subcellular biochemistry: volume 7

Subcellular Biochemistry Volume 7

ADVISORY EDITORIAL BOARD

J. ANDRE Laboratoire de Biologie Cellulaire, 4 Faculte des Sciences, 91 Orsay, France

D. L. ARNON Department of Cell Physiology, Hilgard Hall, University of California, Berkeley, California 94720, USA

J. BRACHET Laboratoire de Morphologie Animale, Faculte des Sciences, Universite Libre de Bruxelles, Belgium

J. CHAUVEAU Institut de Recherches Scientifiques sur Ie Cancer, 16 Avenue VaillantCouturier, 94 Ville Juif, Boite Postale 8, France

C. de DUVE Universite de Louvain, Louvain, Belgium and The Rockefeller University, New York, NY 10021, USA

M. KLINGENBERG Institut fUr Physiologische Chemie und Physikalische Biochemie, Universitat Miinchen, Goethestrasse 33, Miinchen 15, Germany

A. LIMA-de-FARIA Institute of Molecular Cytogenetics, Tornavagen 13, University of Lund, Lund, Sweden

O. LINDBERG The Wenner-Gren Institute, Norrtullsgatan 16, Stockholm, V A, Sweden

V. N. LUZIKOV A. N. Belozersky Laboratory for Molecular Biology and Bioorganic Chemistry, Lomonosov State University, Building A, Moscow 117234, USSR •

H. R. MAHLER Chemical Laboratories, Indiana University, Bloomington, Inaiana 47401, USA

M. M. K. NASS Department of Therapeutic Research, University of Pennsylvania School of Medicine, Biology Service Building, 3800 Hamilton Walk, Philadelphia, Pennsylvania 19104, USA

A. B. NOVIKOFF Department of Pathology, Albert Einstein College of Medicine, Yeshiva University, Eastchester Road and Morris Park Avenue, Bronx, NY 10461, USA

R. N. ROBERTSON Macleay Building, A12, School of Biological Sciences, The University of Sydney, Sydney, N.S.W. 2006, Australia

P. SIEKEVITZ The Rockefeller University, New York, NY 10021, USA

F. S. SJOSTRAND Department of Zoology, University of California, Los Angeles, California 90024, USA

A. S. SPIRIN A. N. Bakh Institute of Biochemistry, Academy of Sciences of the USSR, Leninsky Prospekt 33, Moscow V-7l, USSR

D. von WETTSTEIN Department of Physiology, Carlsberg Laboratory, Gl. Carlsbergvej 10, DK-2500, Copenhagen, Denmark

V. P. WHITTAKER Abteilung fUr Neurochemie, Max-Planck Institut fUr Biophysikalische Chemie, D-3400 Gottingen-Nikolausberg, Postfach 968, Germany

A Continuation Order Plan is available for this series. A continuation order will bring delivery of each new volume immediately upon publication. Volumes are billed only upon actual shipment. For further information please contact the publisher.

Subcellular Biochemistry Volume 7 Edited by

Donald B. Roodyn University College London London, England

PLENUM PRESS • NEW YORK AND LONDON

The Library of Congress cataloged the fust volume of this title as follows:

Sub-cellular biochemistry.

London, New York, Plenum Press. v. iUus. 23 cm. quarterly.

Began with Sept. 1971 issue. Cf. New serial titles. 1. Cytochemistry - Periodicals. 2. Cell organelles - Periodicals.

QH611.S84 574.8'76 73-643479

Library of Congress Catalog Card Number 73-643479

ISBN 978-1-4615-7950-2 ISBN 978-1-4615-7948-9 (eBook) 001 10.1007/978-1-4615-7948-9

This series is a continuation of the journal Sub-Cellular Biochemistry, Volumes 1 to 4 of which were pu bUshed quarterly from 1972 to 1975

© 1980 Plenum Press, New York Softcover reprint of the hardcover 1st edition 1980 A Division of Plenum Publishing Corporation 227 West 17th Street, New York, N.Y. 10011

All rights reserved

No part of this book may be reproduced, stored in a retrieval system, or transmitted, in any form or by any means, electronic, mechanical, photocopying, microfilming, recording, or otherwise, without written permission from the Publisher

Contributors

A. A. Bogdanov A. N. Belozersky Laboratory of Molecular Biology and Bioorganic Chemistry, Moscow State University, Moscow 117234, U.S.S.R.

W. Dierick RUCA-Laboratory for Human Biochemistry and UIA-Laboratory for Pathological Biochemistry, University of Antwerp, Groenenborgerlaan 171, B2020 Antwerp, Belgium

A. A. Hadjiolov Department of Molecular Genetics, Institute of Molecular Biology, Bulgarian Academy of Sciences, Sofia, Bulgaria

H. J. Hilderson RUCA-Laboratory for Human Biochemistry and UIA-Laboratory for Pathological Biochemistry, University of Antwerp, Groenenborgerlaan 171, B2020 Antwerp, Belgium

A. M. Kopylov A. N. Belozersky Laboratory of Molecular Biology and Bioorganic Chemistry, Moscow State University, Moscow 117234, U.S.S.R.

A. Lagrou RUCA-Laboratory for Human Biochemistry and UIA-Laboratory for Pathological Biochemistry, University of Antwerp, Groenenborgerlaan 171, B2020 Antwerp, Belgium

J. Michael Lord School of Biological Sciences, University of Bradford, Bradford BD7 IDP, W. Yorkshire, U.K.

Ian F. Pryme Cell Biology Research Group, Department of Biochemistry, The Preclinical Institute, University of Bergen, Arstadveien 19,5000 Bergen, Norway

C. I. Ragan Department of Biochemistry, University of Southampton, Southampton, HANTS, S09 3TU, U.K.

Milton R. J. Salton Department of Microbiology, New York University School of Medicine, New York, N.Y. 10016, U.S.A.

I. N. Shatsky A. N. Belozersky Laboratory of Molecular Biology and Bioorganic Chemistry, Moscow State University, Moscow 117234, U.S.S.R.

Asbjorn M. Svardal Cell Biology Research Group, Department of Biochemistry, The Preclinical Institute, University of Bergen, Arstadveien 19, 5000 Bergen, Norway

G. Van Dessel RUCA-Laboratory for Human Biochemistry and UIA Laboratory for Pathological Biochemistry, University of Antwerp, Groenenborgerlaan 171, B2020 Antwerp, Belgium

Aims and Scope

SUBCELLULAR BIOCHEMISTRY aims to bring together work on a wide range of topics in subcellular biology in the hope of stimulating progress towards an integrated view of the cell. In addition to dealing with conventional biochemical studies on isolated organelles, articles published so far and planned for the future consider such matters as the genetics, evolution, and biogenesis of cell structures, bioenergetics, membrane structure and functions, and interactions between cell compartments, particularly between mitochondria and cytoplasm and between nucleus and cytoplasm.

Articles for submission should be sent to Dr. D. B. Roodyn, Department of Biochemistry, University College London, Gower Street, London WCI E 6BT, U.K. There are no rigid constraints as to the size of the articles and in general they should be between 9,000 and 36,000 words, with an optimum size of about 20,000 words. Although articles may deal with highly specialized topics, authors should try as far as possible to avoid specialist jargon and to make the article as comprehensible as possible to the widest range of biochemists and cell biologists. Full details of the preparation of manuscripts are given in a comprehensive Guide for Contributors which is available from the Editor or Publishers on request.

Preface

The broad aim of SUBCELLULAR BIOCHEMISTRY is to present an integrated view of the cell in which artificial barriers between disciplines are broken down. The contents of Volume 7 illustrate the interconnections between initially unrelated fields of study and show strikingly how advances along one front become possible because of parallel successes in another. Current research into cell organelles and membrane systems is not only concerned with the elucidation of their structure and function. It also asks such questions as: Which regions of the cell are concerned in the bioassembly of the organelle? How are organelle and membrane precursors transported from the site of synthesis to the newly formed cell constituent? What genetic systems control the biosynthesis and assembly of cell components and how do these systems interact? How did the various cell constituents evolve? How did the genetic and biosynthetic systems making the organelles themselves evolve? The search for the answer to such questions has placed organelle biochemistry on a different level than that of the more restricted studies of the 1950s and early 1960s and promises to produce some fascinating and surprising results.

Volume 7 opens with a detailed chapter by A. A. Hadjiolov on the biogenesis of ribosomes of eukaryotes. The general arrangement of ribosomal genes is discussed, and there is a full account of their transcription. The chapter describes the way in which the primary "pre-rRNA" is processed to produce the final rRNA, as well as how the pre ribosome is processed. There is also an important section on the regulation of ribosome biogenesis. In spite of the many obvious deficiencies in our knowledge, one cannot help but be struck, on reading Hadjiolov's account, by the great advances and wealth of detail we now have on this important process. The most striking achievement is the elucidation of the main features of the organization of rRNA genes, with multiple transcription units that include transcribed spacer sequences, separated by nontranscribed spacer sequences. Our understanding of ribosomal genetics has thus now reached the molecular level, with sequence mapping. When one combines this with the great advances made in our understanding of the structure of ribosomal RNA (see below), it is clear that we are now rapidly reaching the

ix

x Preface

point at which it will be possible to describe the processes involved in ribosome bioassembly in defined molecular terms. Considering the structural complexity of the ribosome this is a formidable achievement.

Chapter 2, by A. A. Bogdanov, A. M. Kapylov, and I. N. Shatsky, deals with the role of RNA in the organization and function of the most commonly studied prokaryotic ribosome, that from Escherichia coli. The question under analysis is: To what extent do the primary structural features of ribosomal RNA determine the final molecular organization of the ribosome? The general conclusion (at least as regards the effect of 16 S RNA on the 30 S subunit) is, to quote the authors: "the major morphological peculiarities of the 30 S subunits are inherent in the very structure of 16 S RNA." From the point of view of establishing molecular mechanisms for ribosome assembly and evolution, this is a most important conclusion, because it places most of the ribosomal proteins in a rather secondary role. The chapter by Bogdanov and his colleagues discusses in a most intriguing manner the complex molecular interactions that can occur between ribosomal RNA and the various ribosomal proteins; this discussion is presented against the background of the striking recent advances in the determination of the nucleotide sequences of E. coli ribosomal RNAs. The interactions of ribosomal proteins with rRNA can now be examined in precise molecular terms and the role of "domains" of RNA-protein interactions accurately analyzed. As with Hadjiolov's chapter, one has a sense of excitement as previously vague concepts are replaced by precise studies based on known structural properties of ribosomal proteins and nucleic acids, and also on extensive nucleotide sequence information. The two chapters powerfully illustrate the close interaction that is now taking place between genetic, structural, and functional studies of the ribosome.

Chapter 3, by A. M. Svardal and I. F. Pryme, surveys recent advances in our understanding of the role of the endoplasmic reticulum in protein synthesis. Again, one can detect the process whereby previously vague concepts have been replaced by more rigorous studies based on the interaction of defined molecular species. The most striking new approach is the now famous "signal" hypothesis of Blobel and Sabatini (1971), which examines the problem of the translocation of proteins from their site of synthesis to their final intracellular location, not by means of generalities but by examining defined amino acid sequences. Our whole understanding of the relationship between ribosomes and the membrane has undergone a great change in recent years; Svardal and Pryme's chapter shows clearly how the problems of the biosynthesis of specific proteins (whether they be immunoglobulin light chains or mitochondrial membranebound enzymes) are currently being studied in rigorous molecular terms. The authors also dwell on the interesting question of "compartmentalization" of the rough endoplasmic reticulum. Are specific proteins made in specific "regions"? Apparently there is good evidence that light-chain immunoglobulin synthesis

Preface xi

occurs on membrane-bound polysomes and that this process is indeed compartmentalized within a distinct region (or subfraction) of the rough endoplasmic reticulum. We are thus moving toward the view of the rough endoplasmic reticulum as being a complex, highly organized system with local regions performing specialized functions. The elucidation of the architecture of such a system will be a formidable task.

Chapter 4, by J. M. Lord, deals with the biogenesis of peroxisomes and glyoxysomes. It leads on naturally from the previous account of the endoplasmic reticulum and includes much discussion of the role of membranebound poly somes and of the "signal" hypothesis in relationship to the biosynthesis and transport of peroxisomal and glyoxysomal proteins. Thus, as a further striking illustration of the integrative tendency I have been discussing, it has become apparent that the study of the biogenesis of peroxisomes and related particles is really just a "special case" of the more general study of the mechanism of protein synthesis by the endoplasmic reticulum. The authors survey the advances that have been made in our understanding of the structure, function and biogenesis of peroxisomes and glyoxysomes and it is intriguing to see how our appreciation of the metabolic importance of these organelles has steadily increased over the years. An important example is the realization that much (but not all) of the fatty acid ,B-oxidation activity previously observed in rat liver mitochondrial fractions was, in fact, attributable to contamination by peroxisomes, which have their own ,B-oxidation system. If this process continues there will have to be much rewriting of biochemistry textbooks! In general, the chapter by Lord clearly illustrates how studies on the structure, function, and biogenesis of these organelles are advancing hand in hand in an integrated fashion.

Chapter 5, by H. J. Hilderson, G. van Dessel, A. Lagrou, and W. Dierick deals with the subcellular biochemistry of one specific tissue, the thyroid. One of the great hazards of cell biochemistry is to believe that what is true for rat liver is true for all mammalian cells. In fact, each cell type has its own specific subcellular enzyme distribution pattern, peculiarly suited to the needs of the cell. The chapter by Hilderson and his colleagues illustrates the great merit of subjecting a given tissue to an in-depth analysis. One of the problems of not working with rat liver is to be sure of one's markers, because an enzyme typical of a rat liver organelle may either be absent from another cell type or even distributed differently. A most useful feature of the chapter is the inclusion of detailed tables that list the subcellular localization of a large number of thyroid constituents and also indicates the usefulness and validity of the various marker enzymes. Because of its detailed analysis of the experimental problems involved in the fractionation of thyroid homogenates and its range of coverage of the literature, "The Subcellular Biochemistry of the Thyroid" could well become established as the definitive review on this important topic.

xii Preface

Chapter 6, by C. l. Ragan, deals with the molecular organization of NADH dehydrogenase. After a useful introductory discussion of the confused terminology of this important enzyme system, the author surveys the various methods used to fragment the multi enzyme system and gives the results of analysis of its polypeptide composition. He then discusses the various approaches that have been made in recent years to establish the physical relationship between the various subunits. The role of phospholipids is also examined as is the organization of NADH in the mitochondrial enzyme. An interesting and important new model is proposed in which central flavoprotein and catalytic fragments are surrounded by a "shell" of protein subunits. The originality of the suggestion is that most of the subunits in the shell are not directly involved in the enzyme-catalyzed reaction, but are there to provide a correct environment in which the catalytic elements can function. Ragan's approach to the problem of the organization of NADH dehydrogenase may well be of relevance to other mitochondrial enzyme complexes (in particular, cytochrome oxidase) and could be of considerable help in the formulation of reasonable mechanisms for the bioassembly of these complexes. This is therefore yet another example of how the various approaches to organelle and membrane biochemistry now interact.

