purified, recombinant RNase L for comparison, it is estimated that the levels of RNase L in normal mucosae and tumors ranged from about 0.0006 to 0.006% of the total soluble protein.
B. Intracellular L o c a t i o n s
Several studies have shown both cytoplasmic and nuclear locations for RNase L in various cell types (e.g., Nilsen et al., 1982c; St. Laurent et al., 1983; Silverman et al., 1988; Schroder et al., 1989; Bayard and Gabrion, 1993), whereas purified NIH 3T3 nuclei lacked RNase L in one study (Salehzada et al., 1991). Bayard and Gabrion (1993) showed that the nuclear form of RNase L is often found complexed with 2 - 5 A or an inhibitor of 2-5A binding. Attempts at more precise localization of RNase L suggest that the cytoplasmic enzyme is largely associated with polysomes (Salehzada et al., 1991) and that the nuclear form of
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RNase L is attached to nuclear matrices (Schroder et al., 1989). More sophisticated analyses, including confocal microscopy with monoclonal antibody to RNase L, will be required to verify these results.
C. Localization of the RNase L Gene, R N S 4
The intrachromosomal location of the human RNase L gene was determined using a genomic RNase L clone and fluorescence in situ hybridization (FISH) (Squire et al., 1994). Positive hybridization signals were consistently found at chromosome band lq25 on all four chromatids in metaphase spreads. The gene was assigned as " R N S 4 " by the H G M W Nomenclature Committee because it was the fourth RNase ( R N S ) gene to be mapped in the human genome. Although there is no direct evidence to link defects in the RNase L gene to known diseases, deletions of lq23-32 occur in many breast cancers and translocation breakpoints at lq25 are found in some oral squamous cell carcinomas (Squire et al., 1994, and references therein). Because RNase L activation can restrict cell growth (Section V,B), it is tempting to speculate that the mutations in the RNase L gene could lead to inappropriate cellular proliferation.
D. Regulation of RNase L Levels
As mentioned previously (Section IV,A), basal levels of RNase L are found in nearly all mammalian cell types. One could infer from this that the RNase L gene is constitutively expressed. However, there are numerous examples of substantial regulation of RNase L gene expression. In murine JLS-V9 R cells, RNase L levels increased 10- to 20-fold in response to interferon treatment or 6- to 10-fold during the transition from actively growing, subconfluent cells to confluent and growtharrested cells (Jacobsen et al., 1983a,b). Growth arrest of murine NIH 3T3 cells led to a 6-fold increase in RNase L levels, which was increased by another factor of two by subsequent treatment with interferon (Krause et al., 1985a). These observations could be due to transcriptional or posttranscriptional regulation. However, interferon treatment of murine L929 cells produced a 3-fold increase in RNase L m R N A levels that was unaffected by the presence of cycloheximide and is therefore
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probably a primary response to interferon treatment (Zhou et al., 1993). Levels of RNase L are also induced as an early event in cell differentiation (Krause et al., 1985b). RNase L was undetectable in three undifferentiated murine embryonal carcinoma cell lines even after interferon treatment, whereas RNase L was interferon inducible in three derivative, differentiated cell lines. The appearance of RNase L was also observed within 3 days of in vitro differentiation of the embryonal carcinoma cell line, PC13, correlating with an increase in the sensitivity of the cells to the antiviral effects of interferon (Krause et al., 1985b). Exposure of murine L929 cells to microwave radiation, at levels insufficient to affect cell proliferation and viability, also resulted in an induction of RNase L (Krause et al., 1991). An increase in RNase L levels in alveolar macrophages after infection with respiratory syncytial virus was reported by Panuska et al., 1995. Finally, a recent study showed that elevated levels of RNase L occur as an early event in colorectal tumorigenesis (Wang et al., 1995). Cellular levels of RNase L were increased by 1.5- to 10fold in 17 of 20 colorectal tumors as compared to normal mucosae. These findings were unexpected because rapidly proliferating cells often contain decreased, rather than increased, levels of RNase L. Perhaps, therefore, a host response involving local production of interferon or another cytokine was responsible for the induction of RNase L in the polyps and tumors.
E. Evolutionary Distribution
As discussed above, RNase L is present in mammals from mice to humans (Section IV,A). Reptiles and avians also contain the 2 - 5 A system (Cayley et al., 1982a; Ball and White, 1978). Members of three reptile families showed higher levels of 2 - 5 A binding activity (an activity associated with RNase L) in liver, kidney, and heart than was seen in some mammalian cell extracts (Cayley et al., 1982a). The reptilian binding activity was shown to be highly specific for 2-5A. Only trace amounts of 2 - 5 A binding activity were observed in three amphibians and no such activity was present in fish, insects, plants, slime molds, or bacteria. We have recently confirmed the absence of RNase L in tobacco plants (SenGupta et al., 1996; Mitra et al., 1996). The 2 - 5 A synthetase and an activity that inhibits protein synthesis in response to addition of 2-5A, presumably RNase L, are present in chick embryo cells (Ball and White, 1978). Therefore, the evidence
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to date seems favor the presence of RNase L in avians, reptiles, and mammals. Rigorous proof of the presence of RNase L in other species is presently lacking.
V. B I O L O G I C A L A C T I V I T I E S O F R N a s e L
A. Antiviral Activity A large body of evidence implicates RNase L in the antiviral activity of interferon for some types of viruses. However, many viruses have acquired mechanisms for blocking the 2-5A system and thus for escaping its antiviral effects. The most compelling evidence that RNase L functions in interferon action is provided by studies involving EMCV, reovirus, and vaccinia virus. A common feature of these viruses is that they lead to production of dsRNA (e.g., Gribaudo et al., 1991; Colby and Duesberg, 1969). In the interferon-treated, virus-infected cells, the dsRNA stimulates 2-5A synthetase to produce 2-5A, thus leading to activation of RNase L. The antiviral effect is caused by the breakdown of both viral and cellular RNA, thus crippling the ability of the cell to produce progeny virus. Picornaviruses are nonenveloped, positive-sense RNA viruses that replicate through a multistranded RNA intermediate. Interferon treatment of cells followed by EMCV infection leads to an accumulation of 2-5A (Williams et al., 1979b; Knight et al., 1980; Silverman et al., 1982a), presumably due to stimulation of 2-5A synthetase by viral dsRNA (Gribaudo et al., 1991). Increased levels of 2-5A were also detected in organs of EMCV-infected mice (Hearl and Johnston, 1987). The detection of specific rRNA cleavage products, characteristic of RNase L digestion of intact ribosomes, provides direct evidence for activation of RNase L in interferon-treated, EMCV-infected cells (Wreschner et al., 1981a; Silverman et aL, 1982a, 1983). However, another picornavirus, poliovirus type 1, did not induce rRNA cleavage products in interferon-treated HeLa cells (Munoz et al., 1983). Strategies that have further implicated involvement of RNase L in antipicornavirus effects of interferons include overexpression of 2-5A synthetase cDNA or inhibition of RNase L function. Expression of cDNA to low molecular forms of 2-5A synthetase greatly reduced (up to 1000-fold) replication of two picornaviruses, mengo virus, and
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EMCV, without affecting replication of a rhabdovirus, VSV, or herpes simplex virus-2 (HSV-2) (Chebath et al., 1987b; Rysiecki et al., 1989; Coccia et al., 1990). Inhibition of RNase L activity has been achieved either by transfecting cells with a 2-5A analog inhibitor of RNase L or by expressing in cells a dominant-negative RNase L mutant (Watling et al., 1985; Hassel et al., 1993). The 2-5A analog, CH3Sp(A2'p)2A2'pp3'OCH3, is a competitive inhibitor of 2-5A binding to RNase L. When the inhibitor was introduced into EMCV-infected, interferon-treated cells by calcium phosphate coprecipitation, it prevented RNase L-mediated rRNA cleavages and reduced the anti-EMCV effect of interferon by as much as 10fold (Watling et al., 1985). The dominant-negative mutant of RNase L, missing 89 carboxy-terminal amino acids, binds 2 - 5 A efficiently but lacks ribonuclease activity, and is a very effective inhibitor of the wild-type RNase L both in vitro and in intact cells (Hassel et al., 1993). When the dominant-negative RNase L was stably expressed in murine SVT2 cells, at a 25ofold excess relative to the endogenous RNase L, it suppressed the anti-EMCV effect of interferon by 250-fold while reducing the effect on VSV replication by only 2.5-fold (Hassel et al., 1993). Lack of substantial inhibition of VSV replication by the 2-5A system contrasts with the presence of 2-5A synthetase in the VSV virions (Wallach and Revel, 1980). The reason for the apparent insensitivity of VSV to effects of the 2-5A system is unknown. However, additional supporting evidence for an antiEMCV effect of RNase L comes from the isolation and characterization of murine RK6 cells, which have only very low levels of RNase L and are resistant to the anti-EMCV effects of interferon (Kumar et al., 1988). Although RNase L functions against EMCV in interferon-treated cells, in the absence of interferon, EMCV infection causes a dramatic reduction in RNase L levels by an unknown mechanism (Cayley et al., 1982b; Silverman et al., 1982a). Interestingly, exposure of cells to interferon protects against EMCV-mediated inactivation of RNase L. Reoviruses contain segmented dsRNA genomes and lead both to accumulation of 2-5A and to breakdown of mRNA and rRNA interferontreated cells (Nilsen et al., 1982a). It was further shown that cells could compensate for the enhanced RNA decay rate, possibly by increasing rates of mRNA production (Nilsen et al., 1983). These studies are at least consistent with the involvement of RNase L in the antireovirus effects of interferon. Vaccinia virus is a member of the poxvirus family containing a large (192 kb) and complex DNA genome. Transcription from opposite strands of vaccinia D N A produces complementary RNA that anneals to form
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dsRNA (Colby and Duesberg, 1969; Boone et al., 1979; Bayliss and Condit, 1993). Viral interference of the 2-5A system was observed in cells that were relatively resistant to the antivaccinia virus activity of interferon. Interferon treatment followed by vaccinia virus infection resulted in an accumulation of high levels of functional 2-5A; however, activation of RNase L, as determined by rRNA breakdown by RNase L, occurred only after a delay (Rice et al., 1984). In contrast, there was a good correlation between time of 2-5A accumulation (to low levels) and rRNA cleavage in another study involving interferon-treated, vaccinia virus-infected cells (Goswami and Sharma, 1984). Esteban et al. (1986) further demonstrated a correlation between the sensitivity of differently cultured murine L cells to the antivaccinia virus effect of interferon and the appearance of specific rRNA cleavage products, apparently the result of RNase L activation. In addition, enzyme activities that can degrade 2-5A were induced in some interferon-treated, vaccinia virus-infected cells (Paez and Esteban, 1984). Interestingly, a mutation in the vaccinia virus A 1 8 R gene causes aberrant transcription, leading to an increase in the accumulation of dsRNA and activation of 2-5A synthetase and RNase L (Bayliss and Condit, 1993). The A 1 8 R gene encodes a 56-kDa protein with homology to an excision repair DNA helicase, further suggesting a role in nucleic acid metabolism (Pacha et al., 1990). Another vaccinia protein that might interfere with 2-5A system, in this case at the level of 2-5A synthetase activation, is the vaccina E3L dsRNA-binding protein (Chang et al., 1992). Therefore, although vaccinia virus infections often lead to activation of 2-5A synthetase and RNase L, in particular in interferon-treated cells, the virus can sometimes prevent or delay RNA decay. In some interferon-treated cells, the 2-5A system probably restricts vaccinia virus replication whereas in other instances the virus effectively escapes. The possible interaction between human immunodeficiency virus type1 (HIV-1) and the 2-5A system has been suggested in several reports. The transactivation-responsive (TAR) RNA at the 5' termini of all HIV-1 mRNAs is sufficiently base-paired to modestly activate 2-5A synthetase in vitro (SenGupta and Silverman, 1989; Maitra et al., 1994; references therein). In addition, during HIV infection there was an increase in RNase L and 2-5A synthetase activity associated with a subcellular fraction containing the nuclear matrix (Schroder et al., 1989). Similarly, Baca et al. (1994) reported an increase in cytosolic 2-5A synthetase following HIVADA infection of human monocytes. Also, expression in cells of a 2-5A synthetase cDNA under control of an HIV long terminal
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repeat resulted in an inhibition in HIV replication (Schroder et al., 1990). It is unknown, however, if RNase L restricts HIV replication in control or interferon-treated wild-type cells. For instance, although an increased rate of cellular m R N A turnover was observed in HIV-infected cells there was a lack of rRNA cleavages typical of RNase L activity (Agy et al., 1990; Katze and Agy, 1990). An intriguing viral escape mechanism was uncovered by careful analysis of the 2' ,5 '-oligoadenylates produced in interferon-treated cells infected with simian virus 40 (SV40), herpes simplex virus, or vaccinia virus. These viruses induce the synthesis of 2 - 5 A per se and/or many related 2',5'-oligoadenylate derivatives, which fail to activate and, in some instances, competitively inhibit 2-5A activation of RNase L (Hersh et al., 1984; Cayley et al., 1984; Rice et al., 1985). The precise structures of the virus-induced inhibitors have not been elucidated. The molecular events responsible for the production of aberrant 2 - 5 A is completely unknown, but it was suggested that perhaps some viral-induced enzymes could rapidly convert active 2-5A to alternative structures that lack the ability to activate RNase L (Hersh et al., 1984). RNase L inhibitors were also found in extracts of interferon-treated cells incubated with activated reovirus cores or poly(I):poly(C) (Williams et al., 1986).
B. Antiproliferative Effects
It is not too difficult to imagine how a ribonuclease could cause the cessation of cell proliferation or even cell death. Simply put, cells cannot tolerate massive and sustained RNA decay. For example, onconase, a frog ribonuclease related to RNase A, is extremely toxic to intact cells (Wu et al., 1993). The possible involvement of RNase L in controlling cell proliferation is less obvious because its activity is completely dependent on 2-5A, the production of which requires both 2-5A synthetase and dsRNA. The potential of RNase L to block cell growth was shown by the direct transfection of murine L929 cells with 2-5A (Hovanessian and Wood, 1980). Growth inhibition by the 2 - 5 A system was also demonstrated by stable expression in mammalian cells of cDNAs for 2-5A synthetase or RNase L. Accumulation of 2-5A synthetase from a transfected cDNA suppressed cell growth rates (Rysiecki et al., 1989), whereas overexpression of RNase L cDNA suppressed and blocked cell growth
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in the absence and presence of interferon, respectively (B. A. Hassel, A. Zhou, S. Shah, and R. H. Silverman, 1996, unpublished). Cell growth regulation by interferons is complex and probably involves several of the many proteins that are inducible by interferon treatment of cells. For example, the activated form of the interferon-inducible and dsRNA-dependent protein kinase, PKR, is known to suppress cell growth rates (Chong et al., 1992). RNase L appears to be one of the mediators of the anticellular effects of interferons. The enzymatic machinery required to degrade RNA, i.e., 2 - 5 A synthetase and RNase L, is induced by interferon treatment of cells (Hovanessian et al., 1977; Jacobsen et al., 1983a; Zhou et al., 1993). In addition, increases in amounts of 2-5A per se have been found after interferon treatment of cells in whole extracts of murine L cells (Knight et al., 1980) as well as in HeLa cell nuclei (Nilsen et aL, 1982c). However, neither 2 - 5 A nor rRNA cleavages could be found in interferon-treated, human Daudi cells despite an exquisite sensitivity to the anticellular effect of interferon (Silverman et al., 1982b). The transient production of low levels of 2 - 5 A could not be completely excluded in these experiments. In contrast, in murine SVT2 cells the anticellular effect of interferon was suppressed with a dominant-negative form of RNase L (Hassel et al., 1993). Although these studies do not prove that RNase L performs a similar function in vivo, they do demonstrate that RNase L activity contributes significantly to the antiproliferative activity of interferons in at least some cell culture systems. What is the current evidence that RNase L functions beyond the interferon system in the more fundamental control of RNA decay? Early indications of a wider role for the 2-5A system arose from the observations that 2-5A synthetase levels increase during cell growth arrest and changing hormone status (Stark et al., 1979; Krishnan and Baglioni, 1980), whereas RNase L levels increase during cell growth arrest and differentiation (Jacobsen et aL, 1983b; Krause et aL, 1985a,b). The increase in RNase L levels in murine JLS-V9R cells during confluency preceded an inhibition in DNA synthesis, suggesting the involvement of RNase L in cell growth control (Jacobsen et al., 1983b). In the same study, the induction of RNase L occurred in the absence of interferon. Hormonal regulation of the 2-5A system was observed in chick oviducts induced to regress by withdrawal of estrogen stimulation. Tissue regression was accompanied by an increase in levels of 2 - 5 A synthetase (Stark et al., 1979) and 2-5A per se and breakdown of 18S rRNA and ovalbumin mRNA (Cohrs et al., 1988). However, the RNA
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degradation that occurred in the oviduct was not rigorously demonstrated to be due to RNase L. Along the same lines of investigation, 2 - 5 A and related oligonucleotides were detected in rat m a m m a r y glands following cessation of lactation (Reid et al., 1984). Also, a decrease in levels of both 2 - 5 A synthetase and of 2 - 5 A per se were observed in regenerating rat liver following partial hepatectomy (Etienne-Smekens et aL, 1983). Patterns of r R N A breakdown in dying cells have led to the suggestion that RNase L is active during apoptosis. H u m a n colon carcinoma cells undergoing cytocidal effects of y-interferon plus tumor necrosis factor, and different cell types undergoing programmed cell death in response to various agents, were observed to produce discrete r R N A cleavage products (Chapekar and Glazer, 1988; Houge et al., 1995). Although RNase L was suggested to be responsible, the endoribonucleases that caused these cleavages in r R N A were not identified. Recently, however, expression of a dominant-negative RNase L mutant in different mammalian cell lines resulted in a marked suppression of apoptotic cell death (J. Castelli, B. Hassel, R. Silverman, and R. Youle, 1996, unpublished data). In addition, a visual demonstration that the 2 - 5 A system results in cell death as a first line of defense against viruses was obtained in transgenic tobacco plants. Infection of plants expressing a human 2 - 5 A system ( 2 - 5 A synthetase and RNase L) by three different types of plant viruses caused local necrotic lesions instead of typical systemic infections (Mitra et al., 1996). The potential involvement in fundamental control of R N A turnover, in particular during programmed cell death, is one of the most intriguing aspects of the currently unfolding biology of RNase L.
ACKNOWLEDGMENTS
I thank Bryan R. G. Williams (Cleveland) for comments made during preparation of the manuscript. This work was supported by United States Public Health Service Grant CA 44059, awarded by the Department of Health and Human Services, National Cancer Institute.
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Agy, M. B., Wambach, M., Foy, K., and Katze, M. G. (1990). Expression of cellular genes in CD4 positive lymphoid cells infected by the human immunodeficiency virus, HIV-
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1: Evidence for a host protein synthesis shut-off induced by cellular RNA degradation. Virology 177, 251-258. Baca, L. M., Genis, P., Kalvakolanu, D., Sen, G., Meltzer, M. S., Zhou, A., Silverman, R., and Gendelman, H. E. (1994). Regulation of interferon-a-inducible cellular genes in human immunodeficiency virus-infected monocytes. J. Leukocyte Biol. 55, 299-309. Baglioni, C., Minks, M. A., and Maroney, P. A. (1978). Interferon action may be mediated by activation of a nuclease by pppA2'p5'A2'p5'A. Nature (London) 273, 684-686. Baglioni, C., De Benedetti, A., and Williams, G. J. (1984). Cleavage of nascent reovirus mRNA by localized activation of the 2'-5'-oligoadenylate-dependent endoribonuclease. J. Virol. 52, 865-871. Ball, L. A., and White, C. N. (1978). Oligonucleotide inhibitor of protein synthesis made in extracts of interferon-treated chick embryo cells: Comparison with the mouse low molecular weight inhibitor. Proc. Natl. Acad. Sci. U.S.A. 75, 1167-1171. Bayard, B. A., and Gabrion, J. B. (1993). 2',5'-Oligoadenylate-dependent RNAase located in nuclei: Biochemical characterization and subcellular distribution of the nuclease in human and murine cells. Biochem. J. 296, 155-160. Bayard, B., Bette-Bobillo, P., and Aliau, S. (1994). Affinity purification and characterization of (2'-5')oligo(adenylate)-dependent RNase from mouse spleen. Eur. J. Biochem. 223, 403-410. Bayliss, C. D., and Condit, R. C. (1993). Temperature-sensitive mutants in the vaccinia virus A18R gene increase double stranded RNA synthesis as a result of aberrant viral transcription. Virology 194, 254-262. Bisbal, C., Salehzada, T., Lebleu, B., and Bayard, B. (1989). Characterization of two murine (2'-5')(A)n-dependent endonucleases of different molecular mass. Eur. J. Biochem. 179, 595-602. Bisbal, C., Martinand, C., Silhol, M., Lebleu, B., and Salehzada, T. (1995). Cloning and characterization of a RNase L inhibitor. A new component of the interferon-regulated 2-5A pathway. J. Biol. Chem. 271), 13308-13317. Blank, V., Kourilsky, P., and Israel, A. (1992). NF-KB and related proteins: Rel/dorsal homologies meet ankyrin-like repeats. Trends in Biochem. Sci. 17, 135-140. Boone, R. F., Parr, R. P., and Moss, B. (1979). Intermolecular duplexes formed from polyadenylated vaccina RNA. J. Virol. 301, 365-374. Bork, P., and Sanders, B. (1993). A hybrid protein kinase-RNase in an interferon-induced pathway? FEBS Lett. 334, 149-152. Breeden, L., and Nasmyth, K. (1987). Similarity between cell-cycle genes of budding yeast and fission yeast and the Notch gene of Drosophila. Nature (London) 329, 651-654. Brown, G. E., Lebleu, B., Kawakita, M., Shaila, S., Sen, G. C., and Lengyel, P. (1976). Increased endonuclease activity in an extract from mouse Ehrlich ascites tumor cells which had been treated with a partially purified interferon preparation: Dependence on double-stranded RNA. Biochem. Biophys. Res. Commun. 69, 114-122. Cayley, P. J., White, R. F., Antoniw, J. F., Walesby, N. J., and Kerr, I. M. (1982a). Distribution of the ppp(A2'p),A-binding protein and interferon-related enzymes in animals, plants, and lower organisms. Biochem. Biophys. Res. Commun. 108, 12431250. Cayley, P. J., Knight, M., and Kerr, I. M. (1982b). Virus-mediated inhibition of the 2-5A system in control cells and its prevention by interferon. Biochem. Biophys. Res. Commun. 104, 376-382.