Chapter 7, by M. R. J. Salton, surveys our current understanding of the structure and biochemical organization of the membrane of Micrococcus lysodeikticus, an organism that is particularly sensitive to lysozyme and hence particularly amenable to subcellular analysis. The chapter indicates the advances that have been made in our understanding of what is clearly a multifunctional membrane system and also points out the difficulties that still remain in the purification and analysis of membrane-bound bacterial enzymes. One striking feature of the chapter is the wealth of information that is now available from the use of sophisticated immunological techniques; another remarkable result of current research is that the F1-ATPase of M. lysodeikticus shows striking resemblances to other F1-ATPases from bacteria, chloroplasts, and mitochondria. Indeed, one cannot but regard the F1-ATPase as a fundamental enzyme assembly, probably present in all energy-transducing systems. Nothing could illustrate more strikingly the underlying unity of living things or confirm the ever-accelerating process of unification that is now taking place in biological research.

As in previous volumes, we include an extensive book review section. Again, one cannot help but be impressed by the range and general excellence of books now being published in cell biology. In many ways th.e student texts describe the advances being made more clearly than in the heavier specialist multiauthor texts. Such is the pace of modern developments, one suspects that the better student texts will be used more and more often by working scientists, simply to orient themselves in the flood of new findings. The authors of

Preface xiii

research-oriented reviews are often reluctant to make generalizations and try, quite justifiably, to present all the arguments for and against a given view. There is also an inevitable and irreducible amount of jargon in any field, and as a result a specialist review is often of little help to an outsider coming fresh to the subject. In contrast, the author of an educational text has as his primary aim communication with the nonexpert. His task is to explain concepts and delineate major advances and he is not under the obligation to justify every statement by reference to the primary literature. Thus, unless we have more reviews written by specialists not for their closest friends, but for the scientific community in general, it will become increasingly frequent for research workers to turn to books that were originally aimed at students. Since it is impossible for an author of a student text to be an expert in every field he surveys, the danger exists that incorrect theories or "dogmas" may become "canonized" or "institutionalized" by inclusion in standard texts.

Since its inception, SUBCELLULAR BIOCHEMISTRY has been concerned with the problems of communication between scientists, and we are most grateful to the bulk of our contributors, who have struggled valiantly to describe complex phenomena in their specialties in relatively simple language. It is easy to obfuscate and confuse-one wonders whether the scientific community appreciates how difficult it is to do the opposite and whether sufficient attention is paid to the problem of how we can learn to talk to each other, rather than at each other.

D. B. Roodyn London

Contents

Chapter J Biogenesis of Ribosomes in Eukaryotes

A. A. Hadjiolov

1. Introduction ................ . 2. Ribosomal Genes .... . ..... .

2.1. Ribosomal RNA Genes 2.2. 5 S rRNA Genes .......... . 2.3. General Features

3. Transcription of Ribosomal RNA Genes ............ . 3.1. Components of the Transcription Complex ..... . 3.2. The Transcription Process ...... . 3.3. Transcription in Vitro .. . ......... .

4. Processing of Primary Pre-rRNA and Preribosomes . . . ...... . 4.1. Structure of Primary Pre-rRNA ....................... . 4.2. Pre-rRNA Maturation Pathways ......... . 4.3. Preribosomes: Structure and Processing .. .

5. Regulation... . ....................... . 5.1. General Considerations .... ......... . ..... . 5.2. Transcriptional Control ............ . 5.3. Posttranscriptional Control

6. Concluding Remarks ... 7. References............ . ..... .

Chapter 2 The Role of Ribonucleic Acids in the Organization and Functioning of Ribosomes of E. coli

A. A. Bogdanov, A. M. Kopylov, and I. N. Shatsky

2 2

12 15 16 16 18 29 31 32 34 42 44 44 46 51 58 60

1. Introduction.... . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 81 2. The Secondary Structures of Ribosomal RNA in Ribosomes 83

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xvi Contents

3. Compact Folding of RNA in Ribosomal Subunits 85 4. Domain Organization of Ribosomal Subunits 86 5. Role of Ribosomal Proteins in the Organization of RNA Tertiary

Structure within Ribosomal Subunits. . . . 91 6. Direct Participation of Ribosomal RNA in Ribosome Functioning 94 7. Topography of rRNA in Ribosomes. 99 8. Addendum: A Preliminary Model for the Secondary Structure of

16 S Ribosomal RNA 105 9. References. 108

Chapter 3 Aspects of the Role of the Endoplasmic Reticulum in Protein Synthesis

Asbjorn M. Svardal and Ian F. Pryme

I. Introduction .. 2. Membranes of the Endoplasmic Reticulum.

2.1. The Composition and Structure of the Endoplasmic Reticulum Membranes ...... .

2.2. Functional Aspects of the Endoplasmic Reticulum. 3. Protein Synthesis.

3.1. Protein Synthesis by Free and Membrane-Bound Polysomes . 3.2. Secreted Proteins May Be Synthesized Solely on Membrane-

Bound Polyribosomes . .. ......... . ......... . 3.3. Proteins Synthesized on Membrane-Bound and/or Free

Polysomes 3.4. Summary ..

4. Polyribosome-Membrane Interactions. 4.1. The Physical Nature of Binding between 60 S Subunits and

Membrane............. . .............. . 4.2. Interaction between Nascent Polypeptides and Membranes .. 4.3. Direct Interaction between Messenger RNA and Membranes

5. Heterogeneity in the Function of Rough Endoplasmic Reticulum with Respect to Protein Synthesis . 5.1. Compartmentalization of the Synthesis of Proteins Destined

for Discharge to the Extracellular Environment 5.2. Compartmentalization of Protein Synthesis in the

Endoplasmic Reticulum and Specific Posttranslational Modifications ........ .

5.3. Compartmentalization of the Synthesis of Specific Proteins at Discrete Sites within the Rough Endoplasmic Reticulum

6. Conclusions . 7. References ..

117 118

119 121 125 125

127

128 136 137

137 143 147

148

148

149

151 154 155

Contents xvii

Chapter 4 Biogenesis of Peroxisomes and Glyoxysomes

J. Michael Lord

1. Introduction .... ..................... 171 2. Morphology and Topographical Relationship to Other Cellular

Organelles ... ....... 172 3. Biochemical Properties and Metabolic Roles . 174

3.1. Liver Peroxisomes ... 174 3.2. Leaf Peroxisomes . . . . . . . . . . . . . 181 3.3. Fatty Seed Glyoxysomes 184

4. Microbody Proliferation. ........... 185 4.1. Liver Peroxisomes .. · ........................ 185 4.2. Leaf Peroxisomes . . . . . . . . . . . . . . . . . . . . . . . 186 4.3. Fatty Seed Glyoxysomes . . " ....... 187

5. Models for the Synthesis of Microbody Components and Their Transfer to the Organelles . , . . . . . . . . . . . . . 187 5.1. Membrane Lipids ... · ........... 188 5.2. Membrane Proteins 191 5.3. Implications for Microbody-Membrane Biogenesis. 193 5.4. Microbody Matrix Proteins .. 194 5.5. Implications for Microbody Matrix Protein Segregation 197

6. The Synthesis of Microbody Components 197 6.1. Membrane Lipids. 197 6.2. Membrane Proteins · ...... 198 6.3. Matrix Proteins. , . . . . . . 201

7. References. . . . . . . . . . . ......... 203

Chapter 5 The Subcellular Biochemistry of Thyroid

H. J. Hilderson, G. Van Dessel, A. Lagrou, and W. Dierick

1. Introduction............................. . . . . 213 2. Cell Fractionation. 215

2.1. Disruption and Homogenization of Thyroid Tissue. 215 2.2. Localization of Marker Enzymes in Thyroid (Preliminary

Studies) . . . . . . . . . . . . . . . . . . . . . . 216 2.3 . Differential Pelleting 218 2.4. Gradient Centrifugation Studies. 221 2.5. Localization of Biochemical Markers (Supplementary

Studies) . . 227

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3. Localization of Enzymes and Constituents in Bovine Thyroid Tissue. 231 3.1. Subcellular Localization of Lipolytic Enzymes 231 3.2. Subcellular Localization of Peroxidase Activities 232 3.3. Subcellular Localization of RNA-Polymerase Activity. . 234

4. Isolation and Characterization of Thyroid Organelles, Subcellular Components, and Membranes . . . . . . . . . . . . . . . . . . . . . . . . . . 238 4.1. Nuclei . . . . . . . . . . . . . 238 4.2. Mitochondria and Lysosomes 241 4.3. Golgi-Rich Fractions 243 4.4. Protein-Synthesizing Polyribosomes . . . . . . . . 244 4.5. Plasma Membranes 245

5. Summary. . 251 6. References. 259

Chapter 6 The Molecular Organization of NADH Dehydrogenase

C. I. Ragan

1. Introduction.. 267 1.1. The Purpose of This Chapter. 267 1.2. Definitions and Terminology. 268 1.3. The Functional Unit. 269

2. The Protein Components of NADH Dehydrogenase. 271 2.1. Fragmentation of the Enzyme. 271 2.2. Fragmentation by Treatment with Chaotropic Agents. 272 2.3. Polypeptide Composition of NADH Dehydrogenase and Its

Subfragments 272 3. The Protein Structure of NADH Dehydrogenase. . 279

3.1. General Properties of Multisubunit Enzymes 279 3.2. The Nature of Chaotropic Resolution. 280 3.3. Isoelectric Points of the Constituent Polypeptides. . . . . . . . 281 3.4. Labeling with Hydrophilic Probes. . 282 3.5. Labeling with a Hydrophobic Probe. 285 3.6. Proteolytic Digestion 286 3.7. Specific Structure/Function Relationships. 289

4. The Phospholipid Components of NADH Dehydrogenase. . 292 4.1. Are Phospholipids Essential? . 292 4.2. Phospholipid Composition of NADH Dehydrogenase. 293 4.3. Phospholipid Function 293 4.4. Specific Lipid-Protein Interactions. 296

Contents xix

5. Organization of NADH Dehydrogenase in the Membrane 298 5.1. Transmembranous Organization. ............... 298 5.2. Lateral Organization 300

6. Conclusion. ........ . . . . . . .. ....... 301 6.1. A Model of NADH Dehydrogenase Structure 301 6.2. Prospects . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 303

7. References. . . . . . . . 304

Chapter 7 Structure-Function Relationships of Micrococcus lysodeikticus Membranes: A Bacterial Membrane Model System

Milton R. J. Salton

1. Introduction ......... . 2. Ultrastructure of Bacterial Membranes 3. Biochemical Characterization of Micrococcus lysodeikticus

Membranes ... 3.1. Enzyme Distribution ................... . 3.2. Enzymes Involved in Wall-Polymer Biosynthesis and

Peptidoglycan Metabolism. 3.3. Lipomannan Biosynthesis. 3.4. Enzymes Involved in Lipid Biosynthesis. 3.5. Electron-Transport Chain Components 3.6. Membrane Adenosine Triphosphatase (Fj-ATPase) ....... .

4. Antigenic Architecture of the Membrane of M. Lysodeikticus 5. Summary and Conclusions. 6. References ...

Some Recent Books in Cell Biochemistry and Biology

1. Cell and Membrane Biology 2. Genetics and Viruses ........ . 3. Muscle and Ca2+ Transport. 4. General Biochemistry .

Index

309 312

320 320

323 325 328 331 336 352 366 368

376 384 388 390

395

Chapter 1

Biogenesis of Ribosomes in Eukaryotes

A. A. Hadjiolov Department of Molecular Genetics Institute of Molecular Biology Bulgarian Academy of Sciences Sofia, Bulgaria

1. INTRODUCTION

The biogenesis of ribosomes in eukaryotes involves the generation in the cell of the constituent core rRNA molecules and their interaction with about 80 distinct proteins to form the two mature ribosomal particles. Therefore, the ribosome is not only the principal organelle in gene expression, but it is also one of the best-understood models of the structure, function, and coordination of a large number of genes operating in the eukaryotic cell. Furthermore, ribosomes are made in the nucleolus, but they operate in the cytoplasm. In this respect, studies on the biogenesis of ribosomes contribute to our understanding of the molecular architecture of the cell.

This review focuses on the structure and transcription of ribosomal genes, the assembly of preribosomes, and their maturation and the transcriptional and posttranscriptional control mechanisms in ribosome biogenesis. In these fields important observations have been made in the last few years. Several comprehensive reviews cover various aspects of the problem (Busch and Smetana, 1970; Attardi and Amaldi, 1970; Maden, 1971; Craig, 1974; Warner, 1974; Perry, 1976; Hadjiolov and Nikolaev, 1976). Because of the rapid expansion of the studies on ribosome biogenesis, this review emphasizes the more recent work.

Abbreviations used in this chapter: (L-rRNA, S-rRNA) rRNA of the mature large and small ribosomal particle, respectively; (pre-rRNA) precursor to mature rRNA; (preribosome) ribonucleoprotein particle containing pre-rRNA; (pre-mRNA) precursor to mRNA; (r-chromatin) chromatin containing rRNA genes; (r-protein) structural protein of the large and small ribosome; (np) nucleotide pairs in DNA; (knp) 1000 nucleotide pairs.

2 A. A. Hadjiolov

More complete information on earlier findings can be obtained from the reviews quoted above and throughout the text.

2. RIBOSOMAL GENES

The genes involved in ribosome biogenesis include (1) the genes of the four known rRNA species: S-rRNA, L-rRNA, 5.8 S rRNA, and 5 S rRNA; (2) the genes coding for the structural ribosomal proteins (there are about 80 such proteins); and (3) the genes for the specific enzymes and other proteins specialized in transcription, mo .. lification, and processing mechanisms leading to mature ribosomes.

Practically nothing is known about the ribosomal genes from groups (2) and (3). If the information derived from studies with E. coli (Nomura et al., 1977) is valid as well for eukaryotes, we can expect that the genes for ribosomal proteins are present in single (or a few) copies clustered in the genome. Evidence for such clustering was obtained in studies with Saccharomyces carlsbergensis (Mager et aI., 1977). Genetic studies with Drosophila melanogaster showed that the genes for at least seven ribosomal proteins are located in the X chromosome together with the rRNA genes (Steffensen, 1973). This finding could be correlated with the observation that Xenopus laevis anucleolate (Onn) mutants (lacking rRNA genes) do not produce ribosomal proteins (Hallberg and Brown, 1969), although selective inhibition of rRNA genes in HeLa cells by low doses of actinomycin D did not by itself alter the synthesis of ribosomal proteins (Warner, 1977). Whereas these studies provide attractive hints, better tools are obviously needed to dissipate the darkness enveloping our knowledge of ribosomal protein genes.

In contrast, the genes that code for rRNA species in eukaryotes are among the best studied. Moreover, in the last few years important new information has rapidly accumulated. Earlier knowledge is analyzed in several competent reviews (Birnstiel et al., 1971; Reeder, 1974; Tartof, 1975). Here, I shall consider only the aspects of rRNA genes more directly related to my subject. It is firmly established that in eukaryotes the genes for S-rRNA, L-rRNA, and 5.8 S rRNA are adjacent and transcribed in a single pre-rRNA molecule, i.e., they form a transcription unit. For this reason, some authors prefer the term prerRNA gene(s). To avoid confusion, I shall continue to designate this group of genes as the rRNA genes, while considering separately the 5 S rRNA genes.

2.1. Ribosomal RNA Genes

The rRNA genes are present in one to several hundred copies, clustered in the genome. During mitosis the clusters of rRNA genes are seen in most cases

Biogenesis of Ribosomes 3

as "secondary constrictions," located at specific sites in one or a few chromosomes. The rRNA genes and the respective chromosomal site (the "nucleolus organizer") possess the capacity to form a nucleolus at telophase. Thus the active rRNA genes are unique in their association in the interphase nucleus with easily identifiable structures. It is considered that the rRNA genes are physically linked to the DNA of a given "nucleolar" chromosome. Yet in several cases multiple extrachromosomal rRNA genes have been found.

2.1.1. Chromosomal Location

Clustering of rRNA genes in the chromosome is typical of prokaryotes (Nomura et al., 1977), and this pattern is perpetuated in eukaryotes, although the number of gene sets per cell is at least one order of magnitude higher. Thus primitive eukaryotes from the Saccharomyces group contain 100-120 rRNA genes per haploid genome (Retel and Planta, 1968; Schweizer et al., 1969; Philippsen et al., 1978). Experiments with monosomic (2n - 1) strains of Saccharomyces cerevisiae demonstrated that about 70% of the rRNA genes are confined to chromosome I, which was shown to yield a DNA of about 4-4.5 X 108

daltons (Finkelstein et aI., 1972; 0yen, 1973; Kaback et al., 1973). Estimations based on a molecular weight for one rRNA gene set of 5.9 X 106 (Philippsen et al., 1978) indicate that almost the entire chromosome I is constituted by clustered rRNA genes. This conclusion is confirmed by an independent genetic analysis approach that showed that about 90% of the rRNA genes are contained in a single chromosome (Petes and Botstein, 1977) identified, however, as chromosome XII (Petes, 1979a,b). The precise physical linkage of the rRNA gene cluster remains to be clarified, but strong evidence exists that clusters of 5-15 rRNA gene sets b'DNA) are separated by aDNA segments, which do not hybridize with rRNA (Kaback et aI., 1973; Cramer et al., 1976).

The chromosomal locations of rRNA gene clusters have been established for a broad variety of higher eukaryotes. Evidence that nucleolar genes are clustered in one Zea mais chromosome (chromosome 6) was obtained already in the pioneer work of McClintock (1934). Further studies with numerous plant and animal species demonstrated that in many cases a single chromosome carries the "nucleolus organizer," although up to six such "nucleolar" chromosome pairs were identified in some species (Busch and Smetana, 1970; Lima-da-Faria, 1976). Introduction of the in situ RNA-DNA hybridization method (Gall and Pardue, 1969), supplemented by the simpler Ag staining technique (Goodpasture and Bloom, 1975) permitted more precise studies on the chromosomal location of rRNA genes. The in situ hybridization analyses provided direct evidence that rRNA gene clusters are found in only one or a few chromosomes, where they are integrated in the "nucleolus organizer." These studies also showed that all true nucleoli in the interphase nucleus contain rDNA. However,

4 A. A. Hadjiolov

in some cases rDNA was detected at chromosome loci that do not organize a nucleolus (Pardue et aI., 1970; Batistoni et al., 1978), thereby suggesting that special regulatory mechanisms should operate in switching on rRNA genes.

Comparative studies led to the generalization that karyotypes with a higher number of "nucleolar" chromosomes are a later event in evolution (Hsu et aI., 1975). A good and well-documented example is given by human chromosomes, five of which (13, 14, 15,21, and 22) carry rRNA gene clusters (Henderson et al., 1972), and a similar high number was also found in some other primates. (Tatravahi et al., 1976). Since the human haploid genome contains 50-200 rRNA genes (cf. Young et al., 1976), it may be deduced that each rRNA gene cluster includes 10-40 genes. The factors specifying the number of rRN A genes in a given chromosome remain unknown, but it should be mentioned that large variations in the distribution of rRNA genes seem to exist among human individuals (Henderson et al., 1973; Warburton et al., 1976).

As a rule, a single rRNA gene cluster is observed in a given "nucleolar" chromosome, although exceptions to this rule have been reported. For example, all 150 genes in Drosophila melanogaster are clustered in a single locus of the X or Y chromosome (see Spear, 1974), but two separate "nucleolus organizer" sites were identified in the Y chromosome of Drosophilahydei (Meyer and Hennig, 1974; Schafer and Kunz, 1975). Analysis of the location of "nucleolus organizers" within the chromosomes of a large number of plant and animal species strongly suggests that it does not occur at random. In the vast majority of cases the rRNA genes are located in the short arm of the "nucleolar" chromosome at a specific distance between the kinetochore and the telomere (Lima-daFaria, 1976).

The studies on the chromosomal location of rRNA genes clearly demonstrate that this is not a random event. The organization of the genome as a whole and the structure of constituent chromosomes are certainly important determinants. To what extent the position of rRNA gene clusters is involved in the regulation of their function remains to be elucidated.

2.1.2. Extrachromosomal Genes

The rRNA genes possess the capacity of differential replication. As a result, in some cases the cell has more or less rRNA genes than corresponds to its ploidy. Generally, the differential replication of rRNA genes takes place in order to meet unusually high demands for ribosome production raised by a given cell. The extra copies of rRNA genes may be integrated in the chromosome or remain extrachromosomal. Here, I shall consider the latter case (see also Tartof, 1975; Tobler, 1975).

The first discovered and still one of the best understood examples of differential replication is the amplification of rRNA genes (Brown and Dawid,


1968; Gall, 1968). Amplification is a process that takes place in the germ cells of a large number of species-amphibia and insects in particular (Gall, 1969). The presence of large amounts of extrachromosomal rONA in the oocytes of several species is now amply documented, and different aspects of the process are reviewed (Birnstiel et al., 1971; MacGregor, 1972; Bird et aI., 1973; Tobler, 1975; Strelkov and Kaffiani, 1978). Amplification begins during maturation of gametes-both oogonia and spermatogonia-where a relatively small amount of amplified rONA can be detected (Pardue and Gall, 1969). However, when gametes enter mitosis, most of the amplified rONA is lost. During meiosis in oocytes (but not in spermatocytes) a second round of amplification takes place resulting in the formation of a large amount of extrachromosomal rONA. Thus amplification of rRNA genes may be considered a characteristic feature in the differentiation of oocytes.

In some cases the amounts of extrachromosomal rONA in oocytes are exceedingly large. For example, the amount of amplified rONA in Xenopus laevis oocytes is about 3000-fold higher than the haploid rONA and corresponds to 1.5 X 106 rRNA genes forming 1000-1500 additional nucleoli (Birnsteil et al., 1971). Most of the amplified rONA is in the form of linear molecules, but up to about 10% is in the form of closed circles containing 1-10 rRNA genes (Hourcade et al., 1973; Rochaix et al .. 1974; Buongiorno-Nardelliet aI., 1976). Different replicative forms, including "tailed" circles are observed leading to ingenious suggestions on the mechanisms of rONA amplification (see Reeder, 1974; Tartof, 1975). At present, there is little doubt that amplification is an extrachromosomal process, most likely occurring by a "rolling circle" mechanism (see Tobler, 1975). Two points remain controversial: (1) Is the first extrachromosomal rRNA gene formed by selective excision or by selective replication? and (2) Is the partial reintegration of amplified rONA into chromosomes possible during embryogenesis? Independently of the solution of these intriguing questions it should be stressed here that the existence of amplified extrachromosomal rONA is now documented in the oocytes of many species. Hence the presence of closed circles and extrachromosomal rRNA genes has been demonstrated in the oocytes of the newt Triturus alpestris (Scheer et al., 1976) and of several insects: Dytiscus marginalis (Gall and Rochaix, 1974; Trendelenburg et al., 1974), Acheta domesticus (Trendelenburg et al., 1976), Colymbetesfuscus (Gall and Rochaix, 1974), and others.

Drosophila melanogaster offers an interesting case of differential replication of rRNA genes, which has been studied in detail (see Spear, 1974; Tartof, 1975). Genetic analysis has shown that when the dose of rRNA genes in the "bobbed" (bb) locus of an X chromosome is below a critical level, differential replication of rRNA genes and compensation of their number takes place in somatic cells during ontogenesis. Analysis of this phenomenon indicates that the compensatory rONA copies may not be completely integrated in the chromo-

6 A. A. Hadjiolov

some. Indeed, circular rDNA molecules were identified in the germ cells of male flies, thereby indicating that the existence of extrachromosomal rDNA may be a stage in this compensation process (Graziani et aI., 1977). Elucidation of the precise mechanisms of extrachromosomal rDNA formation in Drosophila is not yet completed. In this respect it is worth mentioning that extrachromosomal rRNA genes were observed without apparent relation to differential replication mechanisms. Thus, a high percentage of extrachromosomal rDNA is found in Drosophila melanogaster that are females heterozygous for an X chromosome inversion transposing the rRNA gene cluster to the telomere region of the X chromosome (Zuchowski and Harford, 1977; Harford and Zuchowski, 1977).

The wider occurrence of extrachromosomal rRNA genes is supported further by recent studies with some primitive eukaryotes. Analysis of the location of rRNA genes in Tetrahymena pyriformis showed that only a single gene is integrated in the micronucleus chromosomes (Yao and Gall, 1977). The remaining rRNA genes [about 200 per haploid equivalent (Engberg and Pearlman, 1972)] are extrachromosomal and located in the macronucleus. There is also evidence for the occurrence of a large number of extrachromosomal rRNA genes in Physarum polycephalum (Vogt and Braun, 1976) and Stylonychia (Prescott et aI., 1973; Lipps and Steinbruck, 1978).

The widespread occurrence of extrachromosomal rRNA genes in eukaryotes strongly suggests that differential replication of these genes may play an important role in maintaining the genetic balance of rRNA genes. The generation of functional linear or circular extrachromosomal rRNA genes is obviously an important mechanism ensuring adequate production of ribosomes-not only in oocytes, but possibly in some somatic cells as well. Reintegration of some of these rRNA genes in the chromosomal rRNA gene cluster provides an attractive, yet still unproved, possibility.

2.1.3. Organization and Structure

Several powerful techniques introduced in the last few years resulted in a booming expansion of our knowledge on the organization and structure of rRNA genes in eukaryotes. Isolation, cloning, and mapping of rRNA genes from an ever-increasing number of organisms has been achieved. Combined with electron-microscopic techniques of RNA-DNA hybridization and visualization of active rRNA genes, these techniques now permit us to outline in considerable detail the basic features of the organization of rRNA genes. Here I shall summarize the presently available information on the best studied and most typical cases.

The rRNA genes are organized in transcription units. A transcription unit (transcripton) is a DNA segment containing a set of genes (or a single gene) transcribed as one RNA molecule. The rRNA transcription unit contains two


elements: (1) the sequences corresponding to the mature rRNA species, and (2) transcribed spacer (tS) sequences, which are transcribed as part of the initial pre-rRNA molecule, but are not present in the mature rRNA species. The multiple sets of rRNA transcription units are arranged in tandem along the DNA separated by nontranscribed spacer (ntS) sequences. One transcription unit and the adjacent nontranscribed spacer constitute the repeating unit.

The tandem arrangement of a varying number of repeating units constitute the basic principle in the organization of both rRNA and 5 S rRNA genes existing as either chromosomal or extrachromosomal gene sets.

a. Saccharomyces cerevisiae. Treatment of yeast DNA with appropriate restriction endonucleases permitted the isolation of fragments containing the rRNA genes. Several such fragments were cloned and mapped leading to the elucidation of the yeast repeating unit structure (Figure 1). The results obtained by different groups (Nath and Bollon, 1977, 1978; Cramer et ai., 1977; Bell et aI., 1977; Philippsen et ai., 1978) are in good general agreement with each other and may be summarized as follows: The yeast rRNA repeating unit (8.9-9.1 knp) contains the sequences coding for 5 S rRNA and for primary pre-rRNA (containing 18,5.8, and 25 S rRNA sequences). There is no detectable length heterogeneity among individual repeating units, a fact suggesting that ntS sequences in yeasts are identical in size. Although the genes for 5 S rRNA and rRNA are in one repeating unit, their coding sequences are in different DNA strands and are transcribed in opposite directions (Aarstad and 0yen, 1975; Kramer et al., 1978). The mapping of the sequences corresponding to the separate mature rRNA reported from different laboratories is identical. Unfortunately, hybridization with the respective pre-rRNA was not carried out, thereby leaving some uncertainty about their exact position. The molecular weight of the largest pre-rRNA in Saccharomyces cerevisiae was estimated as

I I I I I i I i 2 3 4 5 6 7 8 9 knp

~ 185 5.85 25$

[==~-------.----~ RU nt 5 -tSe tS i

fA ~~ ~G~C I~~ A I FIE I EcoR1

t t t t t + FIGURE 1. Organization of the rRNA + 5 8 rRNA repeating unit (RU) of Saccharomyces cerevisiae. The hatched regions in the fragments (A - E) obtained upon cleavage with EcoRI restrictase (arrows) have been sequenced. 5 8, 18 8 , 5.8 8, and 25 8 designate the segments coding for the respective mature rRNA; nt8, nontranscribed spacer; t8" external transcribed spacer; t8" internal transcribed spacer. The location of the 5' end of the rRNA transcription unit is uncertain.

8 A. A. Hadjiolov

2.5 X 106 (Udem and Warner, 1972) or 2.8 X 106 (Dudov et aI., 1976) indicating that a sizable part of the DNA between the 5 S rRNA and rRNA loci contains tS sequences. Consequently, ntS sequences in yeasts could encompass no more than 8-19% of the repeating unit.

The presence of both rRNA and 5 S rRNA genes in one repeating unit seems to be limited to only a few lower eukaryotes. The organization of rRNA genes in Saccharomyces carlsbergensis (cf. Meyerink and Retel, 1977) shows remarkable similarities with that in Saccharomyces cerevisiae. An analogous type of organization of rRNA genes is found also in Dictyostelium discoideum, although in this case the repeating unit (about 38 knp) is markedly larger (Maizels, 1976).

b. Drosophila melanogaster. Drosophila contains about 150 rRNA genes clustered in the X chromosome and a somewhat smaller number in the Y chromosome (see Tartof, 1975). Cloning of these genes (Glover et al., 1975) led to several important observations on their organization. The "normal" repeating unit is 10.5-12.5 knp in length (Glover and Hogness, 1977; Wellauer and Dawid, 1977; Dawid et al., 1978). In this case a limited length heterogeneity in ntS is found ranging from 3.3 to 5.4 knp (Wellauer and Dawid, 1978). It is also noteworthy that the ntS sequences are internally repetitious as observed earlier in higher animals (see below). The transcription units in Drosophila melanogaster are organized as in all other eukaryotes, although in this case the external transcribed spacer (tSe) sequences are shorter than the internal transcribed spacer (tS;) sequences (Figure 2).