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Torrence, P. F., Imai, J., and Johnson, M. I. (1981). 5'-O-monophosphoryladenylyl(2' ~ 5')adenylyl(2' ~ 5') adenosine is an antagonist of the action of 5'-0triphosphoryladenylyl(2' ~ 5')adenylyl(2' ~ 5')adenosine and double-stranded RNA. Proc. Natl. Acad. Sci. U.S.A. 78, 5993-5997. Torrence, P. F., Imai, J., Lesiak, K., Jamoulle, J.-C., and Sawai, H. (1984). Oligonucleotide structural parameters that influence binding of 5'-O-triphosphoryladenylyl(2' ~ 5') adenylyl(2' ~ 5')adenosine to the 5'-O-triphosphoryladenyly(2' ~ 5')adenylyl(2' ~ 5')adenosine-dependent endoribonuclease: Chain length, phosphorylation state, and heterocyclic base. J. Med. Chem. 27, 726-733. Torrence, P. F., Brozda, D., Alster, D., Charubala, R., and Pfleiderer, W. (1988). Only one 3'-hydroxyl group of ppp5'A2'p5'A2'p5'A (2-5A) is required for activation of the 2-5A-dependent endonuclease. J. Biol. Chem. 263, 1131-1139. Torrence, P. F., Maitra, R. K., Lesiak, K., Khamnei, S., Zhou, A., and Silverman, R. H. (1993). Targeting RNA for degradation with a 2',5'-oligoadenylate-antisense chimera. Proc. Natl. Acad. Sci. U.S.A. 90, 1300-1304. Torrence, P. F., Xiao, W., Li, G., and Khamnei, S. (1994). Development of 2',5'-oligonucleotides as potential therapeutic agents. Curr. Med. Chem. 1, 176-191. Vilcek, J., and Sen, G. C. (1996). Interferons and other cytokines. In "Fields Virology" 3rd edition (B. N. Fields, ed.), 3rd Ed., 375-399. Raven, New York. Walker, J. E., Saraste, M., Runswick, M. J., and Gay, N. J. (1982). Distantly related sequences in the a- and 13-subunits of ATP synthase, myosin, kinases and other ATPrequiring enzymes and a common nucleotide binding fold. E M B O J. 1, 945-951. Wallach, D., and Revel, M. (1980). An interferon-induced cellular enzyme is incorporated into virions. Nature (London) 287, 68-70. Wang, L., Zhou, A., Vasavada, S., Dong, B., Nie, H., Church, J. M., Williams, B. R. G., Banerjee, S., and Silverman, R. H. (1995). Elevated levels of 2-5A dependent RNase L occur as an early event in colorectal tumorigenesis. Clin. Cancer Res. 1, 1421-1428. Watling, D., Serfinowska, H. T., Reese, C. B., and Kerr, I. M. (1985). Analogue inhibitor of 2-5A action: Effect on the interferon-mediated inhibition of encephalomyocarditis virus replication. E M B O J. 4, 431-436. Williams, B. R. G. (1995). The role of the dsRNA-activated kinase, PKR, in signal transduction. Seminars in Virology 6, 191-202. Williams, B. R. G., and Silverman, R. H. (1985). The 2-5A system: Molecular and clinical aspects of the interferon-regulated pathway. Prog. Clinic. Biol. Res. 202. Williams, B. R. G., Kerr, I. M., Gilbert, C. S., White, C. N., and Ball, L. A. (1978). Synthesis and breakdown of pppA2'p5'A2'p5'A and transient inhibition of protein synthesis in extracts from interferon-treated and control cells. Eur. J. Biochem. 92, 455-562. Williams, B. R. G., Gilbert, C. S., and Kerr, I. M. (1979a). The respective roles of the protein kinase and pppA2'p5'A2'p5'A-activated endoribonuclease in the inhibition of protein synthesis by double stranded RNA in rabbit reticulocyte lysate. Nucleic Acids Res. 6, 1335-1350. Williams, B. R. G., Golgher, R. R., Brown, R. E., Gilbert, C. S., and Kerr, I. M. (1979b). Natural occurrence of 2-5A in interferon-treated EMC virus-infected L cells. Nature (London) 282, 582-586. Williams, G., De Benedetti, A., and Baglioni, C. (1986). Inhibition of 2',5'-oligo(A)dependent endoribonuclease by 2',5'-oligo(A) degradation products. Virology 151, 233-242.
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Wreschner, D. H., and Rechavi, G. (1988). Differential mRNA stability to reticulocyte ribonucleases correlates with 3' non-coding (U),,A sequences. Eur. J. Biochem. 172, 333-340. Wreschner, D. H., James, T. C., Silverman, R. H., and Kerr, I. M. (1981a). Ribosomal RNA cleavage, nuclease activation and 2-5A(ppp(A2'p),,A) in interferon-treated cells. Nucleic Acids Res. 9, 1571-1581. Wreschner, D. H., McCauley, J. W., Skehel, J. J., and Kerr, I. M. (1981b). Interferon action-sequence specificity of the ppp(A2' p)nA-dependent ribonuclease. Nature (London) 289, 414-417. Wreschner, D. H., Silverman, R. H., James, T. C., Gilbert, C. S., and Kerr, I. M. (1982). Affinity labeling and characterization of the ppp(A2'p)nA-dependent endoribonuclease from different mammalian sources. Eur. J. Biochem. 124, 261-268. Wu, Y., Mikulski, S. M., Ardelt, W., Rybak, S. M., and Youle, R. J. (1993). A cytotoxic ribonuclease: Study of the mechanism of onconase cytotoxicity. J. Biol. Chem. 268, 10686-10693. Zhou, A., Hassel, B. A., and Silverman, R. H. (1993). Expression cloning of 2-5Adependent RNAase: A uniquely regulated mediator of interferon action. Cell (Cambridge, Mass.) 72, 753-765.
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17 RNA-Processing RNases in Mammalian Cells JEFF ROSS McArdle Laboratory for Cancer Research Department of Pathology University of Wisconsin--Madison Madison, Wisconsin 53706
I. Introduction II. rRNA-Processing Ribonucleases A. Introduction B. Endonucleolytic Cleavage Sites 0 (A0 in Yeast) and 2 (Site D in Yeast) C. Endonucleolytic Cleavage and "Trimming" at/near Site 1 (Site AI in Yeast) D. Endonucleolytic Cleavage at Site A3 in Yeast by RNasc Mitochondrial RNA Processing E. 5'-Exonucleolytic Trimming of Pre-5.8S(S) RNA by Two Yeast Exoribonucleases F. Cleavage at Sites 4b, 5, and 6, in and around 28s rRNA (Sites C2, C1, and B2, Respectively, in Yeast) G. Perspective III. mRNA-Degrading Ribonucleases A. Introduction B. mRNA Decay Products and Putative mRNase Recognition Sites C. 3'---, 5' Exoribonucleases D. Endoribonucleases E. Perspective IV. Summary References
I. Introduction I n this c h a p t e r t h e f o c u s is o n R N a s e s i n v o l v e d in p r o c e s s i n g r i b o s o m a l R N A ( r R N A ) a n d d e g r a d i n g m e s s e n g e r R N A ( m R N A ) in m a m m a l i a n
RIBONUCLEASES: STRUCTURESAND FUNCTIONS
553 Copyright 9 1997by AcademicPress, Inc. All rights of reproduction in any form reserved.
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cells. Where relevant, RNases from other organisms are mentioned, and, because yeast mutants have provided so much useful information about rRNA processing, yeast rRNA-processing RNases are discussed in some detail. The rRNA- and mRNA-processing/degrading enzymes are particularly interesting because of their efficiency and specificity. At each division the average mammalian cell must produce approximately 2 x 106 ribosomes from 50 to 1000 genes, and apparently does so with high efficiency, especially if it is growing exponentially (reviewed in Eichler and Craig, 1994). Each rRNA molecule (18S, 5.8S, and 28S) must be precisely cleaved from a 47S precursor. Similarly, mRNA-degrading enzymes distinguish among thousands of mRNAs whose half-lives range from a few minutes to days (reviewed in Ross, 1995; Beelman and Parker, 1995). Thus, one of the most intriguing questions about these RNases is how they recognize their substrates. How do the rRNA-processing enzymes attack such a large target at only one or a few sites? How do a presumably limited number of mRNases decide which mRNAs to degrade? As will be discussed, a combination of f a c t o r s n R N A structure, RNA-binding proteins, and proximity of enzyme to substratenappears to play an important role in determining RNase specificity and efficiency.
II. r R N A - P R O C E S S I N G R I B O N U C L E A S E S
A. Introduction
The rRNA genes in many organisms are tandemly repeated 50-1000 times and generate precursor RNAs ranging from approximately 35S (yeast) to 47S (mammals) (reviewed in Eichler and Craig, 1994) (Fig. 1). RNase cleavages proceed in more or less a 5' ~ 3' direction using a combination of endonucleases and exonucleases to generate the mature 18S, 5.8S, and 28S rRNAs. Both external and internal transcribed spacer regions (ETS and ITS, respectively) are precisely and efficiently removed and degraded. Several observations imply that processing of both large and small ribosomal subunit rRNAs occurs semiindependently. For example, small nucleolar ribonucleoproteins (snoRNPs) play essential roles in rRNA processing, and mutations in yeast genes encoding some snoRNPs, for example, U3, prevent the production of 18S but not of 5.8S and 25S rRNAs (reviewed in Eichler and Craig, 1994; Morrisey and Tollervey, 1995).
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Fig. 1. Diagram of (a) human 47S and (b) yeast 35S pre-rRNAs. Lines indicate the external transcribed sequences (ETS) that are excised and degraded. Boxes indicate the retained rRNA sequences.
Some but not all of the processing RNases have been identified and will be described here, and it will be evident from this discussion that many pieces of the mammalian rRNA-processing "puzzle" are known but have not been fully put into place. Some yeast cell enzymes are also discussed, because several of them, including the 5'--* 3' exoribonucleases and RNase MRP, are likely to be homologous to enzymes required for processing mammalian rRNA. Each known cleavage site in the primary transcript is designated with a name and/or number but, unfortunately, there are no uniform designations for similar sites in mammals and yeast. Therefore, the diagrams of each rRNA cleavage site indicate both.
B. Endonucleolytic Cleavage Sites 0 (A0 in Yeast) and 2 (Site D in Yeast) As shown in Diagram 1, the early or initial endonucleolytic cleavages are catalyzed by a magnesium-dependent enzyme at site 0, which is located 414 to 419 nucleotides 3' of the 5' terminus of the human 47S
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Diagram 1.
rRNA precursor (+650-656 nt of the mouse precursor; Fig. 2), at site 2, which is located at the 3' terminus of the 18S sequence, and at a third site (not shown), which is located within the internal transcribed spacer 1 (ITS1). When a segment of rDNA including the mouse +650 site is incubated in an $100 transcription/processing extract from mouse tissue culture cells, the RNA is synthesized and rapidly cleaved at both the +650 and +656 sites (Kass et al., 1987). A candidate enzyme responsible for these cleavages was first isolated and characterized by Eichler and Eales (1982). In cell-free reactions containing protein-free rRNA transcribed from the 5' external transcribed spacer (ETS) and including the +650 site, the purified enzyme cleaves at +650 (Shumgard and Eichler, 1988; Shumgard et al., 1990). Using a substrate from the 3' segment of 18S rRNA plus the 5' segment of ITS1, the purified enzyme cleaves at three sites, a major one at the mature 18S 3' end, one at approximately 35 nucleotides 5' of the 18S 3' end, and one approximately 55 nucleotides downstream of the 18S 3' end (Shumgard and Eichler, 1988; Shumgard et al., 1990). The enzyme requires magnesium and has a native molecular mass of approximately 50 kDa (Eichler et al., 1993). The sequence requirements for the +650 cleavage have been investigated in the coupled transcription/processing $100 system. The 5' boundary of the recognition site is quite sharp and includes essentially the cleavage site itself (Craig et al., 1987). The 3' site is not as sharply defined but includes nucleotides 650-875 (Craig et al., 1991). This region is conserved among mammals, from humans to amphibians, consistent with it being a processing signal site. It also forms extensive secondary structure, and cleavages occur within a loop (Fig. 2). Prehybridizing the loop region with antisense RNA interferes with cleavage, confirming that the single-stranded loop is necessary (Craig et aL, 1991).
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C
U
650
C 9A
C
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C G G U U U G C C G C G
A
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Fig. 2. Cleavage sites in the +650 region of the mouse rRNA precursor, providing the sequence and proposed secondary structure of the 5' ETS of the human pre-rRNA segment that is cleaved by the rRNA-processing endoribonuclease (described in Section II,B).
In summary, the RNase responsible for the human +414 cleavage (+650 in mouse) recognizes secondary and tertiary structure in the substrate. However, additional work is required to understand substrate specificity in detail. The highly purified ---50-kDa enzyme is able to bind at or close to the cleavage site in reactions lacking magnesium, which is essential for cleavage (Eichler et al., 1993). This important finding illustrates that the RNase can, on its own, recognize its substrate. On the other hand, the enzyme probably does not function on its own in cells and needs additional factors, including the snoRNPs (reviewed in Eichler and Craig, 1994; Morissey and Tollervey, 1995). For example, as determined by psoralen treatment and UV cross-linking of yeast preribosomal RNA (pre-rRNA), U3 binds just upstream of a highly conserved 11-nucleotide segment located downstream of the A0 cleavage site (Beltrame and Tollervey, 1992). Moreover, the genes encoding the
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yeast U3 snoRNP are absolutely required for cleavage at site A0 (Morissey and Tollervey, 1995), and depletion of U3 snoRNP from mouse cell S100 extracts by oligonucleotide-directed RNase H action inhibits +650 cleavage (Kass et al., 1990). These results are consistent with U3 (and other) snoRNPs having a significant role in the +650 cleavage. Perhaps a group of snoRNPs and the endonuclease form a "cleavosome" complex whose purpose is to position the RNase such that it cleaves only at a few selected sites (Eichler and Craig, 1994). Some sort of positioning of the RNase might be required, because sufficient amounts of the purified endonuclease can cleave protein-free mouse rRNA at sites other than +650 (Eichler et al., 1993).
C. Endonucleolytic Cleavage and "Trimming" at/near Site 1 (Site A1 in Yeast) When a 260-nucleotide human pre-rRNA substrate, including part of the 5' ETS plus the 5' segment of 18S rRNA, is incubated in a HeLa cell nucleolar extract in the presence of KCI and MgCI2, a two-step reaction, depicted in Diagram 2, generates RNA with an authentic 18S 5' terminus (Hannon et al., 1989). First, an endonuclease activity cleaves the substrate at three sites at - 8 , - 3 , and +1, relative to the 18S 5' end. The 5' terminus of each reaction product is hydroxylated. Then, a "trimming activity" found in the cytoplasm clips the - 3 (but not the
Diagram 2.
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- 8 ) RNA. Trimming requires a substrate with a 5'-hydroxyl group, consistent with the structure of the endonucleolytic cleavage product, and it actually generates two products, 5'-pCUACC and 5'-pUACC, the latter being the authentic 18S 5' terminus (Yu and Nilsen, 1992). It is unclear whether trimming activity is located in the cytoplasm or leaks into the cytoplasmic extract during fractionation. As determined by mutagenesis and modification interference assays, nucleotides +6 to +25 (i.e., within the 18S rRNA) are important for accurate trimming. Obviously, it will be important to purify and further characterize both enzymes, in order to assess how such specificity for the 18S 5' terminus is achieved.
D. Endonucleolytic Cleavage at Site A3 in Yeast by RNase Mitochondrial RNA Processing RNase mitochondrial RNA processing (MRP) is a ribonucleoprotein endoribonuclease that is required to produce the major form of 5.8S rRNA. The 260- to 280-nucleotide RNA component of MRP, called 7-2 RNA, is generated from a single-copy nuclear gene transcribed by RNA polymerase III; the 100-kDa protein component is similar or identical to the protein of RNase P (reviewed in Gold et al., 1989; Morrissey and Tollervey, 1995; Lygerou et al., 1994). RNase MRP was first identified and purified from mouse cell mitochondria by virtue of its ability in vitro to cleave RNA primers involved in mitochondrial DNA replication (Chang and Clayton, 1987a,b, 1989). It is magnesium dependent and inactive with calcium or manganese; it is inhibited by N-ethylmaleimide and by salt concentrations above 100 mM, and produces RNAs with 5'phosphoryl and 3'-hydroxyl ends. Over 90% of the RNA and protein components of RNase MRP is located in nucleoli, and highly purified mitochondria contain little, if any, 7-2 RNA (Kiss and Filipowicz, 1992), suggesting that the major function of MRP involves rRNA processing (Yuan et al., 1989; Reimer et al., 1988). Based primarily on studies in yeast, it is now clear that RNase MRP is responsible for producing the smaller of the two forms of 5.8S RNA, called 5.8S(S). RNA 5.8S(S) (Diagram 3) is the predominant form in cells and is 7 to 8 nucleotides shorter at its 5' end compared to the minor form, designated 5.8S(L). Because mammalian cells also contain nucleolar-localized RNase MRP,
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Diagram 3.
it seems reasonable to assume that this RNA is responsible for similar cleavages at site 2B in mammalian pre-rRNA. The yeast 35S pre-rRNA molecule is first cleaved at site A0 (site 0 in mammals), generating 32S RNA, which is then cleaved at site A2 in ITS1 by an unknown endonuclease (see Diagram 3). This cleavage separates the 32S precursor into two RNAs, 20S, which will become 18S rRNA, and 27S, which will become 5.8 and 25S rRNAs. The 20S precursor is then cleaved by RNase MRP in ITS1, 76 nucleotides 5' to the mature 5.8S(S) 5' terminus (Henry et al., 1994). Three observations demonstrate that RNase MRP is required for 5.8S(S) rRNA production: (1) Yeast strains with mutations in the N M E 1 gene, which encodes the RNA component of MRP RNase, accumulate normal amounts of 5.8S(L) R N A but little 5.8S(S) RNA (Shuai and Warner, 1991; Lindahl et aL, 1992; Schmitt and Clayton, 1993; Chu et al., 1994). (2) In yeast mutant strains depleted of snoRNP components but containing RNase MRP, an rRNA precursor with a 5' terminus at the A3 site is readily detected (Henry et al., 1994). This demonstrates that site A3 is used in cells. (3) In strains lacking functional RNase MRP, no A3 site-cleaved R N A is
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detected, but a precursor cleaved at site A2 is readily observed (Henry et aL, 1994; Lygerou et al., 1994). Moreover, cleavage at site A3 is reduced when a 10-nucleotide deletion is made at the A3 site itself. It is not clear how the RNase recognizes the A3 site, which is particularly (A + C)rich, being 5 ' - A A C A A / A C A C A C A A A C A A - 3 ' (Henry et aL, 1994). It is also intriguing that the protein components of RNases MRP and P are identical in yeast cells (Lygerou et al., 1994) and probably in mammalian cells as well, because sera from patients with autoimmune diseases immunoprecipitate both proteins (Gold et al., 1989). It will be important to determine how the same protein binds to two RNAs (in RNases MRP and P) and whether the RNAs or the protein are in limiting amounts in cells.
E. 5'-Exonucleolytic Trimming of Pre-5.8S(S) R N A by Two Yeast Exoribonucleases
One or both 5' ---~ 3' exoribonucleases encoded by the H K E I / R A T I and X R N I / K E M I genes are required to cleave the 76 nucleotides from the 5' end of yeast pre-5.8S(S) RNA (Henry et al., 1994) (see Diagram 3). Apparently, these genes have multiple functions, because mutations in them have pleiotropic effects involving, among other processes, DNA replication and RNA metabolism (reviewed in Larimer et al., 1992). In temperature-sensitive mutant strains cultured at the nonpermissive temperature, the levvel of 5.8S(S) RNA is decreased, and an elongated 5.8S RNA whose 5' terminus maps to the A3 cleavage site accumulates (Stevens et al., 1991; Amberg et al., 1992; Henry et al., 1994). X R N 1 and H K E 1 encode 175- and ll6-kDa proteins, respectively, both of which are 5'--0 3' exoribonucleases in vitro (Larimer et al., 1992; Kenna et al., 1993). They are magnesium-dependent, are unable to degrade capped RNA, generate nucleoside 5'-monophosphates (Stevens, 1978), and play a central role in the degradation of many yeast mRNAs (reviewed in Beelman and Parker, 1995). It is unclear how they recognize pre-5.8S(S) RNA or how they stop at the precise 5' terminus of the mature RNA, which is particularly intriguing, because they apparently degrade most or all of each susceptible (uncapped) mRNA molecule they attack. Perhaps 5.8S(S) rRNA-binding proteins block further attack by the exonucleases. Although mammalian homologs of the yeast exoribonucleases have not been unequivocally identified, a nucleolar
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5' ~ 3' exoribonuclease has been purified from Ehrlich ascites tumor cells (Lasater and Eichler, 1984). This magnesium-dependent, 76-kDa enzyme degrades single-stranded R N A with either 5'-OH or 5'-phosphate groups, releasing 5'-mononucleotides. It is not inhibited by the RNase A inhibitors, and it cannot degrade duplex RNA or single- or doublestranded DNA. It is an obvious candidate for the 5' trimming activity of 5.8S(S) RNA in mammalian cells.
F. Cleavage at Sites 4b, 5, and 6, in and around 28S rRNA (Sites C2, C1, and B2, respectively, in Yeast) Little is known about the enzymes responsible for generating 28S rRNA. Termination of rDNA gene transcription occurs 565 nucleotides downstream of the mouse 28S 3' terminus (reviewed in Grummt et al., 1985; Bartsch et al., 1988) and 15 to 40 nucleotides downstream of the yeast 25S 3' terminus (reviewed in Yip and Holland, 1989). In crude transcription/processing extracts from mouse (Kuhn and Grummt, 1989) and yeast (Yip and Holland, 1989) cells, pre-rRNAs are cleaved by an unknown RNase soon after they are synthesized. Ten nucleotides are cleaved from the 3' end of the mouse primary transcript (Kuhn and Grummt, 1989) (cleavage site designated 6' in the Diagram 4), and the cleavage rate is reduced if a cluster of U residues near the transcription termination site is changed to G residues (Kuhn and Grummt, 1989). In crude extracts from the yeast Saccharomyces cerevisiae, the 3' ETS region undergoes three cleavages, two near the 3' end of the 35S precursor and another at the authentic 25S 3' terminus (Yip and Holland, 1989). The responsible RNase, presumably an endonuclease, might be encoded by the RNA 82.1 gene in yeast, because mutant strains with a defective 82.1 gene fail to process both pre-rRNA and pre-5.8S rRNA (Kempers-Veenstra et aL, 1986; Piper et al., 1983). It is unknown whether
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the authentic 25S 3' end (or the 3' termini of 18 and 5.8S RNAs, for that matter) is generated only by the RNA 82.1 gene product or by a combination of an endonuclease plus a 3 ' ~ 5' exonuclease. A 100kDa magnesium- or manganese-requiring, distributive nucleolar 3' 5' exonuclease that releases 5'-mononucleotides has been purified from Ehrlich ascites tumor cells and is a candidate trimming enzyme (Eichler and Eales, 1985). An endonucleolytic cleavage at site 4b within ITS2 generates 5.8S and 28S pre-rRNAs extended at their 3' and 5' termini, respectively (Bowman et aL, 1983). The subsequent trimming reactions are catalyzed by uncharacterized enzymes, but the nucleolar exonuclease is certainly a candidate.
G. Perspective As noted above, considerable progress has been made in unraveling the puzzle of how mammalian pre-rRNA is processed. The paucity of genetic approaches, so rewarding in yeast systems, has been an impediment to still faster progress. However, many features of yeast and mammalian rRNA processing, including the pathways and some of the enzymes (RNase MRP, for example), seem quite similar. Therefore, future progress will probably follow on two parallel approaches. First, mammalian processing RNases will continue to be purified and then cloned, and their functions in rRNA biogenesis will be tested by their capacity to complement yeast mutations. Second, more sophisticated in vitro rRNAprocessing systems will continue to be developed. As more of the RNases and cofactors are purified, these systems will be used to reconstitute individual processing reactions.
III. m R N A - D E G R A D I N G
RIBONUCLEASES
A. Introduction No mammalian cell RNase devoted exclusively to degrading m R N A has yet been unequivocally identified, for several reasons. First, mammalian cells contain multiple RNases with little or no apparent specificity
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for any RNA. In cell extracts these enzymes, like the bacterial and fungal RNases used in the laboratory, degrade most or all deproteinized R N A substrates. As a result, there exists a high nonspecific "background" of RNase activity in lysates of mammalian cells, making it difficult to distinguish biochemically mRNases from other RNases. Second, mammalian cell mutants with defects in m R N A degradation have not been isolated. In lower organisms such mutant strains have been extremely useful for identifying mRNases and characterizing m R N A decay pathways (reviewed in Deutscher, 1993; Carpousis et al., 1994). These limitations have meant that some basic questions about mammalian mRNases remain unanswered. Are they constitutive RNases with little or no specificity for specific RNA sequences? If so, why are some mRNAs so much more stable than others (reviewed in Ross, 1995)? Are m R N A halflives determined more by the structure of the substrate (mRNA tertiary structure, m R N A - p r o t e i n interactions, etc.) than by the activities of mRNases? Some answers are forthcoming because of progress in purifying putative mRNases and identifying m R N A decay pathways in vitro and in intact cells. Therefore, mammalian mRNases are discussed in two sections. The first deals with the identification and mapping of m R N A decay products in intact cells and cell-free extracts. The structures of decay intermediates provide important information about the enzymes involved in the degradation process. The second deals with RNases that have been partially or completely purified and whose properties are consistent with their being mRNases.
B. m R N A Decay Products and Putative mRNase Recognition Sites It is beyond the scope of this review to describe in detail the various motifs thought to be important as determinants of m R N A half-life; they have been reviewed elsewhere (Sachs, 1993; Beelman and Parker, 1995; Ross, 1995). However, it is important to highlight some principles to serve as a guide for emphasizing areas of ignorance. To the best of our knowledge, no RNase has been described whose properties are exactly comparable to those of restriction endodeoxyribonucleases. No known RNase recognizes a short sequence with the same specificity as, say, E c o R I recognizes the sequence GAAT-FC. Nevertheless, certain sequences/motifs, many of which are located in the 3' untranslated
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region (3'-UT), do play important roles in determining mRNA half-life, presumably as preferential sites for recognition of either the mRNases themselves or other factors (RNA-binding proteins) that augment mRNase activity, either positively or negatively. Poly(A) is clearly a major determinant of mRNase activity, and two observations implicate poly(A) as a shield against rapid and indiscriminate attack by cell RNases. First, although many mammalian mRNAs are degraded endonucleolytically (Sections III,B and Ill,D), deadenylation is the first step in the decay pathway of other mRNAs (reviewed in Mercer and Wake, 1985; Bernstein and Ross, 1989; Herrick et al., 1990; Sachs, 1990) (Fig. 3). Second, the poly(A)-poly(A)-binding protein (PABP) complex at the mRNA 3' terminus protects mammalian mRNAs from rapid destruction in vitro (Bernstein et al., 1989). These observations imply that the susceptibility of poly(A) to RNase(s) is determined to a major extent by the stability of the poly(A)-PABP complex. Regardless of their half-lives, most mRNAs begin their cytoplasmic existence with a 100- to 200-nucleotide poly(A) tail. In many cases, these mRNAs are deadenylated at rates that reflect their half-lives (Swartwout et al., 1987; Brewer and Ross, 1988; Swartwout and Kinniburgh, 1989; Shyu et al., 1989, 1991). Therefore, something besides poly(A) must determine the deadenylation rate, because one poly(A) tail is identical to the next. RNA secondary structure also plays a major role in mRNA stability, and stem-loop motifs in mRNA 3'-UT's are often primary determinants of mRNA half-life. The 3'-terminal stem-loop of histone mRNA (Fig. 3) is a site of exonuclease attack and of binding of regulatory proteins (Melin et al., 1992; Pandey et al., 1991, 1994; Section Ill,C). The mRNAs encoding transferrin receptor and ferritin, both of which affect iron homeostasis, are regulated posttranscriptionally by processes dependent on the intracellular iron concentration. These regulatory processes are effected by an iron-response element (IRE), a 23- to 27-base pair stem with a mismatched C and a 6-nucleotide loop with C at its 5' end (reviewed in Casey et al., 1989; M011ner and K0hn, 1988; Klausner et al., 1993; Harford et al., 1994) (Fig. 3). The IRE functions by binding an iron regulatory protein whose binding affinity, in turn, is determined by intracellular iron abundance. A remarkable stem-loop structure in the 3'-UT of human, mouse, and rat insulin-like growth factor II (IGF-II) mRNAs is both an endonuclease cleavage site and a mRNA stability determinant (Meinsma et al., 1991, 1992; Nielsen and Christiansen, 1992; Christiansen et al., 1994; Scheper et al., 1995). Two observations link the AU-rich elements (AUREs) located in 3'UT's with mRNA instability. First, mRNAs whose 3'-UTs contain an
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A U R E and/or an oligo(U) region tend to be unstable (Caput et al., 1986). Second, if an A U R E from the 3 ' - U T of an unstable m R N A is placed within the 3 ' - U T of a stable m R N A , the chimeric transcript decays with a half-life of less than 30 min (Shaw and Kamen, 1986; B o h j a n e n et al., 1992; Alberta et al., 1994; Chen et al., 1994; Chen and Shyu, 1994). The coding region can play a major role in determining m R N A halflife, as illustrated by three observations: (1) Mutations in the coding region of m R N A s such as c-los, c-myc, and tubulin can result in significant changes in m R N A half-life (Shyu et aL, 1989, 1991; Wisdom and Lee, 1990, 1991; Bachurski et al., 1994; Herrick and Ross, 1994; Schiavi et aL, 1994; Lavenu et al., 1995). Some coding region stability determinants are also protein-binding sites (Bernstein et al., 1992; Chen et aL, 1992). (2) The half-life of c - m y c and c-los m R N A s lacking most of their 3'UTs and lacking the A U R E is only 1 to 2 hr, which is still relatively short as compared with many other m R N A s (Fort et al., 1987; Kabnick and H o u s m a n , 1988; Lachman et aL, 1986). Therefore, the truncated m R N A s contain an instability determinant, which, by default, must be in the 5 ' - U T and/or coding region. (3) For most or all m R N A s thus far investigated, the introduction of a nonsense mutation in the 5' portion of the coding region destabilizes the m R N A (reviewed in Peltz et al., 1994). It is not clear whether 5'-UTs are major determinants of m R N A stability, but they can affect m R N A half-life (reviewed in Ross, 1995). In summary, various "motifs" influence the activities of mRNases. However, it is important to note that m R N A degradation, like r R N A processing, is compartmentalized, probably occurs on substrates that are
Fig. 3. Three mRNA decay pathways in mammalian cells. Top: Poly(A) shortening occurs in a 3' --~ 5' direction, but the responsible nuclease(s) have not been identified (Section III,B). Middle: Histone mRNA is degraded 3' --~ 5' by an exoribonuclease that excises nucleoside 5'-monophosphates (Section III,C). The 6-base pair stem and 4-base loop present in all cell cycle-regulated histone mRNAs are indicated, as are two degradation intermediates that lack approximately 5 or 12 of the 3'-terminal nucleotides. These decay intermediates are thought to arise because RNA-binding proteins temporarily bind to and block decay of the 3' region. As a result, the exonuclease is forced to pause on its way 5'. Bottom: Examples of endoribonucleolytic cleavages in three mRNAs (Section III,D). Arrows indicate the regions of cleavage. An AU-rich region in the 3'-UT of c-myc mRNA is shown (AU), the significance of which is that it indicates the existence of two mRNA stability-determining regions (the coding region determinant and the AU-rich region) in the same mRNA.
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ribonucleoproteins, not RNAs, and is likely to be influenced by many factors besides those mentioned above. Most cellular mRNAs are not freely floating in the cytoplasm but are bound to ribosomes and are being translated. Thus, polysome structure and the rate of translation initiation and elongation probably influence the accessibility and susceptibility of many mRNAs to mRNases (reviewed in Peltz et al., 1991, 1994; Belgrader et al., 1993; Sachs, 1993; Beelman and Parker, 1995). For example, most mRNAs are stabilized in cells exposed to translation inhibitors, but different mRNAs might be stabilized for different reasons, and few mechanistic details are known (reviewed in Ross, 1995).
C. 3' --~ 5' Exoribonucleases
There is compelling evidence that 3' ~ 5' exoribonucleases are involved in mammalian mRNA turnover. Both poly(A) shortening and histone mRNA degradation proceed in a 3 ' ~ 5' direction (Mercer and Wake, 1985; Treisman, 1985; Ross and Kobs, 1986; Ross et al., 1986; Fort etal., 1987; Swartwout etal., 1987; Brewer and Ross, 1988; Swartwout and Kinniburgh, 1989; Peppel and Baglioni, 1991; Shyu et al., 1989, 1991). Two observations demonstrate the 3' ~ 5' decay pathway of histone mRNA. (1) Cell cycle-regulated histone mRNAs are abundant during S but scarce during G1, and the signals responsible for the posttranscriptional regulation of histone mRNA abundance and stability through the cell cycle are located at the mRNA 3' terminus (reviewed in Schumperli, 1988; Osley, 1991; Marzluff, 1992). All of these mRNAs lack poly(A) but contain instead a 6-base pair stem and a 4-base loop located close to their 3' termini (reviewed in Marzluff, 1992). The stem-loop motif is widely conserved and is apparently essential for destabilizing the mRNA at the end of S phase. Thus, a chimeric mRNA containing only the 3'-terminal 30 nucleotides of histone mRNA appended to globin mRNA is regulated postranscriptionally as if it were wild-type histone mRNA (Luscher et al., 1985; Levine et al., 1987; Pandey and Marzluff, 1987). (2) mRNA decay intermediates truncated at their 3' ends have been observed both in cell-free mRNA decay reactions and in intact cells (Ross and Kobs, 1986; Ross et al., 1986; Peltz and Ross, 1987; McLaren and Ross, 1993) (Fig. 3). Cell-free extracts have been exploited to attempt to isolate and characterize the RNase responsible for degrading histone mRNA (reviewed
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in Ross, 1994). In crude extracts, polysome-associated histone m R N A is degraded by a magnesium-dependent exoribonuclease that is active from 0 to 250 mM salt and is insensitive to the RNase A inhibitor (Ross and Kobs, 1986; Peltz et al., 1987; Ross et al., 1987). Several exoribonucleases with these properties have been solubilized by highsalt extraction of polysomes, and one has been purified to homogeneity (Caruccio and Ross, 1994). It is a 33-kDa, divalent cation (Mg2+) dependent protein that, in vitro, degrades single-stranded RNA (histone and many others) and double-stranded RNA in the absence of ATP but does not degrade DNA. It also accelerates the degradation of polysomeassociated histone mRNA when added back to cell-free mRNA decay reactions, and it degrades protein-free poly(A) but not poly(A) in a complex with PABP. The reaction products are nucleoside 5'-monophosphates. Therefore, its properties are indistinguishable from those of the activity that degrades polysomal histone mRNA in vitro. If the purified exoribonuclease is an authentic mRNase, its in vitro substrate specificity suggests that it is not restricted to degrading histone mRNA but might degrade any mRNA. Because it can also degrade poly(A), this exonuclease might be involved in deadenylation. If so, it seems particularly interesting that the RNase fails to degrade poly(A) in a poly(A)-PABP complex, consistent with the notion that the PABP protects mRNA from rapid destruction in cells. Several other mammalian poly(A)-degrading 3 ' - + 5' exoribonucleases have been described, but they are primarily nuclear and are therefore unlikely to carry out cytoplasmic mRNA degradation (Astr6m et al., 1991, 1992). A PABP-dependent nuclease appears to catalyze mRNA deadenylation in yeast (Lowell et al., 1992; Sachs and Deardorff, 1992). We know of no evidence that mammalian mRNAs are degraded 5' to 3', although one or more 5'--+ 3' exonucleases degrade at least some mRNAs in yeast (Hsu and Stevens, 1993; Muhlrad et al., 1994, 1995). A cytoplasmic 5' ~ 3' exonuclease activity of unknown function has been identified in extracts from cultured mammalian cells (Coutts and Brawerman, 1993). A related activity has been purified from the postpolysomal ($100) fraction of reticulocytes (Smoske6y et al., 1995). It is a processive 5' ~ 3' exoribonuclease consisting of three bands of 54, 58, and 62 kDa in a SDS gel. Using capped mRNA as a substrate, the activity first cleaves between N-2 and N-3 of the sequence 5'-GpppGN1 N2 N3 N4 . . . . 3' and then proceeds 5' to 3', generating nucleoside 5'-monophosphates. It is inhibited by millimolar concentrations of ATP and by the cap-
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binding complex eIF-4F. It is unclear why or how ATP inhibits the enzyme, but eIF-4F presumably shields and protects the substrate. The functions of these intriguing enzymes are unknown. However, their very existence, coupled with the evidence that some yeast mRNAs are degraded by 5' ~ 3' exoribonucleases, should prompt a renewed effort to detect 5' --~ 3' mRNA degradation pathways in mammalian cells.
D. Endoribonucleases
Putative endonuclease decay products have been identified for the following mRNAs: 9E3 and gro a (involved in regulating cell growth) (Stoeckle, 1991, 1992; Stoeckle and Guan, 1993), transferrin receptor (Binder et al., 1994), monocyte-derived neutrophil-activating peptide (MONAP; an inflammatory protein synthesized by monocytes) (Kowalski and Denhardt, 1989), apolipoprotein (Bakker et al., 1988; Binder et al., 1989; Cochrane and Deeley, 1989; Ratnasabapathy et al., 1990), IGFII (Meinsma et al., 1991, 1992; Nielsen and Christiansen, 1992; Scheper et al., 1995), albumin (Tharun and Sirdeshmukh, 1995), and preproenkephalin (LaForge et al., 1995). In some cases, both the 5' and 3' segments of the mRNA have been detected, providing definitive evidence for sitespecific endonucleolytic cleavage. It is difficult to detect the intracellular decay products of most mRNAs in mammalian cells, presumably because they are so short-lived, and there is no obvious clue as to why the decay products of these mRNAs are more stable than usual. Perhaps they are protected by proteins or form RNase-resistant duplex structures. The RNases responsible for cleaving gro a, apolipoprotein, MONAP, albumin, preproenkephalin, and transferrin receptor mRNAs have not been identified in cell-free exxtracts, but the nature of the decay intermediates is instructive. The 1.3-kb gro a mRNA is very unstable in growing cells (half-life probably less than 30 min) but is stabilized at least eightfold by interleukin-1 (IL-1). If IL-1 is added to cells for several hours and then removed, the mRNA begins to be degraded at an accelerated rate, and a nonpolyadenylated 0.9-kb decay product appears (Stoeckle, 1992). This fragment is generated by removal of 300 to 400 nucleotides from the 3'-UT and would represent the 5' segment of an endonucleolytic cleavage. Because the downstream (3') segment has not been detected, the responsible enzyme could conceivably be an exoribonuclease that
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begins cleaving at the 3' terminus and pauses within the -300- to -400nucleotide region. However, no intermediate degradation products (less than 1300 but greater than 900 nucleotides) have been detected, implying that the 0.9-kb product is generated endonucleolytically. Moreover, 9E3 mRNA, which is related to gro a, is degraded endonucleolytically, because two 9E3 decay products indicative of endonucleolytic cleavage are observed (Stoeckle and Hanafusa, 1989). Transferrin receptor mRNA decay products have been analyzed in cells transfected with a transferrin receptor gene encoding a constitutively unstable mRNA lacking the 5'-most C of the IRE loop. Using S1 nuclease mapping and primer exxtension assays, two degradation products have been detected, and their structures are consistent with endonucleolytic cleavage in the region noted by the arrow in Fig. 3 (Binder et al., 1994). The 3' degradation product is polyadenylated, indicating that cleavage occurs without prior deadenylation. At least part of the endonuclease recognition site is sequence specific, because changing the sequence in that region from G A A C A A G to CCCCCCC blocks cleavage. An endoribonuclease activity that degrades deproteinized IGF-II mRNA has been identified in crude postpolysomal supernatant (S100) of a rat liver cell line (Nielsen and Christiansen, 1992). The RNase activity cleaves the substrate in a loop located within the long-range stem-loop formed in the mRNA 3'-UT (Fig. 3) and near a G-rich region that could form either a G - G (syn-anti) duplex or a G quadriplex (Christiansen et al., 1994). The specificity of the in vitro cleavage activity is striking, because the unusual long-range stem-loop of IGF-II is an apparent signal for mRNA degradation in intact cells (Meinsma et al., 1992; Scheper et al., 1995). Not all endonucleolytic cleavages occur within 3'-UTs. Other endoribonuclease activities have been described that attack mRNA coding regions. For example, to assess how premature translation stop codons reduce mRNA levels, transgenic mice carrying either of four human fl0_ thalassemic globin genes were generated (Lim et al., 1989, 1992; Lim and Maquat, 1992). (Humans homozygous for thalassemia often suffer from anemia, and the erythrocytes of those with fl~ contain no fl-globin. The fl~ genes used in these exxperiments each had a nonsense mutation.) Erythroid cells from these mice, but not from those carrying wild-type human globin genes, contain both full-length and 5'-truncated globin mRNAs. The truncated mRNAs are polyadenylated and apparently arise in the cytoplasm (Lim et al., 1989, 1992;
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Lim and Maquat, 1992). Surprisingly, their 5' termini are capped and are located within the coding region, but not at the premature translation stop sites. The mechanism by which these intriguing intermediates arise is unclear but is presumed to be endonucleolytic. Polysome-associated c - m y c mRNA is cleaved endonucleolytically in cell-free extracts following the addition of excess competitor RNA corresponding to a segment of the mRNA called the coding region determinant (Bernstein et al., 1992) (Fig. 3). This region encodes the C-terminal 60 amino acids of the c - m y c protein and affects mRNA metabolism in intact cells (Wisdom and Lee, 1990, 1991; Herrick and Ross, 1994). Cleavage occurs near the middle of the determinant by an endoribonuclease activity that is tightly bound to polysomes, is magnesium dependent, and is unaffected by the RNase A inhibitor (Bernstein et al., 1992). A protein that binds to the c - m y c mRNA coding region determinant has been purified (Prokipcak et al., 1994), suggesting that the competitor RNA might function by removing the protein from the coding region determinant, thereby exposing the determinant to the RNase. If so, these data would provide direct evidence for alternate c - m y c mRNA decay pathways, presumably by different mRNases. Endonucleolytic cleavage of c - m y c mRNA has been observed only in reactions supplemented with competitor RNA. In unsupplemented reactions, the mRNA is first deadenylated and then degraded in a 3'--* 5' direction (Brewer and Ross, 1988, 1989). Perhaps this unstable mRNA is degraded by the 3' 5' (default) pathway in most cells under normal cell growth conditions. The endonucleolytic pathway might be activated only under special circumstances, as when cells are starved for nutrients or induced to differentiate (Swartwout et al., 1987; Swartwout and Kinniburgh, 1989). If so, it will be important to purify the RNases responsible for each pathway and determine how they are regulated. As discussed above, no endoribonucleases with specificity similar to restriction enzymes have been identified. However, some evidence indicates that putative endonucleolytic mRNases can degrade deproteinized mRNA substrates at different rates corresponding somewhat to their intracellular half-lives. For example, IL-2 mRNA is at least 10-fold less stable than globin mRNA in cells (Lindsten et al., 1989; Ross and Pizarro, 1983), and a soluble (postpolysomal or S130) ---68-kDa RNase in extracts from a human T cell line degrades deproteinized IL-2 mRNA faster than it degrades globin mRNA (Hua et aL, 1993). The enzyme is magnesium dependent and is unaffected by the RNase A inhibitor. It will be interesting to know what sequences the enzyme recognizes and whether the
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enzyme is regulated during T cell activation. A soluble RNase activity from reticulocytes rapidly degrades an IL-1 m R N A substrate containing four or five clustered A U U U A sequences (Gorospe and Baglioni, 1994; see also Wreschner and Rechavi, 1988; Zhang et al., 1993). The proximity of the A U U U A pentamers is apparently crucial, because RNA with four or five clustered A U U U A s is degraded more rapidly than RNA with four or five dispersed AUUUAs. An as yet uncharacterized activity in reticulocyte lysates degrades a high molecular weight (7-kb) isoform of deproteinized insulin-like growth factor I m R N A faster than a 1-kb isoform (Hepler et aL, 1990). The larger isoform contains additional sequences in its 3'-UT, and these are the presumed target(s) for the RNase. A gene for a mammalian cell endoribonuclease and candidate mRNase has recently been identified by virtue of its capacity to complement a mutant Escherichia coli RNase E gene (Wang and Cohen, 1994). RNase E has a variety of functions in E. coli, including rRNA processing and mRNA degradation (reviewed in Carpousis et al., 1994). The ard-1 RNase is a 13.3-kDa, highly basic, proline-rich protein. Neither RNase E nor the ard-1 enzyme has absolute specificity for a single RNA cleavage site in vitro, but both degrade the same region of 9S RNA, which is a known RNase E substrate in E. coli cells. Therefore, ard-1, like RNase E, is a potential mRNase, although there is no evidence as yet that it actually functions as such in mammalian cells. Most of the candidate mammalian mRNases described thus far are resistant to the RNase Aclass inhibitors. Thus, it will be important to learn whether the ard-1 RNase is similarly resistant. Several endoribonucleases with apparent specificity for certain mRNAs have been identified in amphibian cells and are described here because they are probably homologous to mammalian mRNases. One such enzyme, which is induced in X e n o p u s liver following estrogen treatment, degrades a deproteinized albumin mRNA substrate more rapidly than ferritin (Pastori et al., 1991a). This result is significant because it reflects a physiological response to estrogen, namely, the destabilization of albumin mRNA (Pastori et al., 1991b). Cleavage of the albumin substrate occurs primarily between the U and G of the sequence 5'AYUGA-3', which is found in the mRNA 5' region. This in vitro cleavage generates a 194-nt mRNA degradation product that is similar or identical to an RNA fragment observed in the liver of estrogen-treated animals (Pastori and Schoenberg, 1993; Dompenciel et al., 1995). The enzyme functions without magnesium but is more active with it and is associated
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with 80S ribosomes. Another endoribonuclease specifically cleaves maternal homeobox mRNAs in Drosophila cells and X e n o p u s oocytes (Brown and Harland, 1990). Cleavage occurs in the 3'-UT at a consensus sequence 5'-ANCUACCUA-3'. The 120-kDa enzyme has been partially purified and can degrade deproteinized RNA substrates (Brown et al., 1993). If radiolabeled substrate and a crude oocyte lysate are mixed and incubated. The substrate is degraded at a low rate. However, its decay is accelerated by adding excess unlabeled competitor RNA, implying that the lysate contains a factor that shields the substrate from the enzyme. In fact, many mRNAs appear to be protected somewhat from degradation by cytoplasmic nucleases, presumably by protein binding (Coutts et al., 1993). These findings highlight the notion that mRNAs are often protected from mRNases by the binding of shielding proteins. An RNase encoded by herpes simplex virus (HSV) types 1 and 2 seems to have remarkable specificity for mammalian cell mRNAs. HSV is a lytic virus with a wide host range and is a significant human pathogen (reviewed in Whitley, 1994). Almost all host cell mRNAs, even those with half-lives of over 20 hr in uninfected cells, are degraded within 3 hr after the onset of viral infection (reviewed in Roizman and Sears, 1990). Viral mRNAs also are destabilized, but neither ribosomal nor transfer RNAs are affected (Kwong and Frenkel, 1987; Oroskar and Read, 1989; Krikorian and Read, 1990). The destabilization of host mRNAs presumably serves to permit viral mRNAs ready access to the translation machinery without competition from host mRNAs. The destabilization of viral mRNAs permits transitions between immediate early to early and early to late viral gene expression to occur smoothly. Thus, the switch from one mRNA class to another requires only efficient transcription of the appropriate set of viral genes. mRNA destabilization is effected by the viral-encoded virion host shutoff (vhs) protein, a virion protein that is carried into the cell by the virus (Schek and Bachenheimer, 1985; Kwong and Frenkel, 1987, 1989; Oroskar and Read, 1987, 1989; Fenwick and Everett, 1990; Smibert and Smiley, 1990). The protein is phosphorylated and exists in several forms, with a major 58-kDa form in virions (Krikorian and Read, 1989; Read et al., 1993). Functional vhs protein might be a homodimer or multitimer, because certain UL41 mutants behave as dominant negatives in mixed infections with wild-type virus (Kwong and Frenkel, 1989). When extracts from cells infected with wild-type virus are incubated under appropriate conditions, both polysome-associated and protein-free mRNAs are rapidly degraded, whereas rRNA and tRNA are unaffected
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(Krikorian and Read, 1990; Sorenson et al., 1991). m R N A destabilization does not occur in reactions containing extracts from cells that had been infected with mutants encoding defective vhs proteins. To determine whether the vhs protein is itself a RNase or whether it functions indirectly, for example, by activating a latent host cell mRNase or inactivating a RNase inhibitor, crude viral extracts containing solubilized vhs protein were incubated with deproteinized substrates in reactions lacking any host components. The RNAs were degraded by extracts from the wild-type virus but not by those from viruses containing mutant vhs proteins (Zelus et al., 1995). Moreover, vhs protein prepared by translation in reticulocyte extracts also degraded m R N A substrates in vitro. The RNase activities in solubilized virions and in the translation extract were blocked by incubation with anti-vhs antibody. These data indicate that the vhs protein is an mRNase and raise the intriguing question of how it targets mRNA so specifically. The protein does bind strongly to poly(A) and poly(U) but not to poly(C) and appears to degrade deproteinized, polyadenylated RNA substrates more rapidly than nonpolyadenylated RNAs. Additional work is required to assess whether its specificity for mRNA is related to its RNA-binding specificity.