The most striking feature of Drosophila melanogasterrRNA genes is the finding of nonribosomal insertions ("introns") splitting the L-rRNA gene into two unequal parts (Glover and Hogness, 1977; White and Hogness, 1977; Wellauer and Dawid, 1977; Pellegrini et aI., 1977). Similar insertions are found also in Drosophila hydei (W. Kunz, personal communication), suggesting that they

ntS tSe S-rRNA tSj L-rRNA (3.6)

c:=:::-----:::::::=:::r;:! ~-~-~:J.-IIIII!I~IIIIIIiI·i •• 3.3- 5.4 0.6 1.8 1.2 2.45 a 1.15

Ins~~:6 5.0

FIGURE 2. Organization of the rRNA repeating unit of Drosophila melanogaster. The segments coding for mature S-rRNA, 5.8 S rRNA, and L-rRNA are the black bars. Ins designates the three main types of nonribosomal DNA insertions in the L-rRNA gene. The white dot in the L-rRNA gene indicates the position of the gap in the polynucleotide chain of mature L-rRNA. The indicated size of the separate segments (in knp) is based mainly on values reported by Wellauer and Dawid (1977) and Dawid et al. (1978), corroborated with results on the molecular weight of Drosophila pre-rRNA (Levis and Penman, 1978) in the case of the tS, and tS; segments. Precise values and their variations may be found in the original publications.


are typical of this genus. The insertions in Drosophila melanogaster are of different lengths, resulting in rRNA repeating units that are 0.5, 1.0, and 5.0 knp longer than the "normal" ones (White and Hogness, 1977; Wellauer and Dawid, 1977,1978; Dawid et al., 1978). There is also a second minor type of insertion that is not homologous to the major type (Dawid et al., 1978). It is remarkable that the insertions are more frequent in the X chromosome rRNA genes, where 65% of the repeating units have insertions of the major type (49%) or the minor (16%) type. On the other hand, only 16% of the Y chromosome rRNA genes have insertions, mostly of type 2 (Wellauer et al., 1978). It is also of interest that these nonribosomal insertions belong to a repeated class of DNA sequences encountered outside the rRNA repeating units with an estimated total amount in the genome equivalent to about 400 knp (0.2%) of its DNA (Dawid and Botchan, 1977). The role of the insertions splitting Drosophila rRNA genes remains unknown. Studies with cultured Drosophila cells failed to detect any pre-rRNA larger than the normal 34 S (2.7 X 106 daltons) pre-rRNA (Levis and Penman, 1978). Therefore, unlike the split structural genes coding for eukaryotic mRNA, the split rRNA genes in Drosophila may not be transcribed. If this turns out to be the case, then insertion-mediated shutoff of rRNA genes may have some relation to the capacity for differential replication typical of the X chromosome rRNA genes (Williamson and Procunier, 1975). In any case, it seems at present that splitting of rRNA genes is a relatively rare event. Studies with another insect, Bombyx mori not only did not uncover splitting of rRNA genes, but also established that the repeating units (6.9 X 106 daltons) are remarkably homogeneous in length (Manning et al., 1978). On the other hand, a 400-np insertion was discovered in the macronucleus L-rRNA gene of Tetrahymenapigmentosa. Evidence for the transcription and subsequent splicing of this inserted sequence was obtained (Wild and Gall, 1979).

c. Xenopus [aevis. Studies with Xenopus laevis have contributed considerably to the elucidation of the organization, structure, and regulation of rRNA genes. Owing to peculiarities in the structure of rRNA genes (i.e., the markedly higher GC content of ntS segments) and the availability of anucleolate (OnJ mutants, the 450 (haploid) rRNA gene cluster in Xenopus laevis supplied the starting material for the first unequivocal isolation of pure rRNA genes (see Birnstiel et al., 1971). It is also with Xenopus laevis that the first rDNA repeating unit was characterized as a DNA segment of about 8.7 X 106 daltons, or 13 knp (Wensink and Brown, 1971). Recent studies led to a deeper understanding of the organization of Xenopus laevis repeating units, perhaps a typical model of repeating units in other higher eukaryotes (Figure 3).

The organization of the transcription unit (tSe-S-rRNA-tSj-L-rRNA) in Xenopus follows the pattern typical of higher eukaryotes. Its size (approx. 7.9 knp) is apparently constant within cells and individuals of this species (Wellauer et al., 1974b, 1976a; Botchan et aI., 1977). The position of the 5.8 S

10

ABC o S-rRNA

I I I i 0,45: 0.7 ~IQ31 ~1.3 <'O'~" 1.9 , , , ,

I I , I

10------1 I I '------~

0,9

" '"

I " I ,

I " r--------- - ---------- ---, I I ~----------------------~ ~ 3,5

A. A. Hadjiolov

L-rRNA

0.7 4.5

FIGURE 3. Organization of the rRNA repeating unit of Xenopus laevis. A-D designate ntS segments of constant (A and C) or variable (B and D) length (Wellauer el al., 1976a,b). The length (in knp) assigned to the C and D segments is arbitrary, owing to uncertainty about the size of their boundary (Botchan et al., 1977). The lengths of the separate segments in the rRNA transcription unit are approximate and based mainly on values for the molecular weight of Xenopus laevis pre-rRNA and rRNA (cf. Loening el al., 1969; Wellauer and Dawid, 1974).

rRNA gene between S-rRNA and L-rRNA is now conclusively proved (Speirs and Birnstiel, 1974; Walker and Pace, 1977). Considerable length heterogeneity of the ntS sequences in Xenopus laevis rRNA repeating units, even originating from a single nucleolus organizer, is consistently found (Wellauer et aI., 1974b). This length heterogeneity of ntS is also preserved in amplified rDNA (Wellauer et al., 1976b).

More detailed electron microscopic analysis of RNA-DNA hybrids (heteroduplex mapping) permitted the identification of four regions in the ntS segment (Wellauer et al., 1976a). Whereas segments A and C are constant in size, considerable variations are found in segment B (0.56-0.9 knp) and in segment D (approx. 1.3-3.9 knp). Recent detailed studies with different restriction endonucleases show a somewhat distinct structure of the ntS segment (Boseley et al., 1979). At least three repetitious areas could be identified in the regions corresponding roughly to segments B, C, and D. The first area, adjacent to segment A, contains a repeated sequence (canon) of about 100 np, whereas the second and third areas are constituted of 81/60 np canons. The important genetic implications of the established heterogeneity and the presence of repeated sequences in the ntS segment of Xenopus laevis will not be discussed here. However, it should be noted that a detailed knowledge of the structure of ntS segments is likely to provide more direct evidence on the possibility of their transcription under some physiological or pathological conditions (see Section 3.2.3).

To what extent the structure of Xenopus laevis repeating units is typical of other higher eukaryotes is not yet known. It is likely that some further "improvements" in the organization of rRNA genes-ntS segments in particular-may have evolved in evolution. For example, the size of rRNA repeating units in mice (approx. 40 knp) is markedJy higher, largely because of the increased length of ntS segments (Arnheim and Southern, 1977; Cory and Adams, 1977).


d. Tetrahymena pyriformis. The extrachromosomal rRNA genes in the Tetrahymena macronucleus have attracted considerable interest because they present a rather intriguing organization. About 90% of these genes constitute a homogeneous population of linear rRNA molecules with 12.6 X 106

mol. wt., or about 19 knp. Most of the remaining 10% are circles of the same size (Gall, 1974; Engberg et al., 1974). Analysis of the structure of linear rONA molecules shows that they are built of two identical halves (9.5 knp each) constituting a giant palindrome (Karrer and Gall, 1976; Engberg et al., 1976). The detailed study of this rONA showed that each half contains one transcription unit plus ntS sequences, and thus follows the general pattern of rRNA repeating unit structure (Figure 4). The two transcription units (:::::: 7.2 knp each) have their S-rRNA sequences proximal to the center of the molecule (Engberg et al., 1976). Consequently, divergent transcription of opposite coding strands, starting from centrally located promotors, has been postulated and supported experimentally (Grainger and Ogle, 1978). It is shown that replication of Tetrahymena linear rONA is also divergent (Truett and Gall, 1977). The precise size of the central control sequences is unknown, but apposition of data on the size of primary pre-rRNA (Pousada et aI., 1975) and hybridization of L-rRNA with EcoRl fragments (Engberg et aI., 1976) strongly suggests that in each repeating unit they would encompass only a few hundred nucleotide pairs. The distal ntS sequences ( - 2 knp) also reveal an unusual structure. At approx. 1 knp from the end of the transcription unit a tandemly repeated sequence 5'-[C-C-C-C-A-Aln-3' is found in the coding strand, where n is between 20 and 70 (Blackburn and Gall, 1978). The role of this sequence is unknown, but its location suggests that it is unlikely to be a transcription termination signal.

The palindromic organization of extrachromosomal rRNA repeating units is found also in Physarumpolycephalum (Vogt and Braun, 1976; Molgaard et

rs I nff L tSi S I 5' l-I

S' 10.8 2.0 0.6 3.8 -to 0.4 I. 9.S

FIGURE 4. Organization of the palindromic extrachromosomal rONA of Tetrahymena pyriformiS . This linear molecule is constituted by two rRNA repeating units of about 9.5 knp each (Engberg et al., 1976; Karrer and Gall, 1976). The two transcription units are divergent, opposite + rONA strands coding for the respective pre-rRNA (bars). Land S (black bars) designate the segments coding for L-rRNA and S-rRNA. rs, Repeated sequences in the ntS (Blackburn and Gall, 1978). The values (in knp) assigned to separate segments in the rRNA repeating units are approximate, based mainly on data for the molecular weight of Tetrahymena pre-rRNA and rRNA (cf. Pousada et al., 1975) .

12 A. A. Hadjiolov

al., 1976). In this case the repeating unit is markedly longer (28-29 knp), and a large ntS segment (15 knp) is located at the center of the rDNA molecule (Vogt and Braun, 1976; Hall and Braun, 1977; Steer et al., 1978). The relation of these inverted repeat, extrachromosomal rDNA structures with the tandem arrangement of repeating units in higher eukaryotes (but also in Saccharomyces) remains to be clarified.

2.2. 5 S rRNA Genes

2.2.1. Chromosomal Location

The genes for 5 S rRNA are present in multiple copies, and in most cases their number is markedly higher than that of rRNA genes. The use of in situ hybridization and other techniques made possible detailed studies on their chromosomallocalization in a large number of eukaryotes. These studies indicated a remarkable evolutionary tendency of 5 S rRNA genes to drift apart from rRNA genes. Thus in Escherichia coli the 5 S rRNA genes are in one transcription unit with the genes for S-rRNA and L-rRNA (see Nomura et al., 1977). In Saccharomyces the 5 S rRNA genes are equal in number and are interspersed with rRNA genes in chromosome XII (Rete! and Planta, 1972; Rubin and Sulston, 1973; Kaback et al., 1976). In fact, they are in one rRNA repeating unit, although transcribed separately. The linkage of 5 S rRNA and rRNA genes appears to have been lost early in evolution. Already in some of the lower eukaryotes [e.g., Tetrahymena pyriformis (T0nnesen et al., 1976) and Physarum polycephalum (Hall and Braun, 1977)] the multiple 5 S rRNA genes are no longer linked to rRNA genes. This pattern is fully displayed in higher eukaryotes, where clusters of 5 S rRNA genes are usually located at other than the "nucleolar" chromosomes. The 5 S rRNA gene cluster may be on a single chromosome, as in Drosophila melanogaster, where it is in band 56F of chromosome 2 (Wimber and Steffensen, 1970; Quincey, 1971) and in Drosophila hydei (Alonso and Berendes, 1975), although multiple loci for 5 S rRNA genes are found in other species of Drosophila (Cohen, 1976; Wimber and Wimber, 1977). In humans, the bulk of 5 S rRNA genes is packed in chromosome 1, whereas smaller clusters may exist in chromosomes 9 and 16 (Steffensen et al., 1974; Johnson et al., 1974). A similar clustering of 5 S rRNA genes is found also in some Primates (Henderson et al., 1976). An extreme case seems to be Xenopus laevis, where 5 S rRNA gene clusters are located at the ends of most of its 18 chromosomes (Pardue et al., 1973). Again, such a wide spreading of 5 S rRNA genes is not found in other Amphibia species (Pilone et al., 1974; Leon, 1976). Obviously more extensive taxonomic studies are needed in order to unravel regularities in the chromosomal location of 5 S rRNA genes.


2.2.2. Organization and Structure

The multiple copies of Xenopus laevis 5 S rONA (approx. 24,000 per haploid genome) supplied a convenient system for studying their organization and structure. Analysis of the primary structure of 5 S rRNA from many eukaryotes provided evidence that it is strongly conserved in evolution (see Erdmann, 1980). Therefore, studies on the organization of 5 S rONA in Xenopus laevis are likely to provide a reliable model valid for other eukaryotes as well.

The clusters of 5 S rRNA genes in Xenopus laevis are composed of repeating units arranged in tandem. Sequence analysis of 5 S rRNAs reveals that somatic cells synthesize a 5 S rRNA distinct by several nucleotides from oocyte 5 S rRNA (Wegnez et al., 1972; Ford and Southern, 1973). Therefore, two types of differentially expressed 5 S rRNA genes should exist in Xenopus laevis. These genes are designated as somatic (Xis) and oocyte (Xlo) 5 S rONA. Recently, a second class of minor oocyte 5 S rONA (with 2000 copies per haploid genome) (Xlt) was identified (Brown et al., 1977). It was shown that the Xlt 5 S rRNA repeating unit is markedly shorter and its spacer regions do not cross-hybridize with Xlo 5 S rONA. A similar situation is found when oocyte rONA from Xenopus laevis and Xenopus borealis (formerly Xenopus mulleri) are analyzed (Brown and Sugimoto, 1973). These results show that spacer sequences in 5 S rRNA genes are subject to a remarkably rapid divergence. Unfortunately, Xis 5 S rONA has not yet been isolated. Comparative studies with the three distinct 5 S rRNA repeating units could provide important information on the extent of heterogeneity of spacer sequences within a single species.

Recently, deeper studies on the structure of Xlo 5 S rONA became possible and provided several important observations. The Xlo 5 S rRNA repeating unit (Figure 5) is about 700 np long, and its complete nucleotide sequence is now known (Fedoroff and Brown, 1978; J. R. Miller et al., 1978; J. R. Miller and Brownlee, 1978). The repeating unit has two distinct regions of differing nucleotide composition. The ntS (approx. 360 np) is AT-rich, whereas the remaining part of the molecule (containing the 121 np of the 5 S rRNA gene) has a high GC content. Much of the ntS is constituted by the severalfold repeated sequence in the noncoding strand 5'-[C-A-A-A-G-T-T-T-G-A-G-TT-T-T]-3' extending to about 50 nucleotide pairs before the beginning of the 5 S rRNA gene. A remarkable feature of Xenopus laevis 5 S rONA is the presence of a "pseudogene," which has a sequence identical to that of the 5 S rRNA gene, but lacks the last 20 nucleotide pairs (Jacq et al., 1977). Whether the "pseudogene" is ever transcribed remains unanswered, and its significance is as yet completely unknown.

The sequences flanking the 5 S rRNA gene are of considerable interest, as

14 A. A. Hadjiolov

Hind III Hind III

- - - - - - - - - - A + T - - - - - - - -¥- - - - - - G + C - - - - - or I , I : ,

~ae,J11 Hoe III : I I \ I r I, \ I I

---- - -----400---- - - - - -r;..58*---193-----+43, I r I I

I: 'I I : II I I

I: I : :

I: I r

1 : r 121 * 101W I i , I

: Ge n e! Pseudogene 5': :3'

5S RNA 121

FIGURE 5. Organization of the 5 S rRNA repeating unit of Xenopus laevis. The position of ntS (black bars) and 5 S rDNA gene and "pseudogene" (white bars) is indicated. Hind III (-) and Hae III (- - -) designate cleavage sites for the respective restrictases. [Reproduced from J. R. Miller and Brownlee (1978) with permission of the authors and Nature (London).]

they should contain putative promotor and terminator signals. The importance of these sequences is evident from the fact that, in comparison with other ntS sequences, they are strongly conserved (1. R. Miller and Brownlee, 1978). Nevertheless, examination of sequences in this region failed to reveal any distinctive promotor or terminator structures. It should be stressed here that the 5 S rRNA gene and its surroundings in Saccharomyces cerevisiae have been also sequenced (Maxam et al., 1977; Valenzuela et al., 1977; Kramer et al., 1978). Careful comparison of the respective "control" sequences from Saccharomyces cerevisiae and Xenopus laevis does not show any obvious similarities. Therefore, further sequencing studies on rRNA genes would be needed before valid conclusions about putative recognition signals could be safely drawn.