E. Perspective As is the case with rRNA-processing RNases, many details about mRNA stability are known, but some important issues are unresolved. How many RNases are devoted to degrading mRNAs in mammalian cells? How is their activity regulated? What is the nature of the substrates? That is, how do mRNA-binding proteins, polysome structure, and cytoplasmic localization of mRNAs affect mRNase function? Are mRNases localized? Some candidate mRNases are found primarily in postpolysomal fractions of mammalian cytosolic extracts (Hua et aL, 1993), and others are associated with ribosomes (Ross and Kobs, 1986; Brewer and Ross, 1988; Bernstein et al., 1992) or m R N A itself (Bandyopadhyay et al., 1990). All of the candidate mRNases thus far identified fail to be blocked by inhibitors of the RNase A-class of RNases (Sections III,B-III,D). Is this an important distinction between mRNases and RNases with other functions? It is impossible to predict at this stage if mRNA decay pathways and RNases are similar enough in mammals and S. cerevisiae to
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permit complementation analysis, as suggested for the rRNA-processing RNases (Section II,G). However, in yeast, mammals, and Caenorhabditis elegans, nonsense mutations in the 5' segments of mRNAs shorten their half-lives fivefold or more (reviewed in Cheng and Maquat, 1993; Pulak and Anderson, 1993; Peltz et aL, 1994), and some of the genes responsible for this destabilization effect are being identified and characterized (Leeds et al., 1991, 1992; Pulak and Anderson, 1993; Cui et aL, 1995; He and Jacobson, 1995). Thus, it might be feasible to exploit mutant strains of lower organisms to gain insights into the nature of mammalian mRNases.
IV. S U M M A R Y
Significant progress is being made in identifying the RNases responsible for processing and degrading rRNA and mRNA. Some of the enzymes have been purified and are being cloned, after which it should be possible to test their functions both genetically and biochemically. Perhaps the major questions to be answered concern the biochemical nature and structure of the substrates and how each substrate is recognized by a particular RNase. As has been discussed, the lack of absolute specificity of many RNases in vitro for a particular deproteinized substrate implies that, in cells, processing/degradation sites are influenced to a large extent by R N A - p r o t e i n structure. It seems reasonable to predict that some RNases are themselves regulated. However, controlling RNA processing/degradation by regulating RNases seems too inefficient and too prone to error to be a general mechanism. In the case of mRNA halflife regulation, for example, cells might need tens or hundreds of different mRNases to regulate the several thousand different mRNAs they contain. We think it unlikely that cells contain hundreds of different mRNases. Therefore, although RNases, like other enzymes, might be regulated in any number of ways, rRNA processing rates and m R N A decay rates are probably determined more by the structure and location of the substrates than by the RNases. If so, it is essential to determine the extent to which RNA primary sequence, secondary structure, and protein-binding affinity and specificity influence the processing/degradation rate. Are multiprotein complexes formed on the RNAs prior to their processing/degradation, as occurs during pre-mRNA splicing and 3' end formation (reviewed in Keller, 1995)? If not, how can we explain the fact that rRNA-processing
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enzymes lack absolute specificity for a particular sequence in vitro but cleave pre-rRNA (in cells) at only one or a few sites (Section II)? Do proteins bind to specific segments of RNA and function as beacons, directing RNases to those segments? Do other proteins bind to and shield RNAs from RNase attack? To what extent does compartmentalization of the RNAs, the RNases, or both influence RNase activity? Are there redundancies in terms of RNase function, as is the case with the 3' exoribonucleases that process E. coli tRNAs (Deutscher, 1993)? For example, if the 3' ~ 5' exoribonuclease that is a candidate histone mRNase (Section III,C) were completely inactivated in some way, would other enzymes degrade histone mRNA? Over the past 10 or so years, studies on RNase structure and function have revealed the central role of RNA processing and decay in regulating cell growth and differentiation. The half-lives of many mRNAs encoding cytokine, lymphokine, protooncogene, and transforming viral oncogene products are regulated posttranscriptionally ( Jones and Cole, 1987; Lindsten et al., 1989; Stoeckle and Hanafusa, 1989; Bohjanen et al., 1991; Baer et al., 1992; Chen et al., 1993; Iwai et al., 1993; Kanamori et al., 1994; Nair et al., 1994; Jeon and Lambert, 1995; reviewed in Ross, 1995). It will be important to identify the RNases involved in their regulation, to understand the mechanism of regulation, and, perhaps, to learn how to change rRNA and mRNA processing and degradation selectively.
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Bakker, O., Arnberg, A. C., Noteborn, M. H. M., Winter, A. J., and Ab, G. (1988). Turnover products of the apo very low density lipoprotein II messenger RNA from chicken liver. Nucleic Acids Res. 16, 10109-10118. Bandyopadhyay, R., Coutts, M., Krowczynska, A., and Brawerman, G. (1990). Nuclease activity associated with mammalian mRNA in its native state: Possible basis for selectivity in mRNA decay. Mol. Cell Biol. 10, 2060-2069. Bartsch, I., Schoneberg, C., and Grummt, I. (1988). Purification and characterization of "I'TFI, a factor that mediates termination of mouse ribosomal DNA transcription. Mol. Cell Biol. 8, 3891-3897. Beelman, C. A., and Parker, R. (1995). Degradation of mRNA in eukaryotes. Cell (Cambridge, Mass.) 81, 179-183. Belgrader, P., Cheng, J., and Maquat, L. E. (1993). Evidence to implicate translation by ribosomes in the mechanism by which nonsense codons reduce the nuclear level of human triosephosphate isomerase mRNA. Proc. Natl. Acad. Sci. U.S.A. 90, 482-486. Bernstein, P., and Ross, J. (1989). Poly(A), poly(A)-binding protein and the regulation of mRNA stability. Trends Biochem. Sci. 14, 373-377. Bernstein, P., Peltz, S. W., and Ross, J. (1989). The poly(A)-poly(A)-binding protein complex is a major determinant of mRNA stability in vitro. Mol. Cell. Biol. 9, 659-670. Bernstein, P. L., Herrick, D. J., Prokipcak, R. D., and Ross, J. (1992). Control of c-myc mRNA half-life in vitro by a protein capable of binding to a coding region stability determinant. Genes Dev. 6, 642-654. Beltrame, M., and Tollervey, D. (1992). Identification and functional analysis of two U3 binding sites on yeast pre-ribosomal RNA. E M B O J. 11, 1531-1542. Binder, R., Hwang, S.-P. L., Ratnasabapathy, R., and Williams, D. L. (1989). Degradation of apoliprotein II mRNA via endonucleolytic cleavage at 5'-AAU-3'/5'-UAA-3' elements in single stranded loop domains at the 3' noncoding region. J. Biol. Chem. 264, 16910-16918. Binder, R., Horowitz, J. A., Basilion, J. P., Koeller, D. M., Klausner, R. D., and Harford, J. B. (1994). Evidence that the pathway of transferrin receptor mRNA degradation involves an endonucleolytic cleavage within the 3' UTR and does not involve poly(A) tail shortening. E M B O J. 13, 1969-1980. Bohjanen, P. R., Petryniak, B., June, C. H., Thompson, C. B., and Lindsten, T. (1991). An inducible cytoplasmic factor (AU-B) binds selectively to AUUUA multimers in the 3' untranslated region of lymphokine mRNA. Mol. Cell. Biol. 11, 3288-3295. Bohjanen, P. R., Petryniak, B., June, C. H., Thompson, C. B., and Lindsten, T. (1992). AU RNA-binding factors differ in their binding specificities and affinities. J. Biol. Chem. 267, 6302-6309. Bowman, L. H., Goldman, W. E., Goldberg, G. I., Hebert, M. B., and Schlessinger, D. (1983). Location of the initial cleavage sites in mouse pre-rRNA. Mol. Cell Biol. 3, 1501-1510. Brewer, G., and Ross, J. (1988). Poly(A) shortening and degradation of the 3' AU-rich sequences of human c-myc mRNA in a cell-free system. Mol. Cell Biol. 8, 1697-1708. Brewer, G., and Ross, J. (1989). Regulation of c-myc mRNA stability in vitro by a labile destabilizer with an essential nucleic acid component. Mol. Cell Biol. 9, 1996-2006. Brown, B. D., and Harland, R. M. (1990). Endonucleolytic cleavage of a maternal homeo box mRNA in Xenopus oocytes. Genes Dev. 4, 1925-1935. Brown, B. D., Zipkin, I. D., and Harland, R. M. (1993). Sequence-specific endonucleolytic cleavage and protection of mRNA in Xenopus and Drosophila. Genes Dev. 7, 16201631.
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Shuai, K., and Warner, J. R. (1991). A temperature sensitive mutant of Saccharomyces cerevisiae defective in pre-rRNA processing. Nucleic Acids Res. 19, 5059-5064. Shumgard, C. M., and Eichler, D. C. (1988). Ribosomal RNA processing: Limited cleavages of mouse preribosomal RNA by a nucleolar endoribonuclease include the early +650 processing site. J. Biol. Chem. 263, 19346-19352. Shumgard, C. M., Torres, C., and Eichler, D. C. (1990). In vitro processing at the 3'terminal region of pre-18S rRNA by a nucleolar endoribonuclease. Mol. Cell Biol. 10, 3868-3872. Shyu, A.-B., Greenberg, M. E., and Belasco, J. G. (1989). The c-los transcript is targeted for rapid decay by two distinct mRNA degradation pathways. Genes Dev. 3, 60-72. Shyu, A.-B., Belasco, J. G., and Greenberg, M. E. (1991). Two distinct destabilizing elements in the c-los message trigger deadenylation as a first step in rapid mRNA decay. Genes Dev. 5, 221-231. Smibert, C. A., and Smiley, J. R. (1990). Differential regulation of endogenous and transduced/3-globin genes during infection of erythroid cells with a herpes simplex virus type 1 recombinant. J. Virol. 64, 3882-3894. Smoske0y, S., Rao, M. N., and Slobin, L. (1995). Purification and characterization of a decapping 5 ' ~ 3' exoribonuclease from rabbit reticulocytes. Eur. J. Biochem. 237, 171-179. Sorenson, C. M., Hart, P. A., and Ross, J. (1991). Analysis of herpes simplex virus-induced mRNA destabilizing activity using an in vitro mRNA decay system. Nucleic Acids Res. 19, 4459-4465. Stevens, A., Hsu, C. L., Isham, K. R., and Larimer, F. W. (1991). Fragments of the internal transcribed spacer 1 of pre-rRNA accumulate in Saccharomyces cerevisiae lacking 5'---~ 3' exoribonuclease 1. J. Bacteriol. 173, 7024-7028. Stevens, A. (1978). An exoribonuclease from Saccharomyces cerevisiae: Effect of modifications of 5' end groups on the hydrolysis of substrates to 5' mononucleotides. Biochem. Biophys. Res. Commun. 81, 656-661. Stoeckle, M. Y. (1991). Post-transcriptional regulation of groc~,/3, y, and IL-8 mRNAs by IL-I/3. Nucleic Acids Res. 19, 917-920. Stoeckle, M. Y. (1992). Removal of a 3'-non-coding sequence is an initial step in degradation of groo~ mRNA and is regulated by interleukin-l. Nucleic Acids Res. 20, 1123-1127. Stoeckle, M. Y., and Guan, L. (1993). High-resolution analysis of groc~ poly(A) shortening: Regulation by interleukin-l/3. Nucleic Acids Res. 21, 1613-1617. Stoeckle, M. Y., and Hanafusa, H. (1989). Processing of 9E3 mRNA and regulation of its stability in normal and Rous sarcoma virus-transformed cells. Mol. Cell Biol. 9, 4738-4745. Swartwout, S. G., and Kinniburgh, A. J. (1989). c-myc RNA degradation in growing and differentiating cells: Possible alternate pathways. Mol. Cell. Biol. 9, 288-295. Swartwout, S. G., Preisler, H., Guan, W., and Kinniburgh, A. J. (1987). A relatively stable population of c-myc RNA that lacks long poly(A). Mol. Cell. Biol. 7, 2052-2058. Tharun, S., and Sirdeshmukh, R. (1995). Specific endonucleolytic leavages of mouse albumin mRNA and their modulation during liver development. Nucleic Acids Res. 23, 641-646. Treisman, R. (1985). Transient accumulation of c-los RNA following serum stimulation requires a conserved 5' element and c-los 3' sequences. Cell (Cambridge, Mass.) 42, 889-902.
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Wang, M., and Cohen, S. N. (1994). ard-l: A human gene that reverses the effects of temperature-sensitive and deletion mutations in the Escherichia coli rne gene and encodes an activity producing RNase E-like cleavages. Proc. Natl. Acad. Sci. U.S.A. 91, 10,591-10,595. Whitley, R. J. (1994). Herpes simplex virus infections of women and their offspring: Implications for a developed society. Proc. Natl. Acad. Sci. U.S.A. 91, 2441-2447. Wisdom, R., and Lee, W. (1990). Translation of c-myc mRNA is required for its posttranscriptional regulation during myogenesis. J. Biol. Chem. 265, 19015-19021. Wisdom, R., and Lee, W. (1991). The protein-coding region of c-myc mRNA contains a sequence that specifies rapid mRNA turnover and induction by protein synthesis inhibitors. Genes Dev. 5, 232-243. Wreschner, D. H., and Rechavi, G. (1988). Differential mRNA stability to reticulocyte ribonucleases correlates with 3' non-coding (U)nA sequences. Eur. J. Biochem. 172, 333-340. Yip, M. T., and Holland, M. J. (1989). In vitro RNA processing generates mature 3' termini of yeast 35 and 25S ribosomal RNAs. J. Biol. Chem. 264, 4045-4051. Yu, Y.-T., and Nilsen, T. W. (1992). Sequence requirements for maturation of the 5' terminus of human 18S rRNA in vitro. J. Biol. Chem. 267, 9264-9268. Yuan, Y., Singh, R., and Reddy, R. (1989). Rat nucleolar 7-2 RNA is homologous to mouse mitochondrial RNase mitochondrial RNA-processing RNA. J. Biol. Chem. 264, 14835-14839. Zelus, B. D., Stewart, R. S., and Ross, J. (1996). The virion host shutoff protein of Herpes simplex virus type 1: messenger ribonucleolytic activity in vitro. J. Virol. 70, 2411-2419. Zhang, W., Wagner, B. J., Ehrenman, K., Schaefer, A. W., DeMaria, C. T., Crater, D., DeHaven, K., Long, L., and Brewer, G. (1993). Purification, characterization, and cDNA cloning of an AU-rich element RNA-binding protein, AUF1. Mol. Cell. Biol. 13, 7652-7665.
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18 Messenger R N A Ribonucleases and m R N A Turnover in
Saccharomyces cerevisiae CHRISTINE E. B R O W N A N D A L A N B. S A C H S Department of Molecular and Cell Biology University of California at Berkeley Berkeley, California 94720
I. Introduction II. Pathways of mRNA Decay A. Deadenylation-Dependent Decapping B. Nonsense-Mediated Decay C. Endonuclease-Initiated Turnover Pathway III. mRNA Ribonucleases A. Decapping Enzymes B. Deadenylases C. 5'--, 3' Exoribonucleases D. 3' ---, 5' Exoribonucleases E. Endoribonucleases IV. Discussion and Future Directions References
I. I N T R O D U C T I O N
A m o n g t h e m a n y d i f f e r e n t f u n c t i o n s for r i b o n u c l e a s e s , o n e f u n d a m e n tal r o l e is t o d e g r a d e m e s s e n g e r R N A ( m R N A ) . T h e r e g u l a t e d t u r n o v e r
589 RIBONUCLEASES: STRUCTURES AND FUNCTIONS
Copyright 9 1997 by Academic Press, Inc. All rights of reproduction in any form reserved.
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of mRNA is critical in the control of gene expression. For instance, the stabilization of certain unstable mRNAs encoding protooncogenes can contribute to oncogenesis (reviewed by Schiavi et al., 1992). In addition, mRNA stability can be regulated in accordance with changes in the cell growth rate (for example, Kindy and Sonnenshein, 1986), progression through the cell cycle (for example, Morris et al., 1991), and changes in the cellular environment (for example, Mullner and Kuhn, 1988; Owen and Kuhn, 1987; Yen et al., 1988). In many cases, mRNA abundance seems to correlate more closely with mRNA half-lives than with rates of transcription (reviewed by Peltz et al., 1991). The ability to perform both advanced genetics and biochemistry with Saccharomyces cerevisiae makes it well suited for dissecting the fundamental mechanisms of mRNA degradation in eukaryotes. In yeast, some mRNAs are degraded with half-lives greater than 40 min, whereas others are degraded 20 times faster. Ultimately, differential mRNA decay is determined by the regulation of cellular mRNA ribonucleases (mRNases). Eukaryotic mRNase activities have been previously reviewed by A. Stevens (1993), and are also discussed in Chapter 17 in this volume. The focus in this review will be on mRNases in S. cerevisiae, beginning with an overview of pathways of mRNA degradation in yeast, followed by a discussion in greater detail of the biochemistry of the mRNases involved in these pathways.
II. P A T H W A Y S OF m R N A D E C A Y
Two pathways of mRNA decay in S. cerevisiae have been well characterized. The first, deadenylation-dependent decapping, is thought to constitute a general way by which many polyadenylated mRNAs are degraded. The second, nonsense-mediated decay, is dedicated to rapidly degrading transcripts containing premature stop codons. In addition, mRNAs can be degraded by other pathways, an example of which is endonucleolytic cleavage of the message. Several technical strategies have been important in the elucidation of mRNA turnover pathways in yeast. First, the creation of a synchronous pool of newly synthesized messages by the transient induction of the galactose-inducible promoter (transcriptional pulse-chase experiments) has enabled the timing of mRNA decay events to be followed (Decker and Parker, 1993). Second, a poly(G)18 cassette, which forms a strong
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RNA secondary structure (Sen and Gilbert, 1992) that can stall exoribonucleases (Stevens and Poole, 1995), has enabled mRNA degradative intermediates to be detected (Decker and Parker, 1993; Muhlrad et al., 1994, 1995; Vreken and Raue, 1992). Third, the use of strains lacking the 5' ~ 3' exoribonuclease, Xrnlp, that destroys uncapped mRNA (for further discussion, see Section III,C,1) has allowed for both an analysis of the contributions of this enzyme to mRNA degradation and for a simple way to accumulate uncapped mRNA degradative intermediates (Hsu and Stevens, 1993; Muhlrad et aL, 1994, 1995). Finally, the use of antibodies capable of specifically immunoprecipitating capped mRNA (Munns et al., 1982) has enabled the separation of capped from uncapped transcripts within a population of mRNAs (Muhlrad et al., 1994, 1995).
A. Deadenylation-Dependent Decapping
Detailed studies investigating mRNA turnover in yeast have revealed that both stable and unstable polyadenylated mRNAs can be degraded by a common pathway, termed deadenylation-dependent decapping (reviewed by Beelman and Parker, 1995; Decker and Parker, 1994). For transcripts degraded by this mechanism, poly(A) tail removal triggers decapping, and then the transcript is rapidly destroyed (Fig. 1A). Evidence that mRNA turnover can be initiated by shortening of the poly(A) tail was demonstrated by transcriptional pulse-chase experiments, which showed that mRNA degradation occurred only after the transcript had been deadenylated to tail lengths less than 15 adenylate residues (Decker and Parker, 1993). By using either Xrnlp-deficient yeast strains or an mRNA containing a poly(G) tract to trap decay intermediates, along with antibodies to capped mRNA, it was demonstrated that decapping temporally follows deadenylation (Muhlrad et al., 1994, 1995). How a structural change at the 3' end of the mRNA induces decapping remains to be resolved (for further discussion, see Section III,B,1). Upon removal of these terminal structures, the uncapped, poly(A)- transcript is rapidly destroyed. A major mode of hydrolysis of the body of the message is 5' ~ 3' exonucleolytic digestion by the Xrnl ribonuclease (Hsu and Stevens, 1993; Muhlrad et al., 1994, 1995). A minor mode of hydrolysis is 3' ~ 5' exonucleolytic degradation, which
A
m7Gppp---~AUG
m7Gppp----~AUG
UAA~'(A)60.90
1
Deadenylation
NonsenseMutation
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m7Gppp----~AUG UAA
Translation u~(A)5.15
UPF-dependent recognition of nonsense mutation
I Decapping l~AUG
UAA]'--"(A)60.90
u~(A)5.15
m7GpPP~(A)15-90 Decapping
5'-3' Exonucleolytic Degradation ml~AUG
UA~.. (A)15.90
3'-5 Exonucleolytic Degradation UAA~--,-(A)5.15
5'-Y Exonucleolytic Degradation
m7Gppp-----qA'uG UAA~--'--(A)15.90
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can be detected for some mRNAs (Muhlrad et al., 1995) but not for others (Muhlrad et al., 1994). Transcripts with differing stabilities, such as the stable P G K 1 mRNA (tl/2 = 45 min) and the unstable M F A 2 mRNA (tl/2 = 3.5 min), are known to be degraded by the deadenylation-dependent decapping pathway (Decker and Parker, 1994; Muhlrad et al., 1994, 1995). Differences in mRNA turnover rates appear to be due to differences in both the rates of poly(A) tail shortening and the decapping of the transcript (Muhlrad et al., 1994, 1995). As a result, an understanding of how these two steps are regulated is needed to elucidate the mechanisms underlying the regulation of mRNA degradation in yeast. Is the deadenylation-dependent decapping pathway conserved for other eukaryotes? Deadenylation is the first step in decay for the group of rapidly degraded messages containing the 3' untranslated region (3'UTR)AU-rich (ARE) destabilizing element (Brewer and Ross, 1988; Chen and Shyu, 1995b; Chen et al., 1995a; Shyu et al., 1991; Swartwout and Kinniburgh, 1989; Wilson and Treisman, 1988). However, it is unclear if deadenylation is a prerequisite for the turnover of stable mRNAs or if deadenylation triggers decapping. To a large degree this uncertainty results from the fact that the tools used in yeast to trap degradation intermediates [poly(G) tracts or ribonuclease-deficient strains] have not yet been successfully employed in other eukaryotic systems.
Fig. 1. mRNA turnover pathways in yeast. (A) The deadenylation-dependent decapping pathway of mRNA decay is initiated by poly(A) tail shortening. Following tail removal to an oligoadenylate length of approximately 5-15 residues, the mRNA is decapped and degraded by 5' --. 3' exoribonucleolytic digestion. In addition, a 3' --~ 5' exonucleolytic pathway is also detected for the degradation of certain mRNAs. Other nuclease activities not depicted, such as endonucleases, could be functioning in this general pathway of mRNA turnover, although little is known about their involvement. (B) The nonsensemediated decay pathway is initiated by the up frameshift (UPF)-dependent recognition of early nonsense codons. Upon detection of the nonsense mutation, the mRNA is decapped and rapidly degraded via a 5' --~ 3' exonucleolytic mechanism. In contrast to the deadenylation-dependent pathway, decapping of an mRNA degraded by the nonsense-mediated pathway does not require prior deadenylation. Other mRNases not depicted may also be functioning in this pathway.
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B. Nonsense-Mediated Decay Messenger RNA degradation is also stimulated by the presence of early nonsense codons (Fig. 1B) (reviewed by Maquat, 1995; Peltz et al., 1994). Nonsense codons can arise from a variety of faulty cellular events such as genomic mutations, inaccurate splicing, and RNA editing errors. This decay pathway probably exists in eukaryotic cells to decrease the expression level of potentially toxic truncated polypeptides derived from the mutated mRNAs. In yeast, recognition of the stop codon as being an early nonsense mutation depends on the activity of the up frameshift (UPF) pathway. Mutant Upf proteins were originally identified as allosuppressors and later were shown to lead to the stabilization of mRNAs containing an early nonsense codon (Leeds et al., 1991, 1992). Although this pathway is conserved among eukaryotes, little is known about the functions of the individual proteins comprising it. To begin to understand the mechanism of mRNA turnover for nonsense-mediated decay, the degradation of the stable yeast P G K 1 mRNA harboring a premature stop codon was investigated (Muhlrad and Parker, 1994). The introduction of an early stop codon into the P G K 1 transcript can destabilize this mRNA greater than 10-fold (Muhlrad and Parker, 1994; Peltz et aL, 1993). Rapid degradation of this nonsensecontaining mRNA occurs by the stimulation of decapping, followed by 5' ~ 3' exonucleolytic digestion (Muhlrad and Parker, 1994). Unlike the deadenylation-dependent pathway, mRNA decay is initiated by decapping that is independent of poly(A) tail removal. However, similar to the deadenylation-dependent pathway, nonsense-mediated decay also appears to utilize Xrnlp. Mutated P G K 1 mRNA is stabilized sixfold in yeast strains lacking Xrnlp, suggesting that it is normally degraded by this enzyme (Muhlrad and Parker, 1994). These observations support a model in which early nonsense mutations stimulate, through the UPF pathway, the decapping and rapid destruction of polyadenylated transcripts.