The organization and structure of 5 S rRNA genes in other higher eukaryotes is much less studied, but it seems that they follow the general pattern established for Xenopus laevis. For example, the 5 S rRNA genes of Drosophila melanogaster have been mapped with restriction endonucleases, and it was found that the estimated 160 copies (Tartof and Perry, 1970) are grouped in one or two clusters of about 370 np for each gene (Procunier and Tartof, 1976; Hershey et aI., 1977; Artavanis-Tsakonas et al., 1977). As in Xenopus laevis the ntS segments of Drosophila melanogaster 5 S rRNA repeating units are characterized by their high AT content (Hershey et al., 1977).


2.3. General Features

The investigations carried out in the last few years extended our understanding of the organization and structure of rRNA genes in eukaryotes. Although these studies are still in their initial stage, some general features may be outlined.

(1) Two distinct sets of multiple rRNA genes operate in eukaryotes: (a) the rRNA genes for S-rRNA, 5.8 S rRNA, and L-rRNA; and (b) 5 S rRNA genes. In most eukaryotes these two sets of genes are located in distinct chromosomes. As a rule, the number of 5 S rRNA genes is significantly higher. In some primitive eukaryotes (Saccharomyces. Dictyostelium discoideum) the two sets of genes are equal in number and are located in the same repeating unit, although transcribed separately.

(2) The rRNA and 5 S rRNA genes are always organized in repeating units usually arranged in tandem and located at specific chromosome loci. The rRNA repeating units contain nontranscribed spacer (ntS) sequences and a transcription unit containing transcribed spacer (tS) sequences and sequences corresponding to the mature rRNA species. The organization of 5 S rRNA repeating units is analogous, but in this case the existence and the size of tS sequences is still controversial.

(3) The ntS sequences display considerable heterogeneity in size and structure, not only among species, but even within different cell types. In many cases repetitious simple sequences (approx. 10-15 np) are identified in ntS segments of the repeating units. Similar simple sequences are found also in the chromosomes, outside rRNA gene clusters. These and other observations suggest that ntS sequences may be involved in meiotic (and possibly mitotic) recombination events directing the chromosome location and the maintenance of sequence homogeneity of the multiple of rRNA repeating units. In addition, the ntS contains shorter and highly conserved sequences flanking the transcription unit. These regions contain putative promotor, initiation, termination, and other control signals. The available information does not allow definitive conclusions about the existence and the physical nature of such signals.

(4) The transcription unit of all rRNA repeating units studied until now displays a common pattern of organization. Constituent sequences follow the order: [promotor]-tSe-S-rRNA-tSj-5.8 S rRNA-tSj-L-rRNA. The 5.8 S rRNA gene is invariably located within the internal transcribed spacer. The sequences corresponding to mature rRNA are markedly more conserved in evolution that in the ntS and even tS sequences. This fact indicates that considera ble genetic pressure exists to maintain constant the primary structure of r RN A genes. The character and the operation of postulated rectification mechanisms

16 A. A. Hadjiolov

maintaining identical the multiple copies of rRNA genes are still conjectural. The transcription unit of 5 S rRNA repeating units coincides with the 5 S rRNA gene, but it may include a few nucleotide pairs distal of the gene.

(5) Eukaryotes possess the capacity for differential replication of their rRNA genes. Amplification of rRNA genes in the oocytes of many amphibia and insects results in the formation of up to several thousand extrachromosomal rRNA repeating units. Differential replication of rRNA genes is observed also in some somatic cells as shown by magnification and polytenization phenomena in Drosophila. Formation of extrachromosomal rRNA repeating units and the possibility of their reintegration in the chromosomes of germ or possibly somatic cells is an attractive field for further exploration. The production of a large number of extrachromosomal rRNA genes organized as giant palindromes is another form of differential replication exploited in some lower eukaryotes (e.g., Tetrahymena, Physarum, Stylonychia). It is not yet known whether 5 S rRNA genes are also subject of differential replication mechanisms.

In summary, the complex organization and structure of rRNA and 5 S rRNA genes outlined above creates conditions for their adequate replication and maximally efficient transcription. Inclusion of S-rRNA, 5.8 S rRNA, and L-rRNA genes in one transcription unit ensures-even at the gene level-synchrony in the formation of the two ribosomal particles. Why the 5 S rRNA genes have drifted apart from rRNA genes is totally unknown.

3. TRANSCRIPTION OF RIBOSOMAL RNA GENES

3.1. Components of the Transcription Complex

3.1.1. RNA Polymerases

Transcription is catalyzed by specific enzymes-DNA-dependent RNA polymerases. The information about eukaryotic RNA polymerases has been extensively reviewed (Jacob, 1973; Chambon et al., 1974; Chambon, 1975; Biswas et al., 1975), and only some of their properties will be briefly considered here. The RN A polymerase reaction requires the presence of all four nucleoside-5'-triphosphatesas substrates. Many synthetic or natural analogues can replace the respective nucleoside-5' -triphosphates and be incorporated into the polynucleotide chain (see Suhadolnik, 1970; Langen, 1975). The primary structure of the polyribonucleotide product is a faithful copy of the + strand of the DNA template. In most known cases the RNA chains made in vitro or in vivo start with 5'-terminal triphosphates of adenosine or guanosine. In eukaryotes, three major types of RNA polymerases-A, B, and C [I, II, and III, respectively:


see Chambon (1975)] -have been identified. There is convincing evidence that the separate RNA polymerases are specialized for the transcription of distinct types of genes: A, for rRNA genes; B, for nucleoplasmic genes; and C, for lowmolecular-weight RNA genes (including 5 S rRNA genes). All three eukaryotic RNA polymerases are multicomponent enzymes containing two highmolecular-weight and four to six (or more) low-molecular-weight subunits (see Chambon, 1975). RNA polymerase B is inhibited selectively by very low concentrations (10-8 to 10-9 M) of the Amanita phalloides toxin a-amanitin (Fiume and Wieland, 1970), which binds stoichiometrically to the enzyme molecule. Studies on the binding of 14C-Iabeled amanitin provided evidence that animal cells contain about 4-7 X 104 molecules of RNA polymerase B per haploid genome (Chambon et al., 1972). RNA polymerase C is also inhibited by a-amanitin, but at markedly higher concentrations (10-4 to 10-5 M), whereas RNA polymerase A is fully resistant. There is some indirect evidence that the exotoxin of Bacillus thuringiensis inhibits the RNA polymerase reaction by altering the enzyme, but both RNA polymerases A and B are affected (Smuckler and Hadjiolov, 1972; Beebee et al., 1972).

The specialization of eukaryotic RNA polymerases in the transcription of different types of genes is a remarkable acquisition in evolution, but the molecular recognition mechanisms involved remain unknown.

3.1.2. Nucleolar rDNA and Chromatin

In all eukaryotes the genome DNA (including rRNA genes) is complexed with histones and nonhistone proteins in order to constitute chromatin in the interphase nucleus and chromosomes during mitosis and meiosis. Therefore, it is reasonable to assume that chromatin (rather than "naked" DNA) is the actual physical structure involved in the eukaryotic transcription complex. The studies on the isolation and structure of chromatin-containing rRNA genes (rchromatin) are still at their initial stage. Characterization of chromatin from isolated nucleoli showed that only 10-20% are truly intranucleolar, and still a smaller part (about 1 %) contains rDNA (see Busch and Smetana, 1970). Thus isolation of pure r-chromatin from nucleoli of somatic cells requires additional fractionation. Methods yielding intranucleolar chromatin fractions, substantially enriched in transcriptionally active rDNA were described recently (Bachellerie et al., 1977a,b; Bombik et al., 1977) opening the way for further analytical studies.

The isolation of practically pure r-chromatin (with more than 90-95% rDNA) was achieved by the use of cells containing amplified extrachromosomal rRNA genes: Xenopus laevis oocytes (Higashinakagawa et al., 1977), Tetrahymena pyriformis (Leer et al., 1976; Mathys and Gorovsky, 1976; Jones,

18 A. A. Hadjiolov

1978a), and Physarum polycephalum (Grainger and Ogle, 1978). The r-chromatin obtained contains a large number of ribosomal and other nonhistone proteins (Higashinakagawa et ai., 1977; Jones, 1978b). A detailed analysis of Tetrahymena r-chromatin shows (Jones, 1978b) that it contains histones H2b, H3, H4, and HX (equivalent to H2a) in a ratio similar to that found in total chromatin. The extent of modification of these histones is also comparable in the two chromatin fractions. Histone HI is also present in r-chromatin, although its modification by phosphorylation may be less extensive than in bulk chromatin. The main distinctive feature of Tetrahymena r-chromatin appears to be a histone/DNA ratio of only about 40% of that in total chromatin. The difficulties inherent in chromatin isolation techniques require further studies before generalizations on the composition of r-chromatin could be attempted. At present, it appears that the basic structure of r-chromatin is similar to that of bulk chromatin.

3.2. The Transcription Process

3.2.1. Topology of Primary Pre-rRNA

All eukaryotes are characterized by the presence in the nucleolus of a pool of large pre-rRNA molecules containing the sequences of L-rRNA, 5.8 S rRNA, and S-rRNA. These primary pre-rRNA molecules are the first labeled and therefore can be considered as the initial transcription product (see below). Because the pre-rRNA molecules are a faithful copy of the + strand of the rDNA transcription unit, the arrangement of tS and rRNA sequences in prerRNA should reflect their topology in the respective rRNA gene. In many cases this arrangement is ascertained by direct electron microscopic secondary structure mapping of pre-rRNA (Wellauer and Dawid, 1975; Schibler et al., 1975; see Hadjiolov and Nikolaev, 1976). Considerable variations in the size of primary pre-rRNA exist among species, ranging from molecular weight of 1.9-2.1 X 106 in Acetabularia (Spring et ai., 1976) and 2.5-2.8 X 106 in Saccharomyces (Udem and Warner, 1972; Dudov et al., 1976) to about 4.7 X 106 in mammalian cells (see Wellauer and Dawid, 1975; Hadjiolov and Nikolaev, 1976). Independently of these large variations in size, a general pattern in the topology of primary pre-rRNA molecules (and the respective transcription unit) remained stable in evolution (Figure 6).

Identification of the polarity of primary pre-rRNA molecules remained a tedious problem for a long time. Different experimental approaches were tried leading to conflicting conclusions (see Hadjiolov and Nikolaev, 1976). Recent techniques of gene analysis provided direct evidence (Reeder et al., 1976; Dawid and Wellauer, 1976) that the following 5'- to 3'- polarity of primary pre-rRNA is typical of eukaryotes:


5' end - tSe - S-rRNA - tS; - 5.8 S rRNA - tS; - L-rRNA - 3' end

This polarity is identical to that of primary pre-rRNA in prokaryotes (see Nomura et al.. 1977), thereby suggesting that a common pattern in the sequence of transcription of rRNA genes has remained stable in evolution.

3.2.2. Morphology of Active rRNA Genes

a. General Characteristics. Introduction of the gene-spreading technique by Miller and his co-workers (0. L. Miller and Beatty, 1969a,b; see O. L. Miller and Hamkalo, 1972a,b) allowed the direct visualization of the transcription of rRNA genes. The original studies were performed with amplified rRNA genes of Triturus viridescens and other amphibian oocytes. Since then, extensive studies were carried out with numerous cells and organisms. Generally, the best results were obtained with amplified extrachromosomal rRNA genes. Thus the active rRNA genes in the oocytes of several amphibia [Triturus (0. L. Miller and Beatty, 1969a,b; Scheer et al .. 1973), Xenopus laevis (0. L. Miller and Beatty, 1969b; Scheer et al., 1977), Pleurodeles waltlii (Angelier and Lacroix, 1975)] and of some insects [Dytiscus marginalis, Acheta domesticus (Trende-

DNA

Pri mary pre-rRNA

ntS TU nts TU ntS TU ntS TU

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

S.cerevisiae (Plants)

\ Insects

H'!_~_ •. ·! H ,/

1" '" ............... .

Fishes. Amphibia

................... . ..... ....... ~-=~--. /

S-rRNA t Sj L-rRNA

Birds

Mammals

FIGURE 6. Arrangement and size of specific sequences in primary pre-rRNA (and rRNA genes) of eukaryotes. TU. transcription unit; ntS, nontranscribed spacer; tS" external transcribed spacer; tS;, internal transcribed spacer; S-rRNA and L-rRNA, sequences corresponding to mature rRNA. The dotted arrow indicates uncertainty about the coincidence between 5' end of primary pre-rRNA and the promotor site of the transcription unit.

20 A. A. Hadjiolov

lenburg, 1974; Trendelenburg et al .. 1973,1976)] have now been studied in considerable detail. Extensive studies were carried also with germ and embryonic cells of insects [Drosophila (Meyer and Hennig, 1974; Glatzer, 1975; Laird and Chooi, 1976; McKnight and O. L. Miller, 1976) and Oncopeltusfasciatus (Foe et al .. 1976)].

The gene-spreading technique was successfully applied to the study of green algae [Acetabularia. Chlamydomonas. and others (Trendelenburg et al .. 1974; Spring et al .. 1974, 1976; Berger and Schweiger, 1975a,b; Woodcock et al .. 1975; Berger et al .. 1978)]. Information about other lower eukaryotes is still limited, but the recent visualization of transcription in extrachromosomal rDNA of Physarum polycephalum (Grainger and Ogle, 1978) indicates that more extensive studies may be expected in the near future.

Visualization of active rRNA genes in mammalian cells is technically more difficult and only limited information is available. Nevertheless, active rRNA genes were observed in HeLa (0. L. Miller and Bakken, 1972) and Chinese hamster ovary (CHO) (Puvion-Dutilleul et al .. 1977a) cells, in mouse spermatids (Kierszenbaum and Tres, 1975) and in rat hepatocytes (Puvion-Dutilleul et al .. 1977b), thus allowing the observed morphological image of the transcription process to be extended to all eukaryotes.

The general features of active rRNA genes may be summarized as follows: First, the active transcription units are visualized as an axis of rDNA (r

chromatin) covered with densely packed lateral fibrils forming a gradient of increasing length and resulting in a typical "Christmas tree" pattern. Active transcription units are designated also matrix units by some authors. In the majority of intra- or extrachromosomal rRNA genes studied, several active transcription units, with identical polarity, are arranged in tandem on the rDNA axis and separated by fibril-free spacers. Up to 120-150 matrix units on a single axis have been identified in some cases (cf. Spring et al .. 1976, 1978). Normally the bulk (80-90%) of the active transcription units are of standard size, corresponding roughly to the size of the primary pre-rRNA typical for a given eukaryote. For example, the standard matrix unit in Acetabularia mediterranea is 2.1 JIm, corresponding to a molecular weight for pre-rRNA of about 1.7-2.1 X 106 (Spring et al .. 1976); the matrix unit in Xenopus laevis is 2.2-2.6 JIm, and the molecular weight of its pre-rRNA is in the range of 2.1-2.6 X 106

(Scheer et al .. 1977); in CHO cells, matrix units of 2.6-5.2 JIm correspond to a primary pre-rRNA with a molecular weight of about 4.5 X 106 (Puvion-Dutilleu I et al.. 1977a). Estimates based on the correlation between the molecular weight of primary pre-rRNA and the length of the respective matrix unit show that transcribed rDNA is only slightly shorter than the calculated length for Bform DNA (Miller and Hamkalo, 1972a,b; see below).