C. Endonuclease-lnitiated Turnover Pathway Messenger RNA can also be degraded via an endonuclease-initiated turnover pathway. One example of this pathway in S. cerevisiae is
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the autogenous control of R P L 2 mRNA degradation (Presutti et al., 1991, 1995). R P L 2 encodes the ribosomal protein L2 (rpL2), and excess amounts of rpL2 destabilize its own mRNA by triggering endonucleolytic cleavage within its 5'-UTR (Presutti et al., 1995). A 200-nucleotide element in the R P L 2 5'-UTR is sufficient for rpL2dependent degradation. Interestingly, this sequence promotes endonucleolytic scission even when inserted into an intron of a heterologous message, suggesting that cleavage can occur in the nucleus (Presutti et al., 1995). In other eukaryotes mRNA degradation can also be initiated by endonucleolytic cleavage. The 5' and 3' degradative products produced from an internal cleavage event have been detected for several messages, including the mammalian transferrin receptor TfR mRNA (Binder et al., 1994), the mammalian insulin-like growth factor I G F I I mRNA (Nielsen and Christiansen, 1992), the avian apolipoprotein aport mRNA (Binder et al., 1989), the avian cytokine 9E3 mRNA (Stoeckle and Hanafusa, 1989), and the X e n o p u s homeobox X l h b o x 2 B mRNA (Brown and Harland, 1990). In each of these examples, transcript turnover seems to be initiated by endonucleolytic cleavage at a sequence-specific site. Although the mechanism of mRNA turnover initiated by endonucleolytic scission is not well understood, it appears that the timing of cleavage is independent of poly(A) tail removal (Binder et al., 1994; Brown and Harland, 1990; Stoeckle and Hanafusa, 1989). The turnover of mRNA by an endonuclease-initiated pathway is often controlled by trans-acting factors. As discussed previously, cleavage of the R P L 2 mRNA is regulated by the binding of excess rpL2 to a sequence within the 5'-UTR of the transcript (Presutti et al., 1991, 1995). In other eukaryotes, perhaps the best characterized example of such regulation is the iron-dependent turnover of the TfR mRNA (Casey et al., 1989; Klausner et al., 1993). Furthermore, the decay of many endonucleolytically cleaved mRNAs is highly regulated, particularly by changes in the cellular environment. For instance, the R P L 2 mRNA is under an autogenous feedback control, and TfR mRNA is regulated by cellular iron concentrations, and the aport mRNA is controlled by estrogen levels. Based on these observations, it has been proposed that a general pathway of decay would be initiated by poly(A) tail shortening, and that other pathways such as endonuclease cleavage would be limited to the subsets of mRNAs requiring more complex regulation (Beelman and Parker, 1995; Decker and Parker, 1994).
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III. m R N A R I B O N U C L E A S E S
Five classes of identified mRNA ribonuclease activities will be presented. Messenger RNases that degrade the 5' cap structure and the 3' poly(A) tail, the decapping enzymes and deadenylases, respectively, will be discussed first, because removal of these terminal structures seem to be rate limiting for several pathways of mRNA degradation. Discussion of mRNases that degrade the body of the message exonucleolytically, i.e., the 5'----> 3' exoribonucleases and 3'---> 5' exoribonucleases, will follow. Finally, endonucleases, mRNases that cleave internal to the message, will be presented. Although discussed separately, these mRNase activities work in concert to determine message-specific turnover rates.
A. Decapping Enzymes
It is well established that eukaryotic cellular mRNA possesses a 5' cap structure consisting of a 7-methylguanylate covalently attached to the transcript via a unique 5' to 5' phosphodiester linkage (reviewed by Banerjee, 1980; Shatkin, 1976) (see Figure 2). The 5' cap structure is added cotranscriptionally and is implicated in several processes, including premessenger RNA splicing (Inoue et al., 1989; Izaurralde et al., 1994; Konarska et al., 1984; Krainer et al., 1984; Ohno et al., 1987; Patzelt et al., 1987), nucleocytoplasmic mRNA transport (Hamm and Mattaj, 1990; Izaurralde et al., 1992; Jarmolowski et al., 1994), translation initiation (Gallie, 1991; Rhoads, 1988; Sonenberg, 1988), and mRNA stability (see below). It is generally accepted that the 5' cap stabilizes an mRNA by protecting the message from nonspecific exonucleolytic degradation. In support of this hypothesis, many purified 5' ~ 3' exonucleases are unable to degrade capped RNA (Kenna et al., 1993; Murthy et al., 1991). In addition, a capped mRNA is usually more stable than a message lacking a cap in crude cell extracts and in X e n o p u s oocytes (Furuichi et al., 1977, Green et al., 1983; Shimotohno et al., 1977). Moreover, the importance of the decapping reaction is highlighted by both the deadenylationdependent decapping and the nonsense-mediated decay pathways, wherein the removal of the cap structure can be rate limiting for mRNA turnover (Muhlrad et al., 1994, 1995; Muhlrad and Parker, 1994). Thus far, two decapping activities have been identified in S. cerevisiae.
18
597
mRNA/mRNases in Saccharomyces cerevisiae Gag coat protein of the L-A Virus
Decapping enzyme of S. cerevisiae
HO
HO
1 O
~
-o'
'
O__
- o'
O
5'
Basel
o
H2 N
O
O(Cl~3) O I O= P--O"- CH2
3'
l
O I O=P--O I O
Base 2
/
O(CH3) 9" " " "
Fig. 2. Structure of the m R N A cap structure and sites of decapping. The m R N A cap structure consists of a 7'-methylguanylate residue covalently attached to the 5' end of the m R N A via a 5' ---, 5' phosphodiester linkage. The sites of decapping for the purified yeast decapping activity and for the Gag coat protein are indicated.
1. Decapping Activity Purified from High Salt Wash of Yeast Ribosomes An mRNA decapping activity, with an apparent molecular mass of 79 kDa, has been purified over 10,000-fold from a ribosomal high salt wash fraction of S. cerevisiae (Stevens, 1988). Characterization of this purified enzyme demonstrates that it decaps mRNA by hydrolyzing the /3-pyrophosphate bond of the cap structure, yielding mVGDP and 5'-pRNA as products (Stevens, 1980a) (see Fig. 2). Unlike nonspecific pyrophosphatases, which cleave a wide range of phosphodiester bonds, including the 5' cap structure (Bartkiewicz et al., 1984; Kole et al., 1976; Shinshi et al., 1976a,b), the purified yeast decapping activity is highly specific. For example, the yeast decapping enzyme does not cleave the
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pyrophosphate bond of ppp-RNA, UDP-glucose, or mTGpppA(G) (Stevens, 1980a). This decapping enzyme strongly prefers long RNA chains. For instance, an RNA molecule of 540 nucleotides is decapped 10-fold more efficiently than an RNA molecule of 50 nucleotides (Stevens, 1988). In contrast, an mTGpppN-pyrophosphatase purified from both HeLa cells and human placenta can decap very short oligonucleotides but not oligonucleotides of 10 or more residues or intact mRNA (Nuss et al., 1975, 1982). Substrate recognition by the yeast decapping enzyme does not include the 7-methyl moiety, because GpppG-RNA and mTGpppG RNA substrates are decapped with relatively equal efficiencies (Stevens, 1988). The purified decapping enzyme does not appear to contain any general 5' ~ 3' exonuclease activity (Stevens, 1988). Biochemical analysis of the purified decapping enzyme (Stevens, 1980a, 1988) demonstrates that the optimum pH for enzymatic activity is between 7.5 and 8.5, and the optimum temperature is 30~ Catalysis requires a divalent cation, as indicated by EDTA-dependent inhibition of the decapping activity that is reversible upon the addition of 1 mM Mg 2§ or Mn 2§ The enzyme is sensitive to high ionic strength, being completely inhibited at 200 mM NHaCI. Its Km for yeast RNA is 12.5/xM. How do the characteristics of this purified decapping activity shed light onto the pathways of mRNA turnover? That full-length mRNA species are decapped in the deadenylation-dependent and nonsensemediated decay pathways is consistent with the finding that the purified decapping activity prefers long-chain RNA molecules. This observation may suggest that cap removal from degradative intermediates (potentially occurring by endonucleolytic cleavage) may require a second enzyme, such as the mTGpppG pyrophosphatase activity purified from mammalian cells. Similarly, if pathways of mRNA turnover are to be conserved between eukaryotes, a decapping enzyme that recognizes fulllength mRNA must exist in mammalian cells. 2. Gag Coat Protein of dsRNA Virus L-A in Decapping mRNA UV cross-linking studies aimed at identifying cap-binding proteins in S. cerevisiae detected a protein that binds covalently to the cap structure in the absence of UV irradiation (Goyer et al., 1989). This protein has
been identified as the Gag coat protein of the yeast dsRNA virus L-A (Blanc et al., 1992). Further studies confirm that the Gag coat protein binds to the mRNA cap structure in vivo, and that upon binding it decaps the host mRNA (Masison et al., 1995). The L-A virus is one of five
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commonly harbored dsRNA viruses in laboratory strains of S. cerevisiae (reviewed by Wickner, 1992). The plus-strand RNA of the L-A virus lacks both a cap and a poly(A) tail (Bruenn and Keitz, 1976; Bruenn and Brennan, 1980; Hannig et al., 1984) and contains two overlapping open reading frames (ORFs). ORF1 encodes the Gag coat protein and ORF2, upon a - 1 ribosomal frameshift, encodes the Gag-Pol fusion protein (Dinman et al., 1991; Fujimura and Wickner, 1988; Icho and Wickner, 1989). Investigation into the mechanism by which the Gag protein decaps mRNA reveals that the a-pyrophosphate bond of m7GpppG-RNA is hydrolyzed to produce mTGMP and ppG-RNA (Blanc et al., 1994) (see Fig. 2). Covalent attachment occurs through the His-154 residue of the Gag coat protein and the a-phosphate of the cap structure (Blanc et al., 1994) in a reaction that requires Mg 2§ (Goyer et al., 1989). Mutational analysis of the His-154 residue confirms that it is required for both the covalent attachment and decapping activities, but not for binding to the cap structure (Blanc et al., 1994). The minimal cap structure required for covalent attachment to the Gag coat protein is mTGpppGp; neither m7Gp nor m7GpppG is a substrate. It is believed that the 3'-phosphate of the second G is important for correct positioning of the cap structure in the cap-binding pocket of the Gag protein. For binding, the Gag coat protein requires a methylguanosine and does not bind to unmethylated GpppG-RNA (Blanc et al., 1992). What is the role for the Gag decapping activity? It has been suggested that the decapping activity is required for efficient expression of viral gene products (Masison et al., 1995). Given that the cap structure stabilizes an mRNA, it would be expected that an L-A plus-strand RNA that lacks a 5' cap structure would be highly unstable. However, by decapping other cellular mRNAs through the activity of its Gag protein, the L-A virus is proposed to decoy the mRNase degradative machinery by increasing its saturation with substrate, thereby increasing the relative stability of its own uncapped transcripts and ensuring their expression (Masison et al., 1995). 3. Outstanding Questions The two aforementioned yeast decapping activities function to hydrolyze the pyrophosphate bonds between the cap structure and the first nucleotide of the mRNA, thus leaving the transcript intact. Does this correspond to the site of mRNA decapping in vivo? The putative in vivo
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site of decapping can be determined by reverse transcription experiments when the 5' --~ 3' exoribonuclease pathway is partially blocked by a deletion of the major 5' ~ 3' exoribonuclease, Xrnlp. In such experiments, as might be expected, the decapped M F A 2 m R N A is found to be full length. In contrast, both the decapped P G K 1 (Muhlrad et al., 1995) and R P 5 1 A (Hsu and Stevens, 1993) transcripts are found to be shortened by two nucleotides. This could be due to weak 5' --~ 3' exoribonuclease activity in vivo. Alternatively, the P G K 1 and R P 5 1 A messages may be decapped by a different class of enzymes whose initial cleavage site is within the 5'-UTR of the mRNA. For example, in cultured mammalian cells a 5' --~ 3' exoribonuclease has been identified that can degrade capped transcripts by initiating exonucleolytic degradation a few nucleotides into the body of the message (Coutts and Brawerman, 1993).
B. Deadenylases At the 3' end of most eukaryotic mRNAs is a stretch of adenylate residues known as the poly(A) tail. Poly(A) addition occurs postranscriptionally in the nucleus as part of a two-step process including endonucleolytic cleavage of the precursor RNA to form the mature 3' end and polymerization of ATP to yield the poly(A) tail (reviewed by Sachs and Wahle, 1993; Wahle and Keller, 1992). Poly(A) tail length is organism specific, and can range from 70 to 90 nucleotides in yeast to 200 to 250 nucleotides in mammalian cells (reviewed by Baker, 1993). Similar to the 5' cap structure, the 3' poly(A) tail is believed to function in several processes, including translation initiation (reviewed by Jackson and Standart, 1990; Munroe and Jacobson, 1990b; Sachs, 1990) and mRNA stability (reviewed by Beelman and Parker, 1995; Decker and Parker, 1994; Sachs, 1993). In the cytoplasm, the poly(A) tail is bound by the highly conserved poly(A) binding protein (Pablp). The yeast poly(A) binding protein is encoded by the essential P A B 1 gene (Sachs et al., 1986, 1987). P A B 1 has been shown to be required for both translation initiation (Munroe and Jacobson, 1990a; Sachs and Davis, 1989; Tarun and Sachs, 1995) and aspects of mRNA turnover (Caponigro and Parker, 1995; Sachs and Davis, 1989). Pablp binds to the poly(A) tail with a periodicity of approximately 25 adenylate residues and seems to require a minimum
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of 12 residues for binding (Sachs et al., 1987). Poly(A) function is believed to be mediated by the Pablp-poly(A) complex. The mRNA poly(A) tail is progressively shortened at a messagespecific rate in the cytoplasm. For instance, in S. cerevisiae the stable P G K 1 mRNA is deadenylated 4 ___ 2 residues per minute whereas the unstable M F A 2 mRNA is deadenylated 13 ___ 2 residues per minute (Decker and Parker, 1993). Consistent with a relationship between poly(A) tail removal and mRNA turnover, rates of deadenylation often correlate with mRNA stability (Decker and Parker, 1993; Shyu et aL, 1991). Rapid deadenylation can be promoted by cis-acting elements within an unstable message, and when such instability elements are transferred to a stable transcript, the chimeric message is more rapidly deadenylated and degraded (Decker and Parker, 1993; Shyu et al., 1991). Similarly, mutations in an unstable message that slow deadenylation rates also render the message more stable (C.-Y. A. Chen et al., 1994; Muhlrad and Parker, 1992). These types of experiments strongly suggest that deadenylation rates can be critical in determining mRNA half-lives. As has been discussed, poly(A) tail removal can be a prerequisite for mRNA decapping. Poly(A) tails appear to inhibit mRNA decapping through the activity of Pablp. The role of Pablp in mRNA decay was examined by utilizing yeast strains harboring a conditional allele of the essential P A B I gene or harboring a secondary bypass mutation suppressing a P A B I deletion (Caponigro and Parker, 1995). One of the phenotypes associated with loss of Pablp is the uncoupling of the deadenylation and decapping reactions. In these studies, mRNA decay intermediates harbored poly(A) tail lengths of up to 75 nucleotides, demonstrating that without Pablp mRNA decay initiates without prior deadenylation (Caponigro and Parker, 1995). This is in contrast to wildtype yeast, for which deadenylation is a prerequisite for decapping, and thus intermediates of mRNA decay contained very short or no poly(A) tails (Caponigro and Parker, 1995; Muhlrad et al., 1995; Muhlrad and Parker, 1994). The length of the poly(A) tail for yeast mRNA predicted to stimulate the decapping reaction (5-15 adenylate residues) is similar to the minimal poly(A) length required for Pablp binding (12 adenylate residues) (Sachs et al., 1987). Thus, it is modeled that when the mRNA poly(A) tail is shortened to an oligo(A) length, Pablp is lost from the message and decapping ensues. These observations suggest a fundamental role for mRNA deadenylases in differential mRNA stability. Thus far, only one deadenylase has been identified and characterized in S. cerevisiae.
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1. Pablp-Dependent Poly(A) Nuclease of Saccharomyces cerevisiae The yeast deadenylase, referred to as poly(A) nuclease (PAN), has been purified over 100,000 fold from S100 extracts of S. cerevisiae (Boeck et al., 1996; Sachs and Deardorff, 1992). This mRNase is unique in that it efficiently degrades only RNA bound by Pablp, and thus requires an mRNP substrate (Sachs and Deardorff, 1992). In the presence of Pablp, purified PAN exonucleolytically shortens homopolymers of poly(A), releasing 5'-AMP mononucleotides as products (Lowell et al., 1992). The existence of such a Pablp-dependent poly(A) nuclease was predicted based on several observations demonstrating that Pablp is required for the efficient shortening of poly(A) tails in vivo (Sachs and Davis, 1989). Biochemical characterization of the purified PAN activity (Lowell et al., 1992) reveals a requirement for Mg 2+ and a sensitivity to ionic conditions greater than 50 raM. The enzyme is active between a pH range of 7.0 to 8.5 and a temperature range of 30 ~ to 37~ although PAN can be heat inactivated at 55~ for 10 min. PAN enzymatic activity appears to require a 3'-hydroxyl group, because it will not degrade poly(A) terminating in a 3'-phosphate. Intriguingly, PAN is not absolutely specific for adenylate residues and can degrade non-poly(A) sequences that are bound by Pablp (Lowell et al., 1992). For example, a stretch of poly(A) with 3' non-poly(A) extensions of 7 or 32 nucleotides is hydrolyzed by PAN. Similarly, a non-poly(A) sequence corresponding to the AU-rich region of the M F A 2 3'-UTR, which appears to be bound by Pablp in vitro, is degraded by PAN (Lowell et al., 1992). These studies indicate that the substrate specificity of PAN is determined by the interaction of Pablp with RNA, and raise the possibility that PAN may degrade into the 3'-UTR of the mRNA in vivo, if it is bound by Pablp. Although under most experimental conditions PAN is strictly Pablp dependent, certain conditions render this enzyme Pablp independent. For instance, short poly(A) tails of 25-adenylate residues, but not long poly(A) oligomers, can be slowly deadenylated in the absence of Pablp (Lowell et aL, 1992). In addition, poly(A) preincubated with spermidine can be degraded by PAN with nearly 60% the efficiency found for the Pablp-dependent activity. This spermidine-stimulated activity is specific for adenylate residues (Lowell et al., 1992). Interestingly, PAN deadenylation activity can be regulated by 3'-UTR sequences in vitro. Generally, the purified PAN activity is distributive; however, in the presence of the M F A 2 3'-UTR the PAN enzyme switches
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to a processive mode of tail removal (Lowell et aL, 1992). Consistent with this in vitro regulation of PAN, the M F A 2 3'-UTR sequence is critical for determining the rate of deadenylation and stability of the M F A 2 mRNA in vivo (Muhlrad and Parker, 1992). Furthermore, Pablp appears to be required for message-specific deadenylation rates in vivo (Caponigro and Parker, 1995). For instance, in wild-type yeast, M F A 2 mRNA is deadenylated at about three times the rate of P G K 1 mRNA. However, when yeast are depleted for Pablp, rates of poly(A) shortening slow and both M F A 2 and P G K 1 mRNA are deadenylated at similar rates (1.2 ___0.5 and 1.7 +__0.4 residues per minute, respectively) (Caponigro and Parker, 1995). These observations suggest that PAN may be responsible for mRNA-specific differences in deadenylation rates. To further characterize the in vivo role of PAN, it was necessary to identify the genes encoding the enzymatic activity. Initially, the purification of the PAN enzyme identified a 135-kDa protein copurifying with PAN activity and led to the cloning of the P A N 1 gene (Sachs and Deardorff, 1992). For several reasons, P A N 1 was believed to encode the PAN enzymatic activity. The P A N 1 gene product is found in highly purified fractions of the PAN nuclease (Boeck et aL, 1996; Sachs and Deardorff, 1992). In addition, P A N 1 is an essential gene and conditional mutations in it lead to phenotypes similar to P A B 1 conditional alleles, including an arrest of translation initiation and alterations in mRNA poly(A) tail lengths (Sachs and Deardorff, 1992). Now, however, several lines of evidence indicate that P A N 1 does not encode the PAN deadenylase. For instance, antibodies directed against recombinant Panlp can efficiently immunodeplete Panlp without depleting PAN activity (Boeck et al., 1996). A newly refined large-scale purification from yeast extracts has now identified two proteins of 127 and 76 kDa, encoded by the P A N 2 and P A N 3 genes, respectively, that consistently copurify with PAN activity (Boeck et al., 1996; Brown et al., 1996). Yeast deleted for either P A N 2 or P A N 3 or for both are viable and grow as well as wild-type yeast in all growth conditions thus far tested. Several lines of evidence confirm that P A N 2 and P A N 3 are components of the PAN purified activity (Boeck et al., 1996; Brown et al., 1996). First, yeast deleted for P A N 2 or P A N 3 have longer average poly(A) tail lengths. Second, yeast extracts prepared from deleted strains have no detectable PAN activity. Most importantly, antibodies directed against recombinant Pan3p recognize a 76-kDa protein in the most pure fractions of the PAN enzyme and can immunodeplete enzymatic activity (Brown et al., 1996). Furthermore,
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immunoprecipitation of Pan3p coprecipitates Pan2p, suggesting that these two proteins interact (Brown et al., 1996). Thus, it is modeled that Pan2p and Pan3p are subunits of the Pablp-dependent poly(A) nuclease in yeast. Further analysis of the roles of P A N 2 and P A N 3 in mRNA deadenylation will be important to understand the role of the PAN enzyme in mRNA decay. Given the importance of poly(A) tail shortening in mRNA turnover, it is perhaps surprising that P A N 2 and P A N 3 are not essential for cell viability. However, strains lacking Pan2p and Pan3p have a full range of poly(A) tail lengths, and therefore it appears that PAN is not essential because other deadenylases are present in the cell. 2. Deadenylases in Other Eukaryotes
A poly(A) nuclease has been isolated from HeLa cells. This enzyme degrades poly(A) exonucleolytically and releases 5'-AMP as product (Astrom et al., 1991, 1992). In addition, it appears that the PAN enzyme may be evolutionarily conserved. Pan2p is homologous to an uncharacterized protein in Schizosaccharomyces p o m b e and humans and Pan3p is homologous to an uncharacterized protein in Caenorhabditis elegans. This putative conservation of P A N 2 and P A N 3 suggests the conservation of a PAB-dependent poly(A) nuclease in higher eukaryotes.
C. 5'---> 3' E x o r i b o n u c l e a s e s
Studies in S. cerevisiae have demonstrated a critical role for 5'---> 3' exoribonucleases in mRNA turnover. Whether a message is stable or unstable, degraded via a nonsense-mediated decay pathway or the more general deadenylation-dependent decapping pathway, 5'---> 3' exonucleases seem to be involved (see Sections II,A and II,B). The characterization of two yeast 5' --> 3' exoribonucleases, encoded by the X R N 1 and R A T1 genes, will be described below. 1. X R N I Encoding 5' - . 3' Exoribonuclease
The X R N 1 (Larimer and Stevens, 1990) gene, alias SEP1 (Tishkoff et al., 1991), D S T 2 (Dykstra et al., 1991), K E M 1 (Kim et al., 1990), R A R 5
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(Kipling et al., 1991), and SKI1 ( Johnson and Kolodner, 1995), has been independently identified by several different laboratories. Yeast deleted for the X R N 1 gene are viable but grow slowly (Larimer and Stevens, 1990). The Xrnl protein has been implicated to function in diverse cellular processes (reviewed by Kearsey and Kipling, 1991) such as DNA recombination (J. Chen et al., 1994; Dykstra et al., 1991; Holler et al., 1995; Tishkoff et al., 1991, 1995), initiation of DNA replication (Kipling et al., 1991), nuclear fusion (Kim et al., 1990), microtubule-related functions (Interthal et al., 1995; Kim et al., 1990), sporulation (Kim et al., 1990; Tishkoff et al., 1991), rRNA processing (Henry et al., 1994; Stevens et al., 1991), and mRNA turnover (Hsu and Stevens, 1993; Larimer et al., 1992; Muhlrad et al., 1994, 1995; Muhlrad and Parker, 1994). Although it is possible that the Xrnl protein has several roles within the cell, only its function as a 5' ~ 3' exoribonuclease will be discussed. Xrnlp was purified from a yeast ribosomal high salt wash based on its 5' ~ 3' exoribonuclease activity (Stevens, 1978, 1979, 1980b; Stevens and Maupin, 1987b). The purified enzyme hydrolyzes RNA in a processive manner, producing 5' mononucleotides as products (Stevens, 1979). The optimum pH for catalysis is approximately 8.0, and the enzyme requires a divalent cation. Catalytic activity is sensitive to the 5' structure of the RNA. For instance, Xrnlp greatly prefers substrates terminating in a 5'-phosphate; the same substrates terminating in a 5'-triphosphate or a 5'-OH are not efficiently degraded (Stevens, 1978, 1979). In addition, capped transcripts seem to be resistant to degradation (Stevens, 1978). The purified Xrnl ribonuclease can be stalled by the introduction of a poly(G)~s tract, which imparts a strong secondary structure to the RNA (Stevens and Poole, 1995). This observation is consistent with in vivo studies whereby insertion of a poly(G)~s cassette has been used to trap mRNA degradative intermediates (Decker and Parker, 1993; Muhlrad et al., 1994, 1995; Muhlrad and Parker, 1994; Vreken and Raue, 1992). Xrnlp seems to be the major cytoplasmic 5' ~ 3' exoribonuclease that degrades uncapped mRNA. Yeast deleted for X R N 1 accumulate deadenylated messages that lack the 5' cap structure (Hsu and Stevens, 1993; Muhlrad et al., 1994, 1995). Several messages are more stable in an X R N l - d e f i c i e n t strain. For example, in the absence of Xrnlp shortlived messages such as R P 5 1 A , C Y C 1 , and M F A 2 transcripts are approximately 2.1-, 2.2-, and 3.9-fold more stable, respectively (G. Caponigro and R. Parker, 1996, personal communication; Larimer et al., 1992). In contrast, the half-lives of the stable P G K 1 and A C T 1 transcripts are not significantly affected (Larimer et al., 1992; Muhlrad et al., 1995).
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Moreover, ribonuclease activities in addition to Xrnlp have been detected for the turnover of the P G K 1 mRNA (Muhlrad et aL, 1995; Vreken and Raue, 1992). This suggests that other ribonucleases may function together with Xrnlp in the degradation of stable mRNAs. Why might certain mRNAs, specifically unstable messages, be more susceptible to digestion by Xrnlp? It is possible that this message-specific effect is due to localization or translation differences between stable and unstable messages. Alternatively, instability elements of short-lived transcripts may recruit Xrnlp as part of an mRNase complex. This complex could contain the other mRNases needed for decay, such as the deadenylase and the decapping enzyme. The simultaneous recruitment of all of these enzymes may allow for the rapid destruction of the mRNA. Studies of the double-stranded RNA viruses L-A and M1 in S. cerevisiae also corroborate the role of X R N 1 as a 5' ---> 3' exonuclease that functions to degrade uncapped RNAs. The dsRNA virus M1 is a satellite of the L-A virus, and similar to L-A, the M~ plus-strand RNA is uncapped (reviewed by Wickner, 1991, 1992). The M1 RNA encodes a secreted toxin (killer toxin) that kills yeast that do not harbor M1 (M-o, i.e., sensitive cells). In a genetic screen that searched for increased expression of the killer toxin (super killers, or SKI mutants) (Toh-E et al., 1978), one mutant isolated, skil, contains a mutation in the X R N 1 gene (Johnson and Kolodner, 1995). Unlike many of the other SKI mutants that increase the copy number of M1 dsRNA, yeast harboring a mutation in X R N I / S K I 1 exhibited a super killer phenotype without an increase in M1 copy number (Ball et al., 1984; Ridley et aL, 1984). This increased gene expression is attributed to prolonged survival and translation of the plus-strand RNA encoding the killer toxin. Because this transcript lacks a 5' cap, it is reasoned that loss of Xrnlp function leads to transcript stabilization (Masison et al., 1995). A deletion of X R N 1 / S K I 1 is synthetically lethal with mutations in two other genes identified in this screen, S K I 2 and S K I 3 ( J o h n s o n and Kolodner, 1995). The function of the S K I 2 and S K I 3 gene products is not well understood, but they appear to repress the translation of deadenylated mRNAs (Masison et al., 1995). However, that S K I 2 and S K I 3 are synthetically lethal with X R N 1 may also suggest a role for these gene products in mRNA turnover. X R N 1 encodes an abundant 175-kDa protein that comprises 0.2% of total cellular protein (Heyer et aL, 1995). Compatible with the role of Xrnlp as an mRNase, cell fractionation and indirect immunofluorescence studies show that the majority of Xrnlp is localized to the cytoplasm
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(Heyer et al., 1995). Furthermore, Xrnlp cosediments with microtubules during sucrose cushion centrifugation, suggesting a possible interaction with the microtubule cytoskeleton (Interthal et al., 1995). Given the role of X R N 1 in mRNA degradation, it is interesting that this gene is not essential for cell viability. Although some mRNAs are more stable in the absence of X R N 1 , these transcripts are still degraded. Therefore, other mRNases that exist within the cell can compensate for the loss of Xrnlp activity. In E. coli, such a redundancy of mRNase activities is well established (reviewed by Deutscher, 1993). In summary, an understanding of many Xrnlp-related phenotypes remains elusive. It is possible that Xrnlp has many independent roles within the cell. Alternatively, many of these phenotypes could be indirect, resulting from the translation of stabilized, uncapped mRNA. This possibility is supported by the observation that in an xrnl mutant uncapped, poly(A)-deficient messages are associated with polysomes (Caponigro and Parker, 1995), and aberrant levels of protein expression are detected (Larimer et al., 1992). A more detailed characterization of the Xrnl protein will hopefully distinguish between these possibilities. 2. R A T I Encoding 5' --, 3' Exoribonuclease
A second 5'--~ 3' exoribonuclease, encoded by the R A T 1 ( H K E I , TAP1) gene, has been identified in S. cerevisiae. However, a role for this ribonuclease in mRNA turnover has not been established. R A T I is an essential gene and encodes a l l6-kDa protein. R A T I has been independently identified by several groups and exhibits pleiotropic phenotypes when mutated in yeast. R A T 1 was first identified in a genetic screen designed to isolate mutants deficient in nucleocytoplasmic trafficking of poly(A) § mRNA (Amberg et al., 1992). R A T I, also known as TAP1, has been identified as a gene that alters tRNA transcriptional activation (Di Segni et al., 1993). The R A T1 and X R N 1 gene products share regions of sequence homology (Kenna et al., 1993) and exhibit many similar properties (Stevens and Poole, 1995), consistent with the idea that they may have related functions. Similar to Xrnlp, Ratlp has been purified from a high salt wash of yeast ribosomes based on its ribonuclease activity (Stevens and Poole, 1995). Ratlp is a processive 5' ~ 3' exoribonuclease that releases 5' mononucleotides as products. The optimum pH for Ratlp catalytic activity is approximately 8.0 and catalysis requires a divalent cation.
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The strong RNA secondary structure formed by poly(G)ls, which stalls Xrnlp, also stalls Ratlp (Stevens and Poole, 1995). In addition, it has been demonstrated that both Ratlp and Xrnlp function in ribosomal RNA processing (Amberg et al., 1992; Henry et al., 1994; Stevens et al., 1991). However, the roles of Ratlp and Xrnlp within the cell do not seem to be redundant because overexpression of X R N 1 does not rescue a ratl deficiency (Kenna et al., 1993). Ratlp copurifies with a yet unidentified 45-kDa protein (Stevens and Poole, 1995). It will be interesting to determine the role of this protein with respect to Ratlp exoribonuclease activity. Although 50% of the Ratlp activity in crude extracts can be recovered from a high salt wash fraction of yeast ribosomes (Stevens and Poole, 1995), immunolocalization studies show that the majority of Ratlp appears to be localized to the nucleus (Kenna et al., 1993). The nuclear localization of Ratlp suggests that its major role within the cell is in nuclear RNA metabolism, and further studies are needed to determine if Ratlp also functions in mRNA turnover. 3. 5' --~ 3' Exoribonucleases in O t h e r E u k a r y o t e s
The 5'--> 3' exoribonuclease activities in other eukaryotes have been partially purified (Coutts and Brawerman, 1993; Lasater and Eichler, 1984; Murthy et al., 1991; Stevens and Maupin, 1987a), although none has yet been shown to function in mRNA turnover. As discussed previously, mRNA degradative intermediates in mammalian cells are difficult to detect, and thus the role of 5'---> 3' exonuclease digestion in mRNA degradation has not yet been resolved. Interestingly, both Xrnlp and Rat lp seem to be evolutionarily conserved. Antibodies directed against Xrnlp cross-react with antigens in S. pombe, Drosophila melanogaster, X e n o p u s laevis, and mouse (Heyer et al., 1995). In addition, homologs to Ratlp have been identified in S. p o m b e (Sugano et aL, 1994) and mouse (Shobuike et al., 1995). This putative conservation of the Xrnl and Rat 1 proteins suggests the conservation of the 5' --->3' exonucleolytic pathway in higher eukaryotes.
D . 3 ' - - * 5' E x o r i b o n u c l e a s e s
Of the eight characterized exonucleases in E. coli, all hydrolyze RNA in the 3'---> 5' direction (reviewed by Deutscher, 1993). In contrast, the
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majority of the exoribonucleases characterized in yeast function in the 5' --~ 3' direction, and relatively little is known about the role of 3' --~ 5' exoribonucleases in yeast m R N A turnover. Intermediates of decay shortened at the 3' end can be detected for the P G K 1 mRNA, implicating the existence of 3' ~ 5' exoribonucleases (Muhlrad et al., 1995). However, this 3' ~ 5' activity can be detected only upon mitigation of the 5' --~ 3' exonucleolytic pathway by either a 5'-UTR poly(G)18 track or by a deletion of the X R N 1 gene (Muhlrad et al., 1995). Therefore, although a 3' --~ 5' exonucleolytic pathway exists in yeast, it seems to be less active than the 5' --. 3' pathway in m R N A turnover. Two 3' --* 5' exoribonucleases have been characterized in yeast, although neither have been shown to be involved in m R N A degradation. 1. R R P 4 Encoding 3' --. 5' Exoribonuclease
The R R P 4 gene is essential for yeast viability and encodes a 39-kDa 3'--* 5' exoribonuclease with strong homology to an uncharacterized open reading frame in human cells (Mitchell et al., 1996). In vitro, the Rrp4 protein exhibits a distributive 3'---> 5' exoribonuclease activity and releases 5' mononucleotides as products (Mitchell et al., 1995). In vivo, Rrp4 is necessary for the processing of 5.8S ribosomal RNA, and yeast harboring a conditional allele of R R P 4 accumulate 5.8S rRNA species with 3' extensions (Mitchell et al., 1995). A role for R R P 4 in m R N A turnover has not been established. However, since X R N I and R A T 1 function in both rRNA processing and m R N A degradation, it is possible that R R P 4 also functions in both these processes. 2. Yeast M i t o c h o n d r i a l 3' --~ 5' Exoribonucleases
A second 3'--* 5' exoribonuclease has been isolated from the mitochondria of S. cerevisiae and is proposed to be a prominent activity involved in mitochondrial RNA turnover (Min et al., 1993; Min and Zassenhaus, 1993). Interestingly, this ribonuclease requires nucleoside triphosphates (NTPs) for activity, and any of the eight common riboor deoxyribo-NTPs will suffice. Hydrolysis of the /3-y pyrophosphate bond of the NTP seems to be important in catalysis, because no activity is detected in the presence of nonhydrolyzable NTP analogs. An ATPdependent exoribonuclease has also been identified in HeLa nuclear extracts (Murthy et al., 1991). The observation that some ribonucleases may require nucleoside triphosphates for activity is intriguing, although
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the reason for such a requirement has not been established. It is possible that NTP hydrolysis is important for "unwinding" R N A secondary structures, and thus these RNases may have an intrinsic R N A helicase activity. As of yet, no similar NTP-dependent ribonuclease has been shown to function in yeast cytoplasmic mRNA decay.
E. Endoribonucleases
Two classes of endoribonuclease activities appear to be involved in mRNA turnover. The first, sequence-specific endoribonucleases, cleave an m R N A (or a small subset of mRNAs) at a unique site. As discussed previously (see Section II,C), messages such as the yeast RPL2, the mammalian TfR and IGFII, the avian apoII and 9E3, and the Xenopus Xlhbox2B mRNA appear to be cleaved by this group of endoribonucleases. The second class of endoribonucleases has limited sequence requirements and can cleave the mRNA at multiple sites. The interferonstimulated 2-SA-dependent RNase (RNase L) is a member of this class (reviewed by Silverman, 1994). In yeast, an endoribonuclease activity with limited sequence specificity has been purified from a high salt wash of yeast ribosomes (Stevens, 1985, 1986). This enzyme prefers to cleave the bond between pyrimidine and adenosine residues ( Y - A ) , and releases products terminating with pyrimidine nucleoside 2',3'-cyclic phosphate. Whether this endoribonuclease functions in mRNA stability remains to be established. In general, very little is known about endoribonucleases and their role in yeast mRNA turnover. One difficulty has been the experimental detection of in vivo mRNA degradation intermediates resulting from endoribonuclease cleavage. It is likely that cellular exoribonucleases often degrade endonucleolytic products prior to their achieving a level that is experimentally detectable. Moreover, if cleavage of the mRNA occurs at multiple sites, the isolation of distinct 5' and 3' fragments resulting from the endonucleolytic cleavage event would be even more difficult. Certainly, with continued biochemical and genetic efforts in yeast, the significance of the endonucleolytic pathway in m R N A turnover will be revealed.
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IV. DISCUSSION AND F U T U R E D I R E C T I O N S
The integration of mRNase activities in eukaryotes to yield mRNAspecific degradation rates results from both sequence elements within the mRNA, and from the other proteins associated with the mRNA. Sequence elements within the m R N A that regulate stability can lie within the 5'-UTR, the open reading frame, or the 3'-UTR. As previously discussed, some of these can regulate deadenylation rates, others can provide endonucleolytic cleavage sites, and others can provide as yet undefined stimulation of the nonsense-mediated decay pathway. As presented in Sections III,C and II,E, several of the yeast mRNases have been purified from a high salt wash of yeast ribosomes, which suggests that these activities may be ribosome associated. However, the relationship between mRNase activities and translation remains poorly understood. For nonsense codon-stimulated degradation of m R N A through the UPF pathway, it is assumed that the UPF proteins require the ribosome in order to identify the nonsense codon (Peltz et aL, 1993). The effects of blocking translation on the degradation of normal messages is less simple to model. For instance, some mRNAs in yeast are stabilized by blocking their translation, whereas others are destabilized under the identical conditions (for example, Beelman and Parker, 1994; Muhlrad et al., 1994, 1995). It is probable that these opposing effects have a common explanation, and that this explanation will come from a detailed understanding of how mRNAs are packaged into a translatable form by mRNA binding proteins, how this mRNA-protein (mRNP) complex is traversed by the ribosome, and how the translating mRNP is recognized by cellular mRNases. For a more detailed understanding of m R N A turnover, it will be necessary to define in much greater detail its interaction with the various mRNases and regulatory factors discussed in this review. As a result, the functional reconstitution of the initial steps of the degradation reaction will need to be achieved. In particular, because the degradation rates of mRNA appear to differ due to their differential deadenylation and decapping rates, the likely choices for early reconstitution experiments will be these two events. One significant advantage of a reconstituted reaction would be the availability of an extract that could be fractionated into its integral parts, and the subsequent purification of the enzymes primarily responsible for the observed activities from these
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fractions. This contrasts to the current status of the field, whereby candidate ribonuclease activities need to be analyzed extensively through in vivo approaches to determine whether they in fact contribute to the normal degradation of the cellular mRNA. The paradigms worked out in the yeast system will certainly lead to rapid advances in studies on mRNases from other organisms. For instance, an active search for decapping activities in extracts from higher cells can now be initiated with confidence because there is ample precedent for the importance of this enzyme class in yeast. Similarly, the identification and purification of both 5' ~ 3' exoribonucleases and Pablp-dependent deadenylases, either by molecular or biochemical techniques, will lead to a greater understanding of the roles of these enzymes in the degradation of mRNAs in eukaryotes. Just as work in yeast will lead to new experiments in higher eukaryotes, so has the work in bacteria created important paradigms for studies on mRNases in yeast. For instance, the redundancy of ribonuclease function in bacteria (Deutscher, 1993) strongly suggests that, in yeast, genetic screens that search for essential functions of a ribonuclease may not succeed. Furthermore, the existence of redundant ribonucleases provides ample support for the goal of searching for ribonucleases with overlapping functions in yeast through synthetic lethal screens. Studies in bacteria have also revealed that mRNases can exist as a complex with other mRNases, and that this complex can catalyze the destruction of several different regions of the m R N A in a sequential fashion (Carpousis et al., 1994; Py et al., 1994). By analogy, it can be predicted that the mRNases involved in m R N A degradation in eukaryotes may also exist as a functional complex. Studies on eukaryotic m R N A degradation and on the mRNases involved in degradation have led to significant advances in our understanding of this important step in the posttranscriptional control of gene expression. If the rate of progress over the past few years in this area is an indication of things to come, then we can all look forward to a detailed description of this facet of m R N A biology in the near future.
ACKNOWLEDGMENTS
We thank the membersof the Sachs Laboratoryfor helpful comments on the manuscript. This work was supported by NIH Grant R01-GM50308 and in part by funds from the Searle Scholars Program to ABS.
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19 Ribonuclease Inhibitor JAN H O F S T E E N G E Friedrich Miescher-Institut CH-4002 Basel, Switzerland
I. Introduction II. Biological Properties A. Species and Tissue Distribution B. Biological Function of Ribonuclease Inhibitor C. Molecular Biology of Ribonuclease Inhibitor III. Molecular Properties A. RNasc-RI Interaction B. Primary and Tertiary Structure IV. Applications A. Protection of RNA B. RNasc Assay C. Inhibition of Angiogenesis Refcrenccs
1. I N T R O D U C T I O N
In 1952, P i r o t t e a n d D e s r e u x d i s c o v e r e d in the s u p e r n a t a n t of g u i n e a pig liver an activity that inhibited R N a s e s ; this m a r k e d the b e g i n n i n g of r e s e a r c h to u n d e r s t a n d the biological f u n c t i o n a n d structural p r o p e r t i e s of the r i b o n u c l e a s e i n h i b i t o r ( R I ) . T h e R I p r o t e i n inhibits R N a s e s of the p a n c r e a t i c type ( B e i n t e m a et al., 1988), a n d m o s t studies h a v e used e n z y m e purified f r o m m a m m a l i a n tissues. R I is an acidic 5 0 - k D a p r o t e i n , characterized by a high c o n t e n t of leucine a n d cysteine residues ( B l a c k b u r n et RIBONUCLEASES: STRUCTURESAND FUNCTIONS
621 Copyright 9 1997by AcademicPress,Inc. All rightsof reproductionin any formreserved.
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Jan Hofsteenge
al., 1977). It is not structurally related to the recently identified inhibitor of 2-5A-dependent RNase (Bisbal et al., 1995), nor to the inhibitors of microbial RNases (Chapter 2). Following the initial discovery of this activity, much research was devoted to identifying the nature of the inhibitor molecule and its properties (Roth, 1956, 1958a,b, 1962; Shortman, 1961, 1962b), as well as to its distribution in different animals and tissues (Roth, 1962; Kraft and Shortman, 1970a). In a large number of studies the changes in RI levels in tissues, due to physiological stimuli such as development, disease, and hormones, have been correlated with changes in R N A metabolism, allowing deduction of a biological role for RI. Purification of the protein was, for a long time, a challenging undertaking because of the low abundance of RI in most tissues and its lability, in particular due to thiol modifications (Gribnau et aL, 1969). The use of affinity chromatography, with bovine pancreatic RNase A as the ligand, finally led to purification to apparent homogeneity ( G r i b n a u e t a l . , 1969; Gagnon and de Lamirande, 1973; Graveau et aL, 1974; Blackburn et aL, 1977; Blackburn, 1979). This allowed the detailed study of its interaction with a number of RNases (Lee etal., 1989a,b; Vicentini etal., 1990; Shapiro and Vallee, 1991), the determination of its primary structure (Hofsteenge et al., 1988), the sequencing and recombinant expression of its cDNA (Lee et al., 1988; Schneider et al., 1988; Lee and Vallee, 1989b; Vicentini et al., 1990; Kawanomoto et al., 1992), and the determination of the three-dimensional structure of free porcine RI, as well as the complex with bovine pancreatic RNase A (Kobe and Deisenhofer, 1993, 1995). These developments have previously been discussed in several reviews (Roth, 1967; Blackburn and Moore, 1982; Lee and Vallee, 1993; Shapiro et al., 1995). Despite the fact that many reports on RI have appeared, a number of questions remain. The most intriguing and probably most important one concerns the true physiological role of this protein. At the structural level, interesting questions related to the specificity of protein-protein interactions, as well as to the function of the many thiol groups present in the RI molecule, may now be addressed. It is hoped that this chapter may provide some of the background for such studies.
Ii. BIOLOGICAL PROPERTIES A. Species and Tissue Distribution
The presence of RI in biological samples has generally been determined using two types of assays. First, RI present in its free form in
19
Ribonuclease Inhibitor
623
extracts or homogenates has been detected by adding a known amount of bovine pancreatic RNase A, followed by determination of the remaining activity, using natural (Shortman. 1961) o r artificial substrates (Blackburn. 1979). Although this type of assay can yield quantitative information. it is not universally applicable. R I from many mammalian sources inhibits RNase A (Roth, 1962)-but this is not the case for RI from birds (Kraft and Shortman. 1970a; Dijkstra er al., 1978). amphibians (Roth, 1962; Malicka-Blaszkiewicz and Kubicz. 1979; Kraft and Shortman, 1970a), and trypanosomes (Gbenle. 1990). The presence of RI may be overlooked unless RNases from the same organism are used. The second assay detects RI in complex with endogenous RNases. From the difference in RNase activity before and after inactivation of RI. often by thiol-blocking reagents such as p-hydroxymercuribenzoate (Roth. 1956; Shortman, 1962b) o r N-ethylmaleimide (Girija and Sreenivasan, 1966). the presence of RI is deduced. More recent methods to detect RI have used specific antibodies (Burton and Fucci. 1982; Yasuna and Goto. 1986: Bloemendal and Janscn, 1988; Miura et rrl., 1988; Feldman rf ul., 1988) or activity staining of polyacrylamide gels with an over- and underlay containing RNA and RNase A. respectively (Nadano rr al., 1995). Using either onc o r both of the enzymatic methods dcscribed above, R1 ac~ivityhas bcen detected in mammals (Kraft and Shortman. 1970a). birds (Kraus and Scholtissek. 1974; Dijkstra rr ol., 197.5). amphibians (Nagano ct al., 1976: Malicka-Blaszkicwicz and Kubicz. 1979). insects (Aoki and Natori. 1981; Garcia-Segura el nl., 1985). and a parasitc (Ghenlc. 1990). Despite this widesprcad occurrence. Ihe inhibitor has bcen purified only from mammalian sourccs. i.e.. human placenta and brain (Blackburn pr ~ t l . ,1977; Nadano er (ti., 1994). porcine and bovine liver and brain (Burton rr ul., 1980; Burton and Fucci. 1982; Cho and Joshi, 1989). rat liver and testis (Burton and Fucci. 1982: Fominaya rf al., 1988a), and mouse and sheep liver (Burton and Fucci. 1982). These proteins are very similar wilh respect to size, amino acid composition and immunological properties (Burlon and Fucci. 1'382). R1 from nonmammalian sources has not been characterized in such detail. and the possibility thal both types of RI arc structurally unrelated cannot bc excluded. Generally. they share with the mammalian inhibitors the sensitivity toward thiol-modifying reagents (Malicka-Blaskiewicz and Kubicz. 1981: Ghenle, 1990). but may be different in other respects. For instance. it has been reported that the inhibitor from the bull frog (Runu cateshriana) has a much higher molecular weight than the one from mammals (Nagano ut ul., 1976). More work is needed to clarify the relationship of these proteins to mammalian RI.