The size of spacer intercepts and consequently of rRNA repeating units shows greater variations, not only among different cells and organisms, but even


within the same rDNA axis (see Franke et aI., 1976; Rungger and Crippa, 1977). This observation reflects the length heterogeneity of ntS segments established by other techniques.

The tandem arrangement of rRNA transcription units is not the sole possibility exploited by eukaryotes. Thus, in extrachromosomal rDNA of Physarum polycephalum two matrix units of opposite polarity start from the central part of the linear template molecule (Grainger and Ogle, 1978). A similar arrangement of matrix units is occasionally seen in other eukaryotes (cf. Trendelenburg et al., 1974). An intriguing alternating polarity of rRNA genes along a single rDNA axis was observed recently in Acetabularia exigua (Berger et al., 1978).

Second, the lateral fibrils of matrix units are obviously pre-rRNA chains growing with progress of transcription. Initial studies revealed that the growing pre-rRNA chains are about 5- to 10-fold shorter than expected. This observation, combined with enzymatic and specific staining studies, led to the important conclusion that the growing pre-rRNA chains are already coated with proteins (see O. L. Miller and Hamkalo, 1972a,b). Preliminary studies with specific antibodies indicate that structural ribosomal proteins are also present in the growing pre-rRNP fibrils (Chooi, 1976). A constant length increment of growing pre-rRNP fibrils is observed in some organisms. However, in most cases terminal granules (up to 30-nm diameter) are formed at the free end of the growing fibrils (see Franke et al., 1976; Angelier et al., 1979), thus indicating a tighter RNA-protein packing at the 5'-end of the pre-rRNA chain.

The growing pre-rRNP fibrils are bound to the rDNA axis by granules of about 12-15 nm in diameter considered to be RNA polymerase molecules (see O. L. Miller and Hamkalo, 1972a; Franke et aI., 1976). It appears that rRNA genes are transcribed with a very high efficiency, resulting in a maximal packing of RNA polymerases along the rDNA axis. The packing of RNA polymerases was assessed in several cases. In a broad variety of organisms the observed maximal packing ratios are in the range of 40-50 enzyme molecules/ tLm axis, thus showing that 50-70% of the template is covered. These observations suggest that in fully active rRNA genes spatial hindrance by adjacent RNA polymerase molecules may be an important factor limiting the maximal transcription rate. The observed regular spacing of RNA polymerases along the rDNA axis suggests further that in fully active rRNA genes constant rates of initiation, elongation, and release of pre-rRNP molecules are maintained.

Third, the tandem arrangement of rRNA genes and the observed maximal packing of RNA polymerases poses the question whether each transcription unit has a separate promotor or transcription of several units starts from a single initiation point. The latter possibility was proposed on the basis of detailed quantitative studies on inhibition of rRNA synthesis by actinomycin D (Perry and Kelley, 1970). The observation of granules in ntS segments of the rDNA axis (see Franke et aI., 1976) seemed to support such a possibility. However, detailed

22 A. A. Hadjiolov

analyses provided evidence that in most cases such beaded structure of spacer intercepts results from changes in chromatin conformation rather than from the presence of nontranscribing RNA polymerase molecules (see Franke et al. , 1976, 1978; Scheer, 1978). Studies on the activation or inactivation of prerRNA synthesis in Triturus alpestris oocytes also provided evidence in favor of the existence of independent promotors for each rRNA transcription unit (Scheer et aI., 1975,1976).

Direct proof for the existence of an independent promotor for each rRNA transcription unit was obtained by analysis of the transcription of bacterial plasmids, containing a single Xenopus laevis rRNA gene, injected into oocyte nuclei (Trendelenburg and Gurdon, 1978). In this case (Figure 7) a relatively short ntS segment (about 2.5 knp) of the rRNA repeating unit contained enough information to initiate correctly the transcription of the rRNA gene by oocyte RNA polymerase A and to attain a maximal packing of enzyme molecules and growing pre-rRNP fibrils along the transcription unit axis.

b. Structure of Active and Inactive r-Chromatin. The recent discovery of nucleosome organization of chromatin caused a huge expansion of studies on

A _____ 8 ~ ____ c

FIGURE 7. Transcription patterns of Xenopus laevis rDNA from recombinant plasmid circles (A,S) injected into Xenopus oocyte nuclei and an endogenous rDNA (C) from the same preparation. All three transcription units are identical in size, packing of RNA polymerases, lateral fibril gradients, and their terminal coiling. The ntS segments from rRNA genes (arrows) are predominantly unbeaded. [Reproduced from Trendelenburg and Gurdon (1978) with permission of the authors and Nature (London) .]


the molecular structure of the genome. It is hoped that clues to regulatory mechanisms involved in gene action may be anticipated. The nucleosome organization of inactive rRNA genes, including both transcription units and ntS segments, is now firmly established (Scheer, 1978; Foe, 1978). The "beads-ona-string" image of inactive r-chromatin is apparently the same as in bulk chromatin, suggesting similar structure of nucleosomes. A recent study on the organization of rDNA circles in Dytiscus marginalis oocytes provided a direct approach to the structure of inactive r-chromatin (Scheer and Zentgraf, 1978). It is shown that inactive rDNA organized in nucleosomes displays a packing ratio (,urn B-form DNA/,um chromatin) of 2.2-2.4. Further compaction of the nucleosome chain into supranucleosomal globules, with a diameter of 21-34 nm and a packing ratio of about 11 is also found in inactive rDNA circles. Typical nuclesome organization of inactive r-chromatin, with rDNA packing ratios of 2.0-2.3, is observed also in Drosophila melanogaster (McKnight et al., 1978) and Oncopeltus Jasciatus (Foe, 1978) embryos. Finally, a DNA packing ratio of 2.1 is found in inactive chromatin circles formed after injection of bacterial plasmids, containing rDNA, into oocyte nuclei (Trendelenburg and Gurdon, 1978). These results are in good agreement with the known structure of nucleosomes in bulk chromatin (see Kornberg, 1977; Oudet et al., 1978; Tsanev, 1978). Together with data on the composition of isolated r-chromatin, the above findings indicate that inactive rRNA genes are condensed in nucleosomes of about 200 np rDNA, constituted by a core nucleosome (140 np rDNA plus two sets of histones H2a, H2b, H3, and H4) and a linker rDNA (about 20-30 np plus histone HI). Whether some quantitative or qualitative difference exists in the histone and nonhistone complement of nucleosomes from inactive r-chromatin and bulk chromatin cannot be decided at present.

The DNA in nucleosomes surrounds a histone core and both strands are fixed by strong ionic interactions. Therefore, transcription requires unfolding of nucleosomes. Observations of active rRNA genes clearly show that this is the case with r-chromatin. Measurements of the length of active transcription units yielded an rDNA packing ratio of only 1.1 to 1.4 (cf. McKnight et al., 1978; Foe, 1978; Franke et al., 1976,1978). Such a low rDNA packing ratio requires an almost complete unfolding of the nucleosome (see Weintraub et al., 1976; Tsanev and Petrov, 1976; Richards et al., 1978). Because active rRNA genes are covered by RNA polymerases, the observed shortening of rDNA could, in fact, be caused by local unwinding (0. L. Miller and Hamkalo, 1972b; McKnight et al., 1978). Therefore, we may ask: What is the actual template in active rRNA genes? Is it naked rDNA or r-chromatin? A conclusive answer is not yet possible. However, several observations support the view that transcribed rDNA is still associated with histones and, possibly, nonhistone proteins. These include (1) the thickness of the active axis is 40-70A (rather than the 20A of B-form DNA) and it stains with the basic protein reagent phosphotungstic acid

24 A. A. Hadjiolov

(Foe et al., 1976; Foe, 1978); (2) digestion with micrococcal nuclease of chromatin containing presumably fully active rRNA genes in Tetrahymena pyriformis (Mathys and Gorovsky, 1978), Physarum polycephalum (Johnson et al., 1978a,b; Butler et al., 1978) or Xenopus laevis (Reeves, 1978a,b) yields 200 np rDNA fragments. This indicates that active rDNA is still associated with histones in a spatial arrangement similar to that in bulk chromatin nucleosomes; (3) complete unfolding of bulk chromatin nucleosomes does not result in the removal of core nucleosome histones (see Tsanev, 1978); and (4) the presence of histones H2b and H3 in active nonribosomal DNA axes was demonstrated by immune electron microscopy (McKnight et aI., 1978).

If RNA polymerase A transcribes along a fully unfolded r-chromatin axis, the histones may be expected to remain attached by their highly basic N-terminal segments to the sugar-phosphate backbone of the rDNA chain(s) (Weintraub et aI., 1976; Richards et al., 1978). That transcription of histone-coated DNA is possible is demonstrated by model experiments with T7 phage or calf thymus DNA transcribed by E. coli (Williamson and Felsenfeld, 1978) or Saccharomyces cerevisiae (Karagyozov et aI., 1978) RNA polymerases. It is shown that elongation on native (Williamson and Felsenfeld, 1978) or single-stranded (Karagyozov et aI., 1978) DNA complexed with histones is still possible, although yielding somewhat shorter products. On the other hand, initiation is markedly more affected, suggesting that special structural requirements should be met by chromatin in the promotor region. Summarizing the above studies it may be concluded that r-chromatin, rather than "naked" rDNA, is more likely to be the actual template in active rRNA transcription units.

The mechanisms causing unfolding of nucleosomes and activation of rRNA genes remain totally unknown. However, an important observation in studies on the morphology of active transcription units is that unfolding of rchromatin is not directly connected with transcription. Indeed, fully extended rchromatin is observed at stages preceding or following transcription of rRNA genes (Foe et al., 1976; Laird et aI., 1976; Foe, 1978; Scheer, 1978; Franke et at., 1978). Consequently, unfolding of nucleosomes is unlikely to be induced by RNA polymerase, and the participation of more or less independent control factors (proteins?) should be envisaged (see Wang and Kostraba, 1978).

In conclusion, inactive r-chromatin appears to be tightly packed in nucleosomal and supranucleosomal structures (Figure 8). Activation of rRNA genes seems to proceed in several consecutive stages. The initial stages involve unfolding of supranucleosomal globules (A), followed by unfolding of nucleosomes (B). Fully unfolded r-chromatin is committed for transcription, but still untranscribed (C). At this stage interaction of RNA polymerase A with promotor sites takes place resulting in the switching on of the rRNA gene (D). The overall process is obviously reversible and supplies the cell with efficient mechanisms for transcriptional control of ribosome formation. The molecular mechanisms

Biogenesis of Ribosomes

A

B

c ....

D •• *.'

r DNA/ r-chromotin pac king ratio ............... , •.• ,~,.,. t... ..... -11

I \ I \

/ U \, I \

I , I ,

I , I ...... •• , .. ~

2.2 - 2.4

?

1.1 -1.4 _ ........

25

FIGURE 8. Successive stages in the activation or inactivation of r-chromatin. The dotted lines indicate the changes in the length of r-chromatin from one rRNA transcription unit. The rDNA to r-chromatin packing ratio is approximate, based on data obtained with different cells and organisms.

involved in the activation and inactivation of r-chromatin provide an attractive target for future investigations.

3.2.3. Primary Transcripts and Primary Pre-RNA

As mentioned above, formation of mature rRNA in all eukaroytes starts from a discrete pool of large pre-rRNA molecules containing the sequences of both L-rRNA and S-rRNA. Thus, primary pre-rRNA may be defined as the largest pre-rRNA species accumulating as a distinct pool in the nucleolus. The primary rDNA transcript may be defined as the polynucleotide chain corresponding to the full length of the rRNA transcription unit. It is possible that the primary transcript and the primary pre-rRNA are not identical, i.e., the primary transcript may contain sequences removed before the accumulation of primary pre-rRNA. The possibility that processing takes place before the end of transcription is shown in studies with bacteria. In E. coli separate precursors to L-rRNA and S-rRNA are split normally from the growing rRNA chain, whereas a 30-S pre-rRNA, containing the sequences for both mature rRNA,

26 A. A. Hadjiolov

can be found only in RNase 111- mutants (see Nikolaev et al., 1975; Nomura et al., 1977). Short-term labeling in eukaryotes invariably shows that initially more than 90% of the label is in primary pre-rRNA, without any detectable labeling of S-rRNA (or its immediate precursor) originating from the 5' end of the growing transcript. Therefore, the sequence S-rRNA-tSj-5.8 S-tSj-LrRNA, present in primary pre-rRNA, is not normally processed during transcription. Consequently, if the primary transcript is processed before the formation of primary pre-rRNA, three possibilities may be envisaged: (1) partial removal of tSe sequences from the 5' end of the primary transcript, (2) removal of 3'-end sequences located beyond the L-rRNA segment, and (3) excision of transcribed insertion sequences and splicing of the primary transcript.

Because transcription of insertions in rRNA genes has not yet been observed, I shall discuss here only possibilities (1) and (2).

Introduction of the gene-spreading technique permitted a direct approach to the problem of the identity between primary transcript and primary prerRNA. Since the initial studies (see O. L. Miller and Hamkalo, 1972a), many investigators compared the estimated length of rDNA transcription units and the size of primary pre-rRNA in a variety of cells and organisms. As discussed above, in most cases the size of primary pre-rRNA corresponds to the length of the transcription unit, assuming that rDNA is in the B form. That this is not a mere coincidence is shown by studies with Dytiscus marginalis ooc)'tes, where the contour length of isolated circular rONA coincides with that of actively transcribed circles (Trendelenburg et al., 1976). However, at the molecular level, important limitations exist in evaluating the precise correlation between primary transcripts and primary pre-rRNA. These include (1) the growing transcript is coated with proteins, and direct measurement of its length is impossible; (2) B-form DNA appears to be 1.1- to l.4-fold contracted in the active transcription unit, but the packing ratio cannot be measured precisely; (3) short transcripts (up to 500 nucleotides) may not be visualized, leaving open the possibility that the length of the transcription unit is underestimated; and (4) determination of the molecular weight of primary pre-rRNA is subject to errors of 0.1-0.3 X 106• These limitations leave open the possibility that the primary transcript is, in fact, longer than primary pre-rRNA.

Biochemically, fractionation of short-term labeled RNA reveals in many cases the existence of RNA molecules, apparently larger than the major primary pre-rRNA. Thus rapidly labeled "85 S" and "65 S" RNA fractions were isolated from rat liver nucleoli (Hidvegi et al., 1971) and a "41 S" (3.4 X 106

daltons) RNA was detected in Acer pseudoplatanus L. (Miassod et al., 1973; Cox and Turnock, 1973). In vitro synthesis of RNA molecules larger than the respective primary pre-rRNA was observed in isolated Rana pipiens nuclei (Caston and Jones, 1972) or rat liver nucleoli (Grummt and Lindigkeit, 1973; Grummt et al., 1975).

The existence of RNA molecules larger than primary pre-rRNA does not


imply a precursor-product relationship. It may also reflect heterogeneity among rRNA transcription units. Indeed, studies on the length distribution of transcription units in several organisms revealed size heterogeneity within a given cell, and even within the same rDNA axis (0. L. Miller and Beatty, 1969a; Scheer et at., 1973, 1977; Trendelenburg et at., 1974,1976; Spring et at., 1974, 1976; Berger and Schweiger, 1975a,b; Foe et at., 1976). It seems unlikely that these differences are artifacts inherent in the gene-spreading technique. For example, careful studies with Xenopus taevis oocytes (Scheer et al., 1973, 1977) show that the bulk of rRNA transcription units are homogeneous (2.2-2.6 /-Lm). However, longer transcription units (3.4-4.2 /-Lm) are consistently found.