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Jan Hofsteenge
Although an inhibitory activity toward RNase A has been reported in plants (Bernheimer and Steele, 1957), this seems to be due to a nonproteinaceous inhibitor. In developing apple leaves the presence of an inhibitor has been deduced from the effect of p-chloromercuribenzoate on the RNase activity (Kessler and Engelberg, 1962). Further studies are required to establish whether plants indeed contain an RNase inhibitor protein. RNase inhibitors have also been found in fungi and bacteria, but these proteins are structurally distinct from animal RI (see Chapter 2). The distribution of RI in tissues of higher animals has mainly, but not exclusively, been investigated in the rat. It has been found in the major organs (liver, kidney, lung, pancreas, and spleen (Roth, 1956; Kraft and Shortman, 1970a; Chesters and Will, 1978; Kiyohara and Menjo, 1983; Morisset et al., 1972), in heart and skeletal muscle (Roth, 1956; Kraft and Shortman, 1970a; Little and Meyer, 1970), in the central and peripheral nervous system (Takahashi et al., 1970; Burton et al., 1980; Bates et al., 1985; Cho and Joshi, 1989), in the reproductive organs (Hilz et al., 1968; Bard6n et al., 1969; McGregor et al., 1981; Kumagai et al., 1991; Rao et al., 1994), in all glands examined (Imrie and Hutchison, 1965; Robinovitch et al., 1968; Kraft and Shortman, 1970a; Greif and Eich, 1971; Liu et al., 1975c), in the eye lens (Ortwerth and Byrnes, 1971), in the esophagus (Chesters and Will, 1978), and in adipose tissue (Eichel et al., 1961). RI also occurs in blood cells (Priess and Zillig, 1967; Kraft and Shortman, 1970a,b; Goto and Mizuno, 1971; Bloemendal and Jansen, 1988), but it is absent from extracellular fluids (Roth, 1956; Nadano et al., 1994). Its distribution within an organ has not been investigated in detail. Based on the presence of its activity in the high-speed supernatant of tissue extracts, and its absence from other subcellular fractions, RI is thought to be located in the cytoplasm (Roth and Juster, 1972). However, other studies have reported RI activity in purified nuclei (Chakravorty and Busch, 1967a; Skridonenko, 1973) or in cytoplasmic ribonucleoprotein particles (Gileadi et al., 1984). A detailed examination of the cellular and subcellular distribution of RI, using in situ probes such as antibodies and cRNA, may be worthwhile.
B. Biological Function of Ribonuclease Inhibitor
The observation that RI, or at least its activity, occurs in a wide range of animals and in every tissue or cell examined suggests an essential role
19
Ribonuclease Inhibitor
625
for this protein. Nevertheless, no direct experiments addressing this question have been published. Based on a large variety of observations, three hypotheses have been proposed: (1) RI inhibits intracellular (cytoplasmic) RNases that are involved in the regulation of the amount of the different kinds of RNA. High levels of RI will cause RNA accumulation, whereas low levels favor its degradation (Shortman, 1962a; Roth, 1962). (2) RI is a safeguard against RNases that are destined for secretion, but inadvertently appear in the cytoplasm (Beintema et al., 1988). (3) RI regulates or terminates the physiological action of RNases such as angiogenin, or the eosinophil-derived enzymes (Lee et al., 1989b; Nadano et al., 1994). The first hypothesis is based on the observation that the amount of RI compared to that of RNase is high in tissues with a high rate of R N A synthesis and accumulation (Shortman, 1962a), a situation encountered in cells that are rapidly dividing or are active in protein synthesis. In contrast, in tissues with a high catabolic activity this ratio is low (QuirinStricker et al., 1968). The effects of many natural stimuli~for example, growth and development, disease, and h o r m o n e s ~ o n the level of RI have been examined, and the results often support the hypothesis (Table I). For instance, the phytohemagglutinin-stimulated proliferation of cultured lymphocytes is accompanied by an increase in RI (Kraft and Shortman, 1970b). Furthermore, the increase in RNA and protein synthesis in the mammary gland during pregnancy and lactation correlates with an increase in RI (Liu et al., 1975b), whereas at involution it decreases. An example of such a correlation in a disease state is found in hyperthyroidism (Greif and Eich, 1977). Physical stimuli or chemicals that affect cellular functions have also provided evidence in favor of this hypothesis (Table I). However, contradictory results have been obtained as well. In particular, the changes in RI level observed in tumors are not always consistent. Whereas an increase is observed in the thymus of AKR mice (Kraft and Shortman, 1970a) and in mammary tumors (Liu et al., 1975b), a decrease has been seen most of the time in a series of transplantable hepatomas in rat (Roth et al., 1964; Graveau et al., 1974). Importantly, there seems to be no correlation between the level of RI and the rate of cell proliferation (Roth et al., 1964). In addition, the amount of RI in lymphocytes of patients with chronic lymphocyte leukemia is not altered in a consistent fashion (M~nsson et al., 1974). Interpretation of these kind of studies has been complicated by the nature of the assays used to determine RI. The results summarized in Table I give the amount of free RI, the rest being bound to endogenous
TABLE I Conditions Affecting the Level of Ribonuclease Inhibitor ~
Condition and tissue/cell I. Development and aging 1. Rat liver 2. Rat mammary gland
Stimulus
3. Rat uterus
Pregnancy Pregnancy Lactation Involution Maturation
4. Rat prostate
Aging
5. 6. 7. 8.
Aging Aging Aging Aging
Rat brain Rat liver Chicken liver Rat and human lymphocytes
9. Rat liver, thymus, lymph nodes 10. Rat liver 11. Fleshfly fat-body 12. Cells in culture II. Disease 1. Rat transplantable hepatomas Rat hepatomas 2. Mouse thymus 3. Rat liver 4. Rat mammary gland
RI h
Aging Regeneration after partial hepatectomy Pupation Proliferation Tumor growth Tumor growth Leukemia Implanted carcinoma Tumor growth
Remarks
Increases up to day 16 after conception
Goodlad and Ma (1975) Liu et al. (1975b)
Results from an increase in RNase (see below, III.8) Maximum at 4 weeks after birth (see below, III.12) Maximum at day 5-10 after birth Increases up to day 10 after birth Measured from 2 days before until 450 days after birth (see below, IV.I) Measured from 9 to 52 weeks after birth
Zan-Kowalczewsk and Roth (1975) Munro and Knowler (1982) Kumagai et al. (1991)
Peaks before onset of logarithmic phase
-(5) +(2)
+
Suzuki and Takahashi (1970) Liu et aL (1975a) Sarkar and Pastro (1971) Kraft and Shortman (1970b) Kraft and Shortman (1970a) Shortman (1962a) Chakravorty and Busch (1967a) Aoki and Natori (1981) Garcfa-Segura et al. (1991)
Compared to normal liver; different hepatomas give different responses
Roth et al. (1964)
Walker 256 carcinoma
Graveau et aL (1974) Kraft and Shortman (1970a) Clark and Goodlad (1983) Liu et al. (1975c)
+ n.c.
Refs.
Chronic lymphocytic leukemia Tumor growth Alzheimer's disease
+/n.c.
Alzheimer's disease Alzheimer's disease Alzheimer's disease Hyperthyroidism Cataract Cataract Cataract Muscular dystrophy
n.c. n.c. n.c. +
n.c. + n.c. n.c. n.c.
8. Rat uterus
Hypophysectomy Estradiol Corticosteroids 9c~-Fluoroprednisolone 9o~-Fluoroprednisolone 9o~-Fluoroprednisolone Triiodothyronine Thyroid-stimulating hormone Estradiol
9. Rat uterus
Estradiol
5. Human lymphocytes 6. Rat implanted tumors 7. Human brain
8. Human thyroid 9. Human, rat, and bovine eye lens 10. Mouse muscle III. Hormones 1. Rat liver 2. Rooster liver 3. Rat liver 4. Rat liver Rat thymus 5. Rat lymphosarcoma 6. Rat, mouse thyroid 7. Rat, mouse thyroid
+
Differs from patient to patient
Mhnsson et al. (1974)
In nuclear supernatant
Chakravorty and Busch (1967a) Sadjel-Sulkowska and Marotta (1984) Morrison et al. (1987) Maschhoff et al. (1989) Jones and Knowler (1989) Greif and Eich (1977) Ortwerth and Byrnes (1971) Ortwerth and Byrnes (1972) Cavalli et al. (1979) Little and Meyer (1970)
w
Decrease due to increase of RNase
In vitro experiments on cytosol
Brewer et al. (1969) Dijkstra et al. (1978) Sarkar (1969) Ambellan (1980)
+
In thiouracil-induced goiters (see below, IV.4 and V.1)
Ambellan and Hollander (1968) Greif and Eich (1972) Murthy and McKenzie (1974)
+
Immature rats: total RI increases as shown immunologically; the decrease in free RI is due to an increase in RNase (see above, 1.3) Mature ovariectomized rats
w
McGregor et aL (1981) Brockdorff and Knowler (1986)
Schauer (1981) Schauer (1991) (continued)
TABLE I ( C o n t i n u e d ) Condition and tissue/cell
Stimulus
RI b
Ovariectomy Ovariectomy + estradiol 11. Rat vaginal epithelial cells Estradiol 12. Rat prostate Castration Castration +testosterone 13. Rat adrenal gland ACTH IV. Compounds affecting cellular functions 1. Human lymphocytes Phytohemagglutinin 2. Human lymphocytes 12-O-Tetradecanoylphorbol- 13-acet ate 3. Human amnion cells Cyclohexamide Trenimon + 4. Rat, mouse thyroid Theophylline + Dibutyryl-cAMP + Thiouracil 5. Rat kidney Puromycinaminonucleoside+
Remarks
10. Mouse uterus
Refs. Zan-Kowalczewsk and Roth (1975)
Immature rats; total RI was measured
Rao et al. (1994) Kumagai et al. (1991)
Peaks at 4 days Imrie and Hutchison (1965) In culture In culture; in particular B lymphocytes were examined In culture; both compounds inhibit cell proliferation
Kraft and Shortman (1970b) Kyner et al. (1979) Hilz et al. (1968)
In vitro
Murthy and McKenzie (1974)
Nephrotoxic
Greif and Eich (1972) Nicholls and Markle (1974) Bishay and Nicholls (1973)
6. Rat liver 7. Rat liver 8. Mouse liver 9. Mouse implanted tumor 10. Mouse muscle and liver 11. Mouse muscle 12. Mouse ascites cells V. Diet 1. Rat, mouse thyroid 2. Rat liver 3. Rat liver 4. Pigeon pancreas VI. Physical stimuli 1. Mouse liver Mouse thymus 2. Guinea pig liver 3. Rat cerebral tissue 4. Rabbit muscle
2-Acetamidofluorene Thioacetamide Endoxan 5-Fluorouracil Poly(I)/Poly(C) Botulinum toxin Actinomycin D
+ -
Hepatocarcinogen Carcinogen Alkylating agent
-
Muscle denervation (see below, VI.4)
Low iodine content Protein free Fasting Fasting
+ -
X-ray X-ray X-ray Electroconvulsive shock Immobilization
n.c. n.c. -
See above, 111.6 and IV.4 --
Wojnar and Roth (1965) Chakravorty and Busch (1967b) Comorosan et a t (1968) Ambellan and Hollander (1968) Meyer and Meyer (1979) Kertai and H611osi (1984) Von Tigerstrom (1972) Murthy and McKenzie (1974) Quirin-Stricker et al. (1968) Onishi (1970) Morisset et al. (1972) Kraft et al. (1969)
n D n Total RI was measured
Ferencz et al. (1973) Park et al. (1977)
Both in the soleus and gastrocnemieus
Kiss and Guba (1979)
a The change in free RI is given, unless indicated otherwise. b +, An increase in RI level; - , a decrease in RI level; n.c., no change. In entree II.1 the number of cases is given in brackets.
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Jan Hofsteenge
RNases. Free R I can, in principle, be determined accurately, but whether this reflects the true situation in the cytoplasm is questionable. In most studies it has been implicitly assumed that, like RI, the RNases in the RI-RNase complexes also originate from the cytoplasm. This is most likely not the case. Purification and structural characterization of such RNases from porcine liver revealed that they belong to the secreted, pancreatic type, which is normally not expected to occur in the cytoplasm (Hofsteenge et al., 1989). This is underscored by the cDNA sequence for one of these proteins, which contains a signal sequence for transport over the endoplasmic reticular membrane (Vicentini et al., 1994), and by the presence of the protein in plasma (Zhou and Strydom, 1993). It seems probable that the major fraction of the RNase-RI complexes in tissue extracts does not exist in the cytoplasm, but that it forms during tissue disruption and fractionation. The amount of free RI will therefore depend on the methods used, as well as on the amount of pancreatic type of RNasc present in the tissue. In this context it is important to note that intracellular RNases that have been implicated in RNA metabolism, and that differ from the secreted RNases, e.g., in their size and requirement lor metal ions, are generally not inhibited by R1 (Kumagai et i l l . , I980; Stollc an Bcnz. 1988; Pastori ct o/., 1991; Hua et d., 1903; Herrera et ( I / . , 199s). Unfortunately. no structural data arc available o n these enzymes. Measurement of the amount of RI in RI-R complexes. as detcrmincd from the RNasc activity released by inactivation of RI, is unreliable becausc RNases with widely different specific activities bind to RI (Sect i o n III,B,3,c). Without knowledge about which RNases are present in the complexes, their relative amounts, and their specific activities, no conclusion can be drawn. Another level of complication in many of these studies is that RI has been measured in tissue extracts. I t is often not known whether RI and the RNases occur in the same cells (Brockdorff and Knowler, 1986). From these analyses, it becomes clear that the determination of RI requires more rigorous methods. The advantage o f immunological detection of RI is demonstrated by the work of Knowler and colleagues (McGregor rt a/., 1981; M u n r o and Knowler, 1982; Brockdorff and Knowler, 1986, 1987). The second hypothesis relies on the fact that only secreted RNases bind tightly to RI (see Section IV,A). No experimental evidence directly supporting this hypothesis has been published. In fact, a number of observations argue against it. The very high affinity of RI for RNase
19
Ribonuclease Inhibitor
631
and its large molar excess in most tissues, which increases even further under certain circumstances (Table I), are difficult to reconcile with this hypothesis. Also, the following observation is unexpected. Human HL60 cells contain very low levels of RNase 2, an enzyme known to be inhibited by RI (Section III,A,1). The total amount of RI in these cells is even lower than that of RNase 2. In such a situation one would expect the level of a safeguarding inhibitor to rise in conjunction with increased production of the enzyme. However, induction with butyric acid (Tiffany et aL, 1995) causes a large increase of RNase 2, as demonstrated immunologically, but the amount of RI antigen remains constant (M. A. Doucey, M. Bl~izquez, and J. Hofsteenge, 1995, unpublished results). The third hypothesis also relies on circumstantial evidence. Angiogenin induces neovascularization at femtomolar concentrations, and it has to enter the cell to exert its action (Fett et al., 1985; Moroianu and Riordan, 1994). Any intracellular inhibitor that would regulate or terminate its activity should, therefore, bind to this enzyme with a very high affinity. Human RI binds angiogenin with a subfemtomolar dissociation constant, and would thus be a good candidate for such a regulator (Lee et al., 1989b). In summary, it is clear that the biological function of RI is still not known with certainty. Definitive answers may be expected in the future, because the tools to perform more direct experiments arc now available in the form of specific antibodies and cDNA probes.
C. Molecular Biology of Ribonuclease Inhibitor The cDNA coding for RI from human placenta (Lee et al., 1988), HeLa cells (Schneider et al., 1988), porcine brain (Vicentini et al., 1990), and rat lung (Kawanomoto et al., 1992) has been obtained. So far no gene structure for the RI protein has been reported. Crawford et al. (1989) have demonstrated that in human placenta alternative splicing of an exon in the 5' untranslated region of the m R N A occurs. Interestingly, one of the splice forms corresponds exactly in length to the proteins from pig and rat, which are N-terminally four amino acids shorter than RI isolated from human placenta. The function of the differential splicing remains to be elucidated, but it does not seem to affect the affinity of RI for the enzyme (Section III,A,1).
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Jan Hofsteenge
The gene encoding human RI ( R N H ) has been localized to chromosome 11p15.5 (Weremowicz et al., 1990; Zneimer et aL, 1990). Long-range restriction mapping placed it close to the Harvey ras protooncogene ( H R A S 1 ) (Schneider et aL, 1992), a finding that was refined by Weitzel and Patel (1994), who established the gene order to be H R C 1 , H R A S 1 , R N H ( H R C 1 encodes a transcription factor). In fact, R N H localizes within 30-50 kb of the 3' end of H R A S 1 . Alterations in chromosome 11p15.5 are commonly found in human cancers, and because genes that are functionally related often are also physically linked, this could indicate that RI is involved in cell growth and differentiation (Weitzel and Patel, 1994). It would be of interest to determine whether this synteny has been conserved in other animals. The chromosomal localization of RI, the occurrence of multiple splice forms of the mRNA, and the variety of physiological stimuli that alter RI levels (Section II,B) seem to warrant a more detailed investigation of the mechanism of regulation of this protein.
III. MOLECULAR PROPERTIES A. R N a s e - R l Interaction
1. Mechanism and Kinetics
RI inhibits RNase by forming a complex, as was realized early on (Roth, 1956); this was firmly established by affinity chromatography on immobilized RNase (Gribnau et al., 1970). Although initial studies employing partially purified RI indicated that the enzyme/inhibitor stoichiometry was 2:1 (Fujisawa, 1972b), a variety of approaches, using pure RI, yielded a ratio of 1:1 (Blackburn et al., 1977; Lee et al., 1989a; Vicentini et al., 1990). The reported values for the dissociation constant of the RI-RNase A complex span four orders of magnitude, and different modes of inhibition have been proposed. The initial findings with purified human and porcine RI showed noncompetitive inhibition, with an inhibition constant for RNase A of approximately (3-7) x 10 -1~ M (Blackburn et al., 1977; Burton et al., 1980; Cho and Joshi, 1989). These results were obtained from Lineweaver-Burk plots, with enzyme and inhibitor concentrations in the nanomolar range. Turner et al. (1983) pointed out
19
633
Ribonuclease Inhibitor
that this approach did not take into account inhibitor depletion and that this and the fact that the Michaelis constant for the substrate was much larger than Ki lead to apparent noncompetitive inhibition kinetics. Similar considerations were made by Fominaya et al. (1988b) in the analysis of purified RI from rat testis, yielding an inhibition constant of 3.2 • 10 -12 M. The use of R N A in these studies posed a limitation with respect to substrate homogeneity, as well as sensitivity of determination of RNase activity. Lee et al. (1989a) followed an entirely different approach to measure the association of angiogenin and RNase A with human RI. Making use of the 50% increase in tryptophan fluorescence on complex formation between angiogenin and human RI, they used stopped-flow techniques to study the association process. From the hyperbolic dependence of the pseudo-first-order rate constant on inhibitor or enzyme concentration they concluded a two-step mechanism: k_ 1
k_ 2
E + I -----El -----El* k,1 k+2 Initially, a loose complex (K1 -- k-l/k+l ~ 5 • 10 -7 M) is formed rapidly, followed by a slower conversion (k2 ~ 100 sec -~) into a more stable complex (El*). The overall second-order association rate constant, kass, was found to be 1.8 x 10s M -~ sec -1 (Lee et al., 1989a). The dissociation rate constant, kois, was determined separately by measuring the rate of release of angiogenin from the complex in the presence of a scavenger to prevent reassociation (Lee et al., 1989b). An extremely low value, k-2 -- 1.3 • 10 -7 sec -l, was found, corresponding to a half-life of 62 days for the RI-angiogenin complex. The inhibition constant, Ki, calculated from these rate constants is 7.1 • 10 -16 M. The association rate constant for RNase A has been determined to be 3.4 • 108 M -1 sec -1, based on a competition experiment using angiogenin (Lee et al., 1989a). Together with the dissociation rate constant of 1.5 • 10 -5 sec -1 this yielded an inhibition constant of 4.4 • 10 -14 M (Lee et al., 1989b). The binding was concluded to be competitive from the effect of 2'-CMP on the association rate constant. Vicentini et al. (1990) measured the progress curve of cleavage of the dinucleotide U p A by RNase A in the presence of varying amounts of porcine RI. Analysis of the data according to the equation for slow, tight-binding inhibition kinetics (Morrison and Stone, 1985) yielded values for the three kinetic parameters that
634
Jan Hofsteenge
w e r e very c o m p a r a b l e to those o b t a i n e d for h u m a n R I ( T a b l e II). F r o m the effect of the s u b s t r a t e c o n c e n t r a t i o n on K~ and kass, it could be c o n c l u d e d that also the association of R N a s e A with R I follows a twostep m e c h a n i s m and that a c o m p e t i t i v e m o d e of inhibition applies. T h e s t r e n g t h of binding strongly d e c r e a s e s with increasing ionic strength, as well as with increasing p H ( L e e et al., 1989a,b; Vicentini et aL, 1990). Using e i t h e r of these e x p e r i m e n t a l protocols, the kinetic c o n s t a n t s for the inhibition of h u m a n R N a s e 2 with h u m a n R I ( S h a p i r o and Vallee, 1991) and porcine R N a s e 4 with p o r c i n e R I (U. N e u m a n n , O. Z e l e n k o , and J. H o f s t e e n g e , 1994, u n p u b l i s h e d results) have also b e e n d e t e r m i n e d (Table II). G i v e n the very low values of Ki, s o m e discrepancies in the l i t e r a t u r e b e c o m e u n d e r s t a n d a b l e . For instance, the conclusion f r o m e n z y m o l o g i cal studies that the S-peptide of R N a s e A does not c o n t r i b u t e to binding RI ( B l a c k b u r n and Jailkhani, 1979) is in d i s a g r e e m e n t with the finding that RI p r o t e c t s Lys-7 from chemical modification ( B l a c k b u r n and Gavilanes, 1982). This can be explained by the fact that in the e n z y m o l o g i c a l e x p e r i m e n t s the c o n c e n t r a t i o n of R N a s e A was much h i g h e r than K~ ( B l a c k b u r n and Jailkhani, 1979). Using m u c h lower e n z y m e c o n c e n t r a tions, the S-peptide was found to c o n t r i b u t e 16 kJ/mol to the binding e n e r g y ( N e u m a n n and H o f s t e e n g e , 1994). 2. I n h i b i t o r s
of Interaction
A l t h o u g h the R I - R N a s e interaction is a reversible one, the half-life of dissociation of the c o m p l e x is m e a s u r e d in tens of hours, or even in
TABLE I! Kinetic Parameters for Inhibition of Various RNases by Ribonuclease Inhibitor a
Enzyme
RI
k.... (M I sec i)
RNase A RNase A Angiogenin RNase 2
Human Porcine Human Human
3.4 1.7 1.8 1.9
x • x •
10~ 10~ 10~ 1()x
RNase 4
Porcine
1.5 • 1()x
kai~ (sec l) 1.5 9.8 1.3 1.8
x x • x
10 s 10 6 10 7 10 7
6.1 • 10 7
Ki (M) 4.4 5.9 7.1 9.4
x x • x
Refs.
10 10 10 10
14 Lee et al. (1989a,b) ~4 Vicentini et al. (1990) ~' Lee et al. (1989a,b) ~6 Shapiro and Vallee (1991) 4.0 x 10 ~s t,
"The parameters were determined at pH 6.0, I = 0.15, and 25~ t' U. Neumann, O. Zelenko, and J. Hofsteenge, 1994, unpublished results.
19 Ribonuclease Inhibitor
635
days. A number of compounds have been found, however, that lead to inactivation of RI and/or the dissociation of the RI-RNase complex. From the very early studies it is known that acid readily inactivates RI (Pirotte and Desreux, 1952). Also the sensitivity toward thiol-modifying reagents has been extensively documented (Blackburn and Moore, 1982). Both Cu(II) and Zn(II) prevent the formation of the human R I angiogenin complex (Badet et aL, 1991), which in the case of Zn(II) could result from an effect on the inhibitor (Fominaya et al., 1988a). An effect of these ions on angiogenin is possible as well. The inhibition of complex formation between bovine brain RNase and RI by AI(III) is due to conformational changes in the RNase (Cho and Kim, 1991). Finally, antibodies specific for RI dissociate the complex with RNase (Yasuna and Goto, 1986). Few physiological inactivators of RI have been described. Oxidized glutathione inactivates RI, probably by forming mixed disulfides (Fominaya et al., 1988a), whereas cathepsins can proteolytically inactivate RI (Fuhge and Otto, 1980). This process may well explain the finding of an RI-inactivating factor in rat spleen, but not in liver (Fujisawa, 1972a).
B. Primary and Tertiary Structure 1. P r i m a r y S t r u c t u r e
The amino acid sequences of RI from pig (Hofsteenge et al., 1988), human (Lee et al., 1988; Schneider et al., 1988), and rat (Kawanomoto et al., 1992) cells share 75-77% sequence identity (Fig. 1). The protein from pig and rat cells consists of 456 residues, whereas the human one is 460 residues long. The polypeptide chain is N-terminally blocked with an acetyl moiety (Hofsteenge et al., 1988). The primary structure of RI reveals two interesting properties. First, it consists nearly completely of leucine-rich repeats (LRRs). Second, it contains an unusually large number of cysteine residues that all occur in the sulfhydryl form. LRRs are sequences of 24-29 amino acids that contain leucine (or another hydrophobic amino acid) at constant positions (Fig. 1). They have been found in proteins with a diverse range of functions, including enzyme inhibition or activation, DNA repair, RNA processing, signal transduction, extracellular matrix interactions, and bacterial virulence. Furthermore, this structural motif has been found in organisms ranging from bacteria to humans. All LRR-containing proteins seem to be in-
IS
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X L E X L X L X X C X L T X X X C X X L X X a L X X X X <... helix .> X L R E L X L X X N X L G D X G a X X L X X X L X X P X X <. helix .>
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90 100 K I Q K L S L Q @ C S L T E A G C G V L P S T L R S L P D V . . . . S C G . . . . . . S . . . . T . .
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110 120 130 T L R E L H L S @ N P L G D A G L R L L C E G L L D P Q C N E . . K . . . . . . R . . . . . . Q L . . . A . . Q . . . . . . . . . . .
. . . . . . . . . . . . . . . . . . . .
140 150 160 H L E K L Q L E Y C R L T A I S C E P L A S V L R A T R R N . . . T . . . . . . . . . . V I P R S . S . . . . . . . . . . . . . K P
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170 180 190 A L K E L T V S N N D I G E A G A R V L G Q G L A D S A C D F . . . V L . . . . F H . . . I H T . C . . . K . . . . D F T . . . . . . N . . . V . . . C . . . K . . P .
82
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200 210 210 Q L E T L R L @ N C G L T P I N C K D L C G I V A S Q A 9 . K I . S . . . . . . . D V . . . K . . . .A.K..S..V.SD..R...G....K.
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230 240 250 S L R E L @ L G S N G L C D A G I A E L C P G L L S P A S Q K . . N T . . . A . . S . . . L . S C A . . . . K . . . V . Y . . . . P . . . E . S .
.. . . . . . . . . . . . .
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.........
A4
85
Fig. 1. Comparison of the primary structures of porcine, human, and rat RI. (Top): Consensus sequences of the A- and B-type leucine-rich repeat, with the most conserved regions underlined (Kobe and Deisenhofer, 1994). (Bottom):Only residues that are different from those in porcine RI have been shown. Contact residues to RNase A in the enzyme-inhibitor complex have been circled.
19
Ribonuclease Inhibitor
637
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Fig. 1
(Continued)
volved in specific macromolecular interactions, suggesting that the LRR provides a versatile framework onto which specialized functions can be grafted. An excellent review of this class of proteins has been published (Kobe and Deisenhofer, 1994). It is important to note that LRRs are structurally not related to the 39-amino acid repeats of the 65-kDa regulatory subunit of protein phosphatase 2A, which also contains aliphatic residues at constant positions (Hemmings et al., 1990), nor to the leucine zipper (Branden and Tooze, 1991). RI consists of 16 homologous LRRs of which the 14 central ones alternately belong to the A type, characterized by the sequence CXLTXXXC, and the B type, with the sequence N X L G D X G a at positions 10-17 (Fig. 1, where "a" stands for aliphatic). The N- and Cterminal repeats (denoted N and C in Fig. 1) are less homologous and cannot be assigned to either type. It is difficult to deduce the tertiary structure of the LRR or the way in which such repeats assemble from the primary structure alone. Attempts using model peptides based on the LRRs of chaoptin (Krantz et al.,
638
Jan Hofsteenge
1991) and Toll protein (Gay et al., 1991) indicated a preponderance of /3 structure, but the results may have been influenced by the solubility properties of the hydrophobic peptides. RI from pig, human, and rat cells contains 30, 32, and 30, cysteine residues, respectively, 27 of which have completely been conserved (Fig. 1). Of these, 14 occur in the CXLTXXXC motif of the A-repeat and 11 occupy positions 21 and 29 of the A- or B-repeat. The other two conserved cysteines (Cys-7 and Cys-404) contact RNase A in the enzymeinhibitor complex (Section III,B,3,a). The conservation of so many cysteines, both between species and with respect to their position in the LRR, suggests an important function. In addition to being structurally important (Section III,B,2) it has been proposed that the cysteines provide a target for the inactivation of RI in physiological processes (Aoki and Natori, 1981; Fominaya et al., 1988b). In vitro RI can readily be inactivated by thiol-modifying reagents (Roth, 1956; Blackburn et al., 1977). In particular, the oxidation with 5,5'-dithiobis(2-nitrobenzoic acid) has been studied in some detail. Complete inactivation of porcine RI is accompanied by the formation of 15 disulfide bridges and greatly increases its sensitivity toward protcolysis. The process is highly cooperative, occurring in an all-or-none fashion (Fominaya and Hofsteengc, 1992). Whether such reactions also play a role in vivo is unknown, but it has been observed that creating oxidative stress in cells in culture decreases RI activity and antigen (Bl4zquez and Hofstecnge, 1996). 2. T h r e e - D i m e n s i o n a l S t r u c t u r e
The repetitive primary structure of RI suggested that also the threedimensional structure would be highly regular. The structure, determined by Kobe and Deisenhofer (1993), fulfilled all expectations and revealed, for the first time, the basic structural features of the LRR (Fig. 2). Each of the 16 LRRs forms a/3o~ unit, which assembles into a right-handed supercoil with a horseshoe-like shape of overall dimensions 70 x 62 x 32 A. Such a supercoil is not unique to an LRR-containing protein; it also occurs in pectate lyase and alkaline protease, where exclusively/3 strands are involved in the repeating unit (Yoder and Jurnak, 1995; Kobe and Deisenhofer, 1994). The conserved residues in the LRR of RI fulfill structural roles. The leucines at positions 2, 5, 7, 12, 20, and 24 and the residue at position 17 (Fig. 1) form a hydrophobic core between the /3 sheet and the c~ helix, and between the/3c~ units. The LRRs in most proteins contain an asparagine at position 10 of the repeat,
19
639
Ribonuclease Inhibitor
N
N
Fig. 2. Three-dimensional structure of porcine RI. Stereo view of the three-dimensional structure of porcine RI (Kobe and Deisenhofer, 1993). For the sake of clarity, only the C,, trace is shown. The positions of the N and C termini have been indicated. The coordinates were taken from the Brookhaven National Laboratory Protein Data Base (lbnh.pdb).
as in the B repeat of RI (Kobe and Deisenhofer, 1994). The side chain of this residue, as well as that of the corresponding cysteine in the A repeat, is buried and forms a hydrogen bond to the polypeptide backbone. Interestingly, a repeated asparagine with the same function also occurs in pectate lyase (Yoder et al., 1993). Conserved residues at the surface of RI are involved in three hydrogen bonding networks. These involve the alternating positively and negatively charged residues at position 3 of the repeats, the glutamic acid at position 4 of the B repeat with the hydroxyamino acid in the A repeat, and the threonine and aspartate at positions 13 and 14 of the A and B repeat, respectively. Nearly all of these structurally important residues have been conserved in porcine, human, and rat RI. If exceptions occur they tend to be located in the less homologous repeats, N or C (Fig. 1). The sensitivity of RI toward thiol modification is not simply the result of the large number of thiol groups. Inspection of the three-dimensional model reveals that only Cys residues in positions 7, 57, 371, and 404 are located at the surface. This agrees with the observation that these cysteines, except Cys-7, are the most easily alkylated ones (Hofsteenge et al., 1991a). The cysteines in the CXLTXXXC motif of the A repeat and at repeat position 29 are completely buried. Those at repeat position 21 are on the outside of the horseshoe, more or less in the surface, but most often with the sulfhydryl group tucked between the helices. The
640
Jan Hofsteenge
highly cooperative oxidation of the 30 thiol groups to 15 disulfides (Fominaya and Hofsteenge, 1992) raises the possibility that one or m o r e of the cysteines function as a kind of trigger. G u i d e d by the three-dimensional structure, further biochemical studies are necessary to resolve this question and to establish the structural features underlying the cooperativity.
3. Three-Dimensional Structure of Complex with RNase A
Description of the Complex. RI inhibits R N a s e by blocking access of the substrate to the active site (Kobe and Deisenhofer, 1995). The enzyme docks with the active site onto the C-terminal end of RI (Fig. 3), w h e r e b y the lobe containing the 65-72 loop and the region a r o u n d H i s - l l 9 interact with the surface of the horseshoe. The lobe containing
Fig. 3. Three-dimensional structure of the RI-RNase complex. Schematic structure of the RI-RNase A complex. The backbones of RI (dark grey) and RNase A (light grey) are shown as ribbons. The RI molecule has been rotated approximately 45~ around the axis, compared to the orientation in Fig. 2. Adapted from Kobe and Deisenhofer (1995).
19
Ribonuclease Inhibitor
641
the Lys-41 and 86-96 loops tuck into the central cavity. A total of 28 residues from RI and 24 residues from the enzyme provide 56 intermolecular contacts. On complex formation 2551 .~2 of surface area becomes buried, but no significant structural changes take place in the enzyme. Throughout the RI molecule, however, small changes occur that cumulatively widen the central cavity (Kobe and Deisenhofer, 1995). The residues on RI that are involved in the binding of RNase A cluster in three separate groups ( A - C in Fig. 4; see also Fig. 1). The complementary residues in RNase A (Fig. 5) lie mainly in the active site groove and in the loops around it. In addition, contact residues on the backside of the molecule are located in the c~2 helix and in the 86-96 loop. Group A in RI is located in the N-terminal repeats N, A1, A2, and B2 and provides 8 contacts with RNase residues around Lys-31 in the c~2 helix. Group B, situated in repeats A4, B4, A5, B5, and A6 (Figs. 1 and 4), makes 12 contacts with residues in the RNase 86-96 loop. The largest number of contacts is provided by group C, whose residues densely cluster in repeats B7 and C and in the C-terminal/3 strand (Fig. 4). This region of RI makes 36 contacts to the active site of RNase A and the loops surrounding it. These contacts involve the catalytic residues Lys-41 and His-119, as well as most of the residues that bind the substrate
Fig. 4. Residues in RI that bind to RNase A. Stereo view of the structure of the Ca backbone of RI, with the side chains of contact residues shown as ball-and-stick models. The three clusters (A-C) have been indicated. The coordinates were taken from Brookhaven National Laboratory Protein Data Base (lbnh.pdb).
642
Jan Hofsteenge N
K31
C
N
K31
C
Fig. 5. RNase A residues that bind to RI. Stereo view of the structure of the C,, backbone of RNase A, with the side chains of the contact residues shown as ball-andstick models. Residues representative of the different regions have been indicated.
(Fig. 5), including Val-43 (B~ site), Lys-66 (P0 site), Gln-ll (p~ site), Gin-69, Asn-71, Glu-111 (B2 site), and Lys-7 (P2 site) (see Chapter 9 for the nomenclature of the RNase substrate-binding site). Although the amino acid sequences of RI from pig, human, and rat cells differ at approximately 110 positions throughout the polypeptide chain, 23 of the 28 contact residues are identical, and three have been replaced conservatively in human and/or rat RI (Fig. 1). This explains the essentially identical affinities of human and porcine RI for RNase A (Table II). It is, however, in strong contrast to the puzzlingly large degree of variability observed in the RI-binding domain of the enzyme (Section III,B,3,c). b. B i o c h e m i c a l D a t a . Comparison of structural features of the R I RNase A complex with the results obtained from biochemical experiments provokes some interesting questions (see also Section III,B,3,c). A number of deletion mutants and a proteolytically shortened form of RI have been kinetically characterized. Removal of residues 1-90 (or 93) of porcine RI deletes 6 of the 56 contacts to RNase A. Nevertheless, the binding energy reduces by only 3% (Hofsteenge et al., 1991b), suggesting that the interaction of the c~2 helix of RNase A does not contribute greatly in energetic terms. In agreement with the observation that the majority of RI-RNase A contacts take place at the C terminus of the inhibitor, Lee and Vallee (1990) found that deletion mutants missing
19
Ribonuclease Inhibitor
643
residues 144-257 or 315-371 of human RI (residues 140-253 or 311-367 of porcine RI) yielded inhibitors with K~ values in the picomolar range. Each of these deletions removes only 3 of the 56 contacts, but large decreases in binding energy were observed, 19.4 and 16 kJ/mol for A(144257) and A(315-371), respectively. This suggests that either the deleted contact residues contribute extraordinarily to the binding energy (see also Section III,B,3,c) or that the deletions alter the conformation of the RNase binding site(s). Contact residues on RNase A and RI have been identified by differential labeling experiments. Cys-371 and Cys-404 in RI are protected from alkylation in the complex (Hofsteenge et al., 1991a). For Cys-404 this can be explained by its direct contact with RNase A, but the reduced modification of Cys-371 must result from its location between clusters A and B. In RNase A, Lys-7, Lys-31, Lys-41, and Lys-91 are fully protected by RI against amidination (Blackburn and Gavilanes, 1982). Lys66, which is in contact with Cys-404 in RI (Kobe and Deisenhofer, 1995), is not protected. Chemical modification of Lys-41 and Lys-7 (Blackburn and Gavilanes, 1980; Neumann and Hofsteenge, 1994) also showed that these residues are important for binding. An attempt to identify contact regions by using peptides as competitors for the interaction (CrcvclThieffry et al., 1992) has been unsuccessful (Fominaya et al., 1993). c. I n t e r a c t i o n w i t h O t h e r R N a s e s . One of the most rcmarkablc propcrtics of RI is its high-affinity binding to RNases that diffcr widely in amino acid sequence. In fact, of the 24 contact residues of RNase A, only 6 have been conserved in all RNases for which the binding to RI has been quantitated, yet the energies of complex formation, AGb, do not differ by more than 13% (Table Ill). A possible explanation would be that these six residues are part of a "hot spot of binding energy," i.e., a small number of contact residues that provide the major proportion of binding energy, in analogy to the binding of human growth hormone to its receptor (Clackson and Wells, 1995). Alternatively, RI interacts with each of these RNases in a different way. This would, however, require the existence of still unidentified interaction areas on the RNases and/or on RI (Shapiro et al., 1995). The results obtained by mutagenesis of angiogenin provide some evidence for the second hypothesis. For instance, in angiogenin the 65-72 loop (RNase A nomenclature) contains a number of deletions and has a different spatial arrangement (Acharya et al., 1994). Surprisingly, replacing residues 58-70 by residues 59-73 of RNase A minimally affects
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Jan Hofsteenge
T A B L E III Conservation of Residues in RI-Binding Domain of RNase a AGb (kJ/mol)b:
-75.5
-86.5
-85.8
- 82.2
RNase A
Angiogenin
RNase 2
RNase 4
K7 Qll N24 Q28 K31 $32 L35 D38 R39 K41 P42 V43 K66 N67 Q69 N71 E86 G88 $89 $90 K91 A 109 E111 Hl19
H8 Q12 R24 $28 R31 R32 L35 -P38 K40 D41 I42 ~ N68 L83 G85 G86 $87 P88 A 106 El08 Hl14
W10 Q14 Q21 N25 Q28 V29 N32 R35 R36 K38 N39 Q40 P63 $64 K66 N70 L85 T87 P88 $89 P90 A 110 D112 H129
c Onconase
R7 Qll
T5 K7
L27 Q30 R31 M34 H37 Y38 K40 R41 F42 K65 N66 Q68 N70 E83 G85 $86 $87 R88 A 106 El08 Hl16
N21 $24 F28 H29 K31 D32 K33
K55 V70 $72 R73 P74 T89 E91 H97
Residues that have been conserved in all four inhibited RNases have been underlined. b Calculated from Table II, using AGb = R T In Ki, where R is the gas constant and T is the absolute temperature. " No inhibition was observed at 0.5/zM of enzyme and inhibitor.
RI binding (Harper and Vallee, 1989). Apparently, the mutated loop, which in RNase A contributes 11 contacts to RI, does not participate in angiogenin. Furthermore, replacement of residues that have a counterpart in RNase A that contact RI (Table III) does not always alter the affinity of angiogenin for the inhibitor. The mutations K40A and R32A
19
Ribonuclease Inhibitor
645
increase Ki 1300- and 10-fold, respectively (Lee and Vallee, 1989a; Shapiro and Vallee, 1992), but the substitutions R31A (Shapiro and Vallee, 1992) and H114A (Shapiro and Vallee, 1989) hardly affect the inhibition constant. On the other hand, the mutation of a residue not expected to bind RI, Arg-5, resulted in a 50-fold decrease in affinity (Shapiro and Vallee, 1992). Much less information is available for RNases 2 and 4. RNase 2 differs from RNase A in 17 of the 24 contact residues, yet it binds approximately as tightly to RI as angiogenin, with whom it also differs in 17 of the putative contact residues (Table III). This implies the existence of additional interactions between RI and RNase 2 that are different from those between RI and angiogenin. Also, RNase 4 binds RI more tightly than RNase A. It differs in particular in the residues of the Lys-41 loop. Either these residues interact more favorably with RI, or, also in this case, additional interactions with RI exist. Replacement of the N-terminal amino acid sequence of RNase 4 with residues 1-10 of RNase A increased the affinity 13-fold (U. Neumann, A. Vicentini, and J. Hofsteenge, 1994, unpublished results), a phenomenon also observed with angiogenin (Bond and Vallee, 1990). Most likely this results from a specific interaction of Lys-7, which has been shown to contribute 10 kJ/mol to the binding energy (Neumann and Hofsteenge, 1994). These results strongly indicate that the four RNases interact with RI in slightly different ways, thus supporting the second hypothesis. However, at present neither of the two hypotheses alone is completely satisfactory, and further studies, using kinetic analysis of site-directed mutants and X-ray crystallography, will be required to resolve this intriguing problem. Onconase, which is not at all inhibited by human RI (whether it binds RI is unknown) (Wu et al., 1993), differs from RNase A in 21 of the 24 contact residues (Table III). This is similar to the number of differences found for RNase 2 (17) and angiogenin (16). The threedimensional structure of onconase has been determined (Mosimann et al., 1994). All three loops in RNase A that interact with RI, but in particular the 65-72 loop, have a different structure (Fig. 6). The RI found in frog cells is of higher molecular weight and inhibits frog RNases (Nagano et al., 1976). It will be of interest to determine the structure of this RI, to reveal how it deals with an RNase having such different binding regions.
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Jan Hofsteenge
,,•
Lys-.41
6-96
Lys-.41
Fig. 6. Comparison of the structures of RNase A and onconase. Stereo view of the superimposed C,, backbones of RNasc A (thick line; 7rsa.pdb) and onconasc (thin line; lonc.pdb). The loops in RNasc A containing contact residues have bccn indicated. The coordinates wcrc taken from the Brookhavcn National Laboratory Protein Data Base.
IV. APPLICATIONS A. Protection of R N A
Even before highly purified RI became available, its protective effect against RNA degradation was employed. In fact, RI was "rediscovered" as a polyribosomc-stabilizing factor (Bontet al., 1965), and addition of rat liver supcrnatant to preparations of polyribosomcs improved recoveries (Lawford et al., 1966; Blobel and Potter, 1966). The use of crude RI in other assays was hampered, however, by contaminating protease activity (Berns et al., 1971). The availability of purified RI allowed its inclusion in in vitro mRNA translation systems, in particular wheat germ extracts (Scheele and Blackburn, 1979; Robbi and Lazarow, 1978), resulting in more and longer polypeptides. After the original publication by de Martynoff et al. (1980), showing that the presence of RI greatly improved the yield of full-length thyroglobulin eDNA, inclusion of RI in cDNA synthesis has become common practice. Similarly, longer transcripts are obtained in cell-free RNA synthesis by including RI (Eichler et al., 1981). In this context it is of interest to examine the specificity of mammalian RI. Of the mammalian RNases, only those belonging to the superfamily of pancreatic-type RNases (Chapter 8) are inhibited. Two noticeable exceptions are the RNase dimer from bovine seminal vesicles, which is
96
19 Ribonuclease Inhibitor
647
only inhibited when dissociated into monomers (Blackburn and Gavilanes, 1980; Murthy and Sirdeshmukh, 1992), and onconase from the frog Rana pipiens (Wu et al., 1993). RNases from crude extracts from chicken and frog liver are not inhibited by mammalian RI (Roth, 1962; Kraft and Shortman, 1970a), indicating that cross-reaction between species breaks down somewhere between mammals and birds (see also Section II,A). It is therefore surprising that a purified endoribonuclease from the insect Ceratatis capitata is inhibited by human RI (GarcfaSegura et al., 1985). The effect of RI on plant RNases varies. Crude barley RNases and purified RNase LV3 from tomato, an enzyme belonging to the RNase T2 family (Chapters 3 and 5), are not inhibited by human RI (KOck et al., 1995). In contrast, an RNase from C u c u m i s sativus seeds, probably belonging to the same family (Rojo et al., 1994), and an unrelated RNase from timothy grass pollen (Bufe et al., 1995) have been reported to be inhibited. No bacterial or fungal RNases that are inhibited by RI have yet been found (Shortman, 1962b; Cho and Joshi, 1989).
B. RNase Assay
The ability of RI to bind a variety of pancreatic-type RNases suggested the interesting possibility to assay RNases with very low ribonucleolytic activity, e.g., angiogenin (Bond, 1988; Bond and Vallee, 1988). For this purpose, a known amount of RI is added to a solution containing an unknown quantity of enzyme. RI remaining unbound can then be determined by "backtitration" with an RNase having a high specific activity, e.g., RNase A.
C. Inhibition of Angiogenesis
A potentially very fruitful application of RI is based on the observation that it inhibits the ribonucleolytic and angiogenic activity of angiogenin (Shapiro and Vallee, 1987). Evidence has been presented strongly suggesting that the growth of solid tumors is dependent on increased neovascularization (Folkman, 1990, and references therein). Consequently, angiogenesis inhibitors are potential antitumor agents. Low doses of human
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RI (100 ng/mouse), administered intratumorally or intraperitoneally, inhibit the growth of Meth A sarcoma, MM46 mammary adenocarcinoma, and B16 melanoma, and increase the lifespan of the recipient animal (Fukushima et al., 1990). Furthermore, RI, together with tumor amputation, has been claimed to reduce metastasis in a Lewis lung carcinoma system. These results are in contrast with those of Norioka et al. (1992), who did observe inhibition of tumor-induced angiogenesis, but no decrease in tumor size from B16 melanoma cells. The detailed study by Polakowski et al. (1992, 1993) used penetration of polyvinyl sponge disks by endothelial cells in mice, demonstrating strong inhibition of angiogenesis by RI. Unexpectedly, the increased penetration induced by basic fibroblast growth factor or sodium orthovanadate is also inhibited. Furthermore, the growth of C755 mammary tumor cells is at least partially inhibited by human RI. All studies, except that by Fukushima et al. (1990), show that the route of administration of RI is important. Systemic application seems to be precluded due to instability of RI activity, and slow release from implanted polymers has been the most successful method so far. The mechanism by which RI reduces tumor growth remains to be established. Importantly, in the cases where RI affects the growth of tumors in vivo, it does not affect the proliferation of the tumor cells (or normal endothelial cells, in culture). This would be consistent with inhibition of angiogenesis being involved in the suppression of tumor growth. Whether interaction of RI with other proteins is required as well (Norioka et al., 1992; Polakowski et al., 1993) is unclear. It has been suggested that the inhibition of metastasis by RI is due to its interaction with extracellular matrix components (Shinimiya et al., 1993). Finally, the use of RI as an inhibitor in other diseases that depend on increased angiogenesis, e.g., rheumatoid arthritis, diabetic retinopathy, and psoriasis, remains to be investigated. The engineering of a more stable RI, e.g., by replacing sensitive cysteine residues, seems to be a prerequisite for such applications. Knowledge of the structural and functional properties of RI should facilitate this.
ACKNOWLEDGMENTS
I would like to thank Dr. Andreas LOftier for reading the manuscript.
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Index
Entry followed by f denotes figure.
Actin binding to angiogenin, 473. 480 to BS-ribonuclcasc, 414 Adhesion. ccll. stimulation by angiogcnin. 477 Amphibia. see spec([ic types Amphibian ribonuclcasc inhibitor. 479, 499 Amphibian ribonuclcascs, 573 antitumor activity, 499 Angiogcncsis, 446-450 assays, 449 inhibition of by RI, 462, 647 Angiogcnic factors. 448, 475 Angiogcnin, see also spec