The established heterogeneity of rRNA matrix units should be correlated with the well-documented homogeneity of rRNA transcription units found in studies using restriction enzymes and heteroduplex mapping (see above). Aside from possible artifacts, two explanations of this apparent contradiction are plausible. First, in some cases RNA polymerases may not initiate or terminate at the correct sites. Alternatively, such sites may be multiple, thereby allowing some flexibility in initiation or termination points. Second, the primary transcript is processed normally during transcription to the size of primary prerRNA. Thus oversize matrix units would reflect their true length and result from defective processing of the primary transcript.

The first possibility is supported by observations showing that the pool of primary pre-rRNA molecules is itself heterogeneous (Loening, 1970, 1975). Three distinct primary pre-rRNA peaks were identified in a mouse cell line (Tiollais et aI., 1971) and it was shown that two of them differ by a piece of about 700 nucleotides (Galibert et al., 1975). Heterogeneity of primary prerRNA was observed also in liver, mainly owing to differences in the tSe segment (Dabeva et aI., 1976). It is important that tracer kinetics experiments with Acer pseudoplatanus L. cells (Cox and Turnock, 1973) or rat liver (Dabeva et at., 1976) failed to show a precursor-product relationship between rapidly labeled "oversize" RNA molecules and primary pre-rRNA. However, it should be stressed that heterogeneity of primary pre-rRNA may be reasonably correlated with electron-microscopic observations, but is not sufficient to decide between the above two possibilities.

The second possibility, implying processing of the primary transcript during transcription, has been discussed in detail by several authors (Loening, 1975; Franke et aI., 1976; Hadjiolov and Nikolaev, 1976; Rungger and Crippa, 1977). The evidence may be summarized as follows:

(1) The length of the bulk of matrix units in some species is markedly larger than expected from the size of the primary pre-rRNA. The most striking example is given by oocytes in Acheta domesticus (5.4 /-Lm and about 15.9 knp B-form DNA) and Dytiscus marginalis (3.6 /-Lm, 10.6 knp B-form DNA), exceeding by far primary pre-rRNA of about 8100 nucleotides (Trendelenburg

28 A. A. Hadjiolov

et al., 1973, 1976; Trendelenburg, 1974). Such a large discrepancy could hardly be explained by overstretching of transcribed rONA.

(2) In several cases RNA polymerases and attached lateral fibrils are seen in ntS segments of the rONA axis. Such transcriptional complexes, showing a length gradient of attached fibrils, are often seen in front of normal matrix units and are named "prelude transcripts" (Scheer et al., 1973; see Franke et al., 1976). They correspond to RNA chains of about 2000 nucleotides and seem to be cleaved before the RNA polymerase starts the transcription of tSe

sequences in primary pre-rRNA. (3) Unusually long lateral fibrils are occasionally seen protruding above

the regular gradient in apparently normal matrix units (see Franke et al., 1976). The possibility that a block of processing of growing pre-rRNP chains may cause visualization of transcription in ntS segments is indicated by studies on the action of 5-fluorouridine on Xenopus laevis oocytes (Rungger and Crippa, 1977; Rungger et al., 1978).

These observations support the concept that processing of the primary transcript occurs simultaneously with transcription. In most cases, processing of 5'-end tSe sequences appears more plausible. Nevertheless, the available evidence is not conclusive, and alternative interpretations are not ruled out (see Loening, 1975; Franke et al., 1976).

Conclusive evidence could be obtained by comparative sequence analysis of 5/- and 3/-end segments of the rRNA transcription unit, its transcript and primary pre-rRNA. Because promotor and terminator sites are not known and the primary transcript cannot be isolated, such studies are not yet feasible. Fortunately, present techniques allow the identification of initial nucleotides in DNA transcripts. It is known that transcription starts with ATP or GTP. Thus, identification of 5'-terminal triphosphates (pppN) in primary pre-rRNA would provide evidence for its identity with the primary rONA transcript and the absence of 5' -end processing.

Initially, studies with animal cells failed to show the presence of pppNp structures in primary pre-rRNA. In Xenopus laevis pGp was found (Slack and Loening, 1974), whereas the 5'-end nucleotides in MH 134 (Kominami and Muramatsu, 1977) and Novikoff (Nazar, 1977) hepatoma cells were heterogeneous. However, more recently evidence for the presence of 5'-terminal pppAp in Xenopus laevis primary pre-rRNA was obtained by showing that it can be a capping enzyme substrate (Reeder et al., 1977). Direct demonstration of 5'-triphosphates was soon achieved in several eukaryotes. Thus pppAp and pppGp in Saccharomyces cerevisiae (Hadjiolov et al., 1978; Nikolaev et al., 1979), pppAp in Tetrahymena pyriformis (Niles, 1978), and pppAp in Drosophila (Levis and Penman, 1978) primary pre-rRNA were identified. The identification of pppAp in a precursor to S-rRNA from Dictyostelium discoideum (Batts-Young and Lodish, 1978) further extends the list.

The identification of 5'-terminal triphosphates in primary pre-rRNA


shows that in eukaryotes processing of 5'-end tSe sequences during transcription does not take place. These results show also that processing of the primary rDNA transcript cannot start normally before the transcription of its L-rRNA segment is completed. Thus, the absence of pre-rRNA processing, simultaneous with transcription, clearly distinguishes eukaryotes from prokaryotes. The above results further suggest that the primary transcript and the primary prerRNA in eukaryotes are identical. Accordingly, the observed heterogeneity of primary pre-rRNA (Loening, 1970, 1975; Tiollais et al., 1971; Dabeva et al., 1976) and its tSe segments (Dabeva et al., 1976) would be explained by the presence of multiple promotor sites rather than by processing during transcription. This possibility is strongly supported by recent evidence, which shows that the promotor sequence of Xenopus laevis rRNA genes is reduplicated and located in the ntS region (Boseley et al., 1979; Moss and Birnstiel, 1979). The absence of processing of 5'-end segments in primary pre-rRNA has its counterpart in 5 S rRNA, where 5'-end triphosphates (pppGp in most cases) were demonstrated in several cases (see Erdmann, 1980).

Whether eukaryotic primary rDNA transcripts contain some 3'-end tSe

sequences (beyond the L-rRNA segment), which are rapidly removed to yield primary pre-rRNA, cannot be answered at present. The available evidence on the structure of rDNA repeating units makes unlikely the possibility that the normal termination signal is too far away from the 3' end of the L-rRNA segment in primary pre-rRNA (see Sollner-Webb and Reeder, 1979). However, some of the morphological evidence, showing the presence of oversize transcripts, may reflect occasional read-through events owing to faulty termination of transcription. This is certainly not the case of the intriguing "prelude transcripts," which could represent the synthesis of some distinct RNA species of unknown function. The unequivocal interpretation of all experimental findings reviewed here is still not possible, but there is little doubt that further analysis of rDNA transcripts and their relation to primary pre-rRNA will contribute greatly to our understanding of the mechanisms of transcription of rRN A genes.

3.3. Transcription in Vitro

The complete understanding of transcription of rRNA genes and its regulation requires in vitro analysis and reconstitution of active transcription complexes. The subject has been reviewed (Jacob, 1973; Chambon, 1975; Biswas et al., 1975), and only some aspects will be briefly considered here.

3.3.1. rRNA Genes

A straightforward approach would be the transcription of isolated rRNA genes by purified RNA polymerase A. Transcription of isolated DNA (includ-

30 A. A. Hadjiolov

ing rONA) by purified RNA polymerase A shows that the enzyme initiates at nicks, ends, or denatured regions of the template acting as pseudopromotors. Thus, such simplified systems have not yet been mastered to yield correct transcription of rRNA genes. In fact, recent studies provided evidence that purification of ONA results in loss of specificity in transcription of rRNA genes (Ballal et aI., 1977; Matsui et al., 1977). Transcription of bacterial plasmids containing Drosophila rRNA genes, with homologous RNA polymerase A is also unspecific (Luse and Beer, 1978). These and other similar results strongly suggest that r-chromatin is the actual template structure transcribed by RNA polymerase A (see above). On the other hand, native chromatin structure may not be a stringent requirement for accurate transcription. At least in the case of yeasts, evidence was obtained for a selective transcription of rONA by purified RNA polymerase A (Holland et aI., 1977), including data for the faithful transcription of complete pre-rRNA molecules (Van Keulen and Retel, 1977). These results indicate that RNA polymerase A (or some associated factor) may playa critical role in the accurate transcription of rRNA genes. Whatever the case, accurate in vitro transcription of isolated r-chromatin (or rONA) by purified RNA polymerase A would require a better knowledge of the native structure of both template and enzyme.

At the present stage, transcription of rRNA genes in isolated nuclei or nucleoli, with endogenous template and enzyme, seems to be a more promising in vitro system. Indeed, numerous studies have shown (see Chambon, 1975; Biswas et aI., 1975) that in such systems, transcription of rRNA genes is taking place. Evidence for selective transcription was obtained by characterization of the product by nucleotide composition, nearest-neighbor frequency, fingerprinting, methylation of nucleotides, and hybridization with isolated rDNA (see Hadjiolov and Milchev, 1974; Udvardy and Seifart, 1976; Onishi and Muramatsu, 1978). However, analysis of the size of the product, attempted in some studies, yielded variable results, thus suggesting that reproducibly faithful transcription is not easily attained. For example, isolated rat liver nucleoli yielded rDNA transcripts larger than the 45 S pre-rRNA made in vivo (Grummt and Lindigkeit, 1973). On the other hand, transcription products with a maximal size of about 45 S were reported recently with isolated nucleoli from liver or other animal cells (Onishi and Muramatsu, 1978; Grummt et al., 1979). Treatment of nucleoli with the detergent Sarkosyl L removes more than 90% of the proteins, but transcription of rRNA genes is still possible, although the fidelity of transcription is apparently altered (Samal et aI., 1978). These and other studies indicate that a better characterization of the experimental systems used will be needed before accurate transcription of rRNA genes could be attained.

One specific point deserves further comment. Transcription of rRN A genes in isolated nuclei or nucleoli seems to involve only elongation of rONA transcripts initiated in vivo (Hadjiolov and Milchev, 1974; Udvardy and Seifart, 1976; Onishi et al., 1977). Thus, initiation of rRNA chains seems to be an


extremely vulnerable process. Whether these observations reflect the need for a continuous supply of cytoplasmicfactor(s) or, more likely, the alteration of transcription complexes during isolation of nuclei or nucleoli, remains to be clarified.

3.3.2. 5 S rRNA Genes

The use of in vitro systems for the faithful synthesis of 5 S rRNA has been more successful. Several studies with isolated nuclei from different animal cells revealed the in vitro formation of discrete low-molecular-weight RNA components identified tentatively as 5 S rRNA and pre-tRNA (Price and Penman, 1972; Marzluff et aI., 1974; Hadjiolov and Milchev, 1974; Weinmann and Roeder, 1974; Udvardy and Seifart, 1976; Weil and Biatti, 1976). In some of these studies, long-term linear incorporation of labeled substrates into a putative 5 S rRNA peak suggested reinitiation in vitro, while inhibition by high concentrations of a-amanitin indicated the involvement of RNA polymerase C (Weinmann and Roeder, 1974; Marzluff et aI., 1974,1975; Udvardy and Seifart, 1976; Wei! and Blatti, 1976). The recent use of plasm ids containing 5 S rDNA permitted a more precise analysis of RNA polymerase C products in isolated nuclei (Yamamoto and Seifart, 1977a). It was conclusively shown that discrete 5 S rRNA molecules are made in vitro having the correct size and sequence. Under optimal conditions (Mg2+ and ionic strength below 50 mM) reinitiation and recycling of RNA polymerase C was taking place. A more detailed analysis (Yamamoto and Seifart, 1977b) showed a slight ambiguity in termination of 5 S rRNA chains, one or two additional residues being present in some molecules. It should be noted that formation of 3' end extended 5 S pre-rRNA molecules was found also in studies in vivo (B. Jacq et aI., 1977). Furthermore, accurate transcription of 5 S rDNA was obtained not only with isolated chromatin with the endogenous enzyme, but also using exogenous RNA polymerase C (Yamamoto et at., 1977), opening the way to a detailed analysis of the control factors involved in the transcription of 5 S rRNA genes.

Along an independent line, injection of purified or cloned 5 S rDNA into Xenopus taevis oocytes also resulted in its faithful transcription, thus supplying another valuable system for the study of in vitro 5 S rRNA synthesis (Brown and Gurdon, 1977, 1978).

The successful transcription of 5 S rRNA genes in in vitro systems achieved recently opens prospects that similar techniques for the transcription of the larger rRNA genes may soon be forthcoming.

4. PROCESSING OF PRIMARY PRE-rRNA AND PRERIBOSOMES

The maturation of rRNA in eukaryotes starts from a more or less homogeneous pool of primary pre-rRNA molecules containing the sequences of S-

32 A. A. Hadjiolov

rRNA, 5.8 S rRNA, and L-rRNA. Therefore, the polynucleotide chain of primary pre-rRNA should be complete and have the "correct" structure in order to enter the sequence of reactions involved in its processing. Moreover, the primary pre-rRNA is, in fact, part of a primary preribosome and its accurate assembly and processing involve important RNA-protein and protein-protein interactions. These features of primary pre-rRNA and preribosomes stress the point that their processing is prepared in many respects during transcription of rRNA genes and should be considered as the second stage of ribosome biogenesis.

4.1. Structure of Primary Pre-rRNA

4.1.1. Size and Nucleotide Composition

The mature rRNA species in eukaryotes are characterized by (1) size variations of L-rRNA in the range of 1.3-1.75 X 106 daltons and (2) size homogeneity of S-rRNA with a molecular mass of approx. 0.7 X 106 daltons (see Loening, 1970; Attardi and Amaldi, 1970). The size of primary pre-rRNA molecules displays a marked increase in evolution, reflecting mainly divergence in both external and internal nonconserved tS sequences (see Figure 6).

The nucleotide composition and primary structure of rRNA species in eukaryotes displays several characteristic features (see Gerbi, 1976; R. A. Cox, 1977): (1) considerable evolutionary divergence exists in both L-rRNA and SrRNA, as shown by nucleotide composition, RNA-DNA hybridization, and fingerprint or partial-sequence analyses; (2) conserved sequences (about 50% in S-rRNA and 30-40% in L-rRNA) seem to include large segments characterized by the presence of modified nucleotides; (3) large (100-500 np) [G + C]-rich segments (up to 78% G + C) are characteristic for L-rRNA in higher eukaryotes (see also Hadjiolov and Nikolaev, 1976); and (4) the 5.8 S rRNA sequences, typical of eukaryotes, are strongly conserved in evolution. They do not reveal any apparent similarity to either prokaryotic or eukaryotic 5 S rRNA (see also Erdmann, 1980).

The nonconserved tS sequences in pre-rRNA species are much less studied, aside from the facts that (1) they do not contain methylated nucleotides or probably pseudouridine (see Maden et al., 1977); and (2) they have a higher [G + C] content, and distinctive large double-stranded loops are found in animal cells. These and other features show that considerable evolutionary changes in tS sequences are typical of eukaryotes.

4.1.2. Modifications

Extensive modification of nucleotides is typical of eukaryotic pre-rRNA. These include (1) methylation of bases or the 2'-OH group of ribose, and (2) conversion of uridine into pseudouridine. These studies were reviewed (Maden


et al., 1974, 1977) and will be considered only briefly here. Most of the methylated nucleosides and pseudouridines are already present in primary prerRNA. Therefore, modification is largely taking place during or shortly after transcription. The respective enzymes should be tightly associated with transcription complexes in nucleoli, but they are poorly characterized mainly because of difficulties in obtaining specific polynucleotide substrates (Liau and Hurlbert, 1975; Liau et al., 1976). Detailed studies with HeLa cells on the oligonucleotides containing methylated nucleosides demonstrated that methylations of L-rRNA and S-rRNA sequences in primary pre-rRNA follow a highly specific pattern (Maden et al., 1974; Maden and Salim, 1974). Also, the number of pseudouridine and 2'-O-methyl-nucleoside residues in LrRNA, S-rRNA, and 5.8 S rRNA, as well as in primary pre-rRNA, is almost equal (see Maden et al., 1977), a striking observation, indicating that these two types of modification of the pre-rRNA chain could be interdependent. In animal cells there are 200 modification sites-lOO for methylation (about 90% 2'-O-methyl groups) and 100 for pseudouridylation. In lower eukaryotes the number of modified nucleotides is markedly lower. Comparative studies with vertebrates showed that the oligonucleotides containing modified nucleotides are more strongly conserved in evolution than is the rest of the polynucleotide chain (Maden et al., 1977; Khan et al., 1978). The highly specific modification pattern of L-rRNA and S-rRNA sequences in primary pre-rRNA suggests that methylation and pseudouridylation play an important role in the assembly and processing of preribosomes, as well as in ribosome structure and function.

4.1.3. Secondary Structure

Until recently, there was no information on the conformation of primary pre-rRNA. Studies on the denaturation spectra of 45 S pre-rRNA from ascites tumor cells showed that its secondary structure is closer to that of the homologous L-rRNA (Hadjiolov and Cox, 1973). The electron-microscopic observation of pre-rRNA molecules, partly denatured in formamide-urea, permitted the visualization of characteristic double-stranded loops in pre-rRNA (Wellauer and Dawid, 1973; see Wellauer and Dawid, 1975). The distinctive location of these loops along the chain of pre-rRNA and rRNA molecules permitted their unmistakable identification. In all vertebrates studied until now such loops (approx. 100-500 np) were observed in the tS., tS j , and L-rRNA segments ofpre-rRNA molecules (Wellauer and Dawid, 1975; Schibler et al., 1975; Dabeva et al., 1976). Taking into consideration their resistance to denaturation, these loops most likely correspond to the high-molecular-weight [G + C)-rich segments identified earlier in animal rRNA (see Hadjiolov and Nikolaev, 1976; R. A. Cox, 1977). Such large [G + C)-rich segments are not found in lower eukaryotes and should be considered as a late acquisition in evolution with yet unknown significance.

34 A. A. Hadjiolov

4.2. Pre-rRNA Maturation Pathways

4.2.1. General Remarks

The conversion of primary pre-rRNA and preribosomes into mature rRNA and ribosomes is a multistep process. Basically, this process takes place in the nucleolus, but some late events occur in the nucleoplasm, and even in the cytoplasm. The processing of pre-rRNA is much better characterized than are the conversions involving the preribosome. Because pre-rRNA and rRNA chains constitute the backbone of the respective ribonucleoprotein particles, the mechanisms of pre-rRNA processing constitute an important aspect of the overall process of ribosome biogenesis. Maturation of pre-rRNA is a relatively fast process, and only minute amounts of primary and intermediate pre-rRNA species are present in the cell under steady-state conditions (Table I). Therefore, the correct analysis of pre-rRNA maturation pathways depends critically on the techniques for the identification and analysis of large pre-rRNA molecules. The introduction of gel-electrophoresis techniques for the separation of RNA permitted the identification of several intermediate pre-rRNA species. Degradation of RNA during the isolation of nuclei or nucleoli, interference by highly labeled hnRNA, formation of aggregates, and so forth, are hazards that require carefully controlled experimental conditions and critical evaluation of the results. Several methods were used to identify separate pre-rRNA com-

Table I Absolute Pool Sizes of pre-rRNA and rRNA Molecules in Rat

Liver Nuclei"

rRNA species (S)

45 41 36 32 21 28 18

10-4 X number of molecules per nucleus

1.2 ± 0.2 [1.0]" 0.9 ± 0.2 1.2 ± 0.2 2.9 ± 0.4 [4.0] 2.5 ± 0.5' 9.6 ± 1.5 [7.0] 8.7 ± 1.7 [6.0]

"The amounts of pre-rRNA and rRNA molecules are determined by A260 measurements of RNA extracted from detergent-purified nuclei and fractionated by gel electrophoresis (Dabeva e/ al., 1978). The sum of all nuclear pre-rRNA and rRNA molecules is about 3.5% of rRNA molecules in the liver cell.

'The values in brackets are for HeLa cells nuclear pre-rRNA and rRNA measured by methods similar to the ones used for liver (Weinberg and Penman, 1968).

'This value is likely to be overestimated owing to interference by RNA species distinct from 21 S pre-rRNA.


ponents: estimation of molecular weights, competition hybridization with total DNA or purified rONA, comparison of methylation patterns or fingerprint analysis of oligonucleotides obtained upon hydrolysis of purified pre-rRNA, electron-microscopic construction of secondary structure maps of pre-rRNA after partial denaturation in formamide-urea and others. The combination of these techniques allowed the unequivocal identification of primary and intermediate pre-rRNA species in a variety of cells and organisms. All these methods permit the outline of possible pre-rRNA maturation pathways leading to mature rRNA. The actual precursor-product relationships require detailed labeling kinetics and pulse-chase studies. Because of the very rapid transformations involved and numerous methodical imperfections, only in a limited number of cases have the actual pre-rRNA maturation pathways been thoroughly documented. Consequently, the tentative character of the pre-rRNA maturation pathways proposed for different eukaryotes has to be kept in mind.

4.2.2. Common Features

Studies on the structure and topology of primary pre-rRNA show that a common pattern is shared by all eukaryotes, suggesting that basically similar mechanisms operate in pre-rRNA maturation. The available information on pre-rRNA maturation pathways in different cells and organisms was reviewed (Hadjiolov and Nikolaev, 1976) and will not be considered in detail here. Welldocumented schemes of pre-rRNA maturation are available for primitive unicellular eukaryotes (Saccharomyces, Tetrahymena pyriformis), plants (Phaseolus vulgaris, Acer pseudoplatanus), insects (Drosophila melanogaster), lower vertebrates (Xenopus laevis), and several mammalian cells and organisms (HeLa and L cells, rat liver). A basic similarity emerging from these studies is that discrete intermediate pre-rRNA species can be identified. Therefore, primary pre-rRNA processing has to involve at least primarily a succession of endonuclease attacks generating intermediate pre-rRNA and mature rRNA species (see Winicov and Perry, 1975; Perry, 1976; Hadjiolov and Nikolaev, 1976). Whether some further trimming of pre-rRNA and rRNA molecules by specific exo- or endonucleases is also needed still remains an open question (see below). In any case, it seems likely that normally the structure of pre-rRNA and the preribosome specify a few "marker sites" in the polynucleotide chain defining its selective cleavage by endonuclease(s). The endonuclease attacks may follow a rigid sequential pattern so that each processing step induces conformational changes that trigger the next one. As a result, a single pre-rRNA maturation pathway is predominant for a given cell type (see Perry, 1976; Hadjiolov and Nikolaev, 1976). Taking into consideration the available data, a general processing scheme shared by all eukaryotes may be outlined (Figure 9).

36 A. A. Hadjiolov

CD 5.8S rRNA tSe \.is-rRNA t Sj"'/ L rRNA

5' CI ::::::jl.-=~ltI· ••• _ 3'

S-rRNA L-rRNA

FIGURE 9. General pattern of primary pre-rRNA maturation in eukaryotes. The major endonuclease attack sites are numbered 1-4. tS" external transcribed spacer; tS" internal transcribed spacer; S-rRNA, 5.8 S rRNA, and L-rRNA, sequences corresponding to the respective mature rRNA.

The proposed general scheme is, of course, tentative. Several of its features deserve further comment. The first step resulting in the removal of a rather large 5'-end tSe segment is well documented in a variety of vertebrates (Wellauer and Dawid, 1975; Schibler et aI., 1975; Dabeva et at., 1976) and in HeLa cells the tSe segment split by endonuclease attack was identified by its characteristic secondary structure (Wellauer and Dawid, 1973). The existence of such tSe sequences and their initial removal in other eukaryotes are suggested by several observations. For example, the size of the 37 S primary prerRNA in Saccharomyces cerevisiae is markedly larger than the sum of the intermediate precursors to L-rRNA and S-rRNA (Dudov et aI., 1976; Nikolaev et aI., 1979). Correlations with data on the yeast rRNA repeating unit indicate the existence of a 5'-end tSe segment of about 0.3 X 106 daltons. Furthermore, the existence of a small 5'-end tSe segment (about 0.2 X 106 daltons) was also inferred from studies with Drosophila melanogaster cells and the respective pre-rRNA (containing both L-rRNA and S-rRNA segments), obtained after its removal was identified (Levis and Penman, 1978). Whether the removal of 5' -end tSe segments is, indeed, a compulsory first step in prerRNA processing remains to be established with certainty. It should also be stressed that in several cases the identification of the expected intermediate pre-rRNA is difficult. Furthermore, in Dictyostelium discoideum, the presence


of 5'-end pppAp was shown in an intermediate S-rRNA precursor (BattsYoung and Lodish, 1978). These data indicate that an endonuclease cut at site 2 or 3 may, in fact, precede the cut at site 1 (see Figure 9).

The second step in pre-rRNA processing results in the formation of intermediate pre-rRNA for both L-rRNA and S-rRNA. The formation of a precursor to L-rRNA has been proved in all studies on pre-rRNA processing in eukaryotes (see Hadjiolov and Nikolaev, 1976). The formation of an intermediate precursor to S-rRNA is found in several eukaryotes, the best documented cases including Saccharomyces (Udem and Warner, 1972; Trapman and Planta, 1975), Tetrahymena pyriformis (Pousada et al., 1975), Musca domestica (Hall and Cummings, 1975, 1977), mouse and rat liver (Egawa et al., 1971, Fujisawa et al., 1973; Hadjiolov et al., 1974a; Dabeva et aI., 1976), and HeLa cells (Weinberg and Penman, 1970; Maden et al., 1972; Wellauer and Dawid, 1973). However, in a broad variety of eukaryotes including Drosophila melanogaster (Levis and Penman, 1978), Xenopus laevis (Loening et al., 1969; Wellauer and Dawid, 1974), and mouse L cells (Wellauer et al., 1974a) a precursor to S-rRNA could not be detected. Instead, several intermediate precursors to L-rRNA were identified, thereby suggesting that in these organisms the endonuclease split at site 3 is, in fact, preceding the one at site 2. Such a switch in the sequence of endonuclease attacks results in a presumably faster output of mature S-rRNA, but its physiological significance remains obscure.

The last steps (endonuclease cuts at sites 3 and 4) result in the formation of mature rRNA and possibly mature ribosomes (see below). The formation of L-rRNA is of particular interest. In all eukaryotes studied until now, the existence of a relatively large pool of an immediate precursor to L-rRNA in the nucleolus has been ascertained. Conclusive evidence was obtained that this pre-rRNA contains the sequences for both L-rRNA and 5.8 S rRNA (Maden and Robertson, 1974; Nazar et al., 1975). Therefore, the formation of LrRNA, hydrogen bonded with 5.8 S rRNA, is obviously a rather complicated process. It involves at least three endonuclease cuts shaping the ends of both L-rRNA and 5.8 S rRNA. The sequence and molecular mechanisms are still unknown. The identification of a 7 S precursor to 5.8 S rRNA in Saccharomyces carlsbergensis (Trapman et aI., 1975a) suggests that its formation and further processing may be an important intermediate step. It is likely that sequence analysis of the respective pre-rRNA and L-rRNA will provide more detailed information on the formation of 5.8 S rRNA and its interaction with L-rRNA. The last step leading to mature S-rRNA (site 3) is less understood. The nucleolar pool of the immediate precursor to S-rRNA may be rather low in some cells. Moreover, in some lower eukaryotes, Saccharomyces cerevisiae (Udem and Warner, 1973) and Tetrahymena pyriformis (Pousada et aI., 1975), this pre-rRNA is rapidly transferred in the cytoplasm, where its conversion into S-rRNA takes place.

38 A. A. Hadjiolov

4.2.3. Multiplicity

Until recently it was generally agreed that a single pre-rRNA maturation pathway is typical for a given cell type. Labeling kinetics studies with yeast cells (Udem and Warner, 1972; Trapman and Planta, 1975) provided evidence in support of this concept, at least for the case of primitive eukaryotes.

The possibility that processing of pre-rRNA may take place along parallel pathways was raised by observations showing the simultaneous existence of several "minor" pre-rRNA components. Thus "aberrant" intermediate prerRNAs were observed in HeLa cells, in particular upon viral infection (Weinberg and Penman, 1970). Also, comparison of pre-rRNA species in mouse L cells (Wellauer et at., 1974a) and mouse liver (Hadjiolov et at., 1974a) suggested the predominance of distinct pathways in the two types of mouse cells, most likely caused by an inverted sequence of endonuclease attacks at sites 2 and 3 in the 41 S pre-rRNA molecule (see Winicov, 1976; Figure 9). Studies with human lymphocytes showed that in resting cells, both 36 Sand 32 S prerRNA are formed. Phytohemagglutinin stimulation caused a fourfold increase in the rate of processing only through 32 S pre-rRNA (Purtell and Anthony, 1975).

Further evidence favoring the simultaneous operation of two pre-rRNA processing pathways was obtained with the mutant BHK cell line 422E showing a temperature-sensitive lesion in the conversion of 32 S pre-rRNA into LrRNA (Toniolo et at., 1973). It was found that at the permissive temperature these cells generate L-rRNA from 32 S pre-rRNA and S-rRNA from 20 S pre-rRNA. In contrast, at the restrictive temperature, the temporal order of endonuclease attacks at site 2 and site 3 was altered generating 36 S prerRNA and S-rRNA (Winicov, 1976; Toniolo and Basilico, 1976). The simultaneous existence of 36 S, 32 S, and 21 S pre-rRNA in rat liver further supports the versatility in nuclease attacks at sites 2 and 3 and the simultaneous processing of pre-rRNA along at least two parallel pathways (Dabeva et at., 1976).

These observations raise the question about the actual precursor-product relationships in pre-rRNA maturation. There is little doubt that the structure of primary pre-rRNA, and possibly the preribosome, specify a fewendonuclease attack sites along the polynucleotide chain (see Perry, 1976). The established flexibility in the sequence of endonuclease attacks prompts the analysis of pre-rRNA maturation as a stochastic process, i.e., a process whereby attack at each site in a given pre-rRNA is considered possible. Such an analysis was attempted with rat liver, a population of cells operating under steady-state conditions (Dudov et aI., 1978). Detailed tracer kinetics experiments on the turnover of pre-rRNA and rRNA were carried out. An algebraic approach to computer analysis of the relations among a large number of interconnected pools


was developed, and different models were analyzed, postulating the possibility of simultaneous cleavage of each pre-rRNA at its cleavage sites. This analysis permitted the selection of a most-probable model for existing connections among pre-rRNA and rRNA pools (Figure 10). The following main features may be deduced: (1) considerable flexibility exists in the sequence of endonuclease attacks at the early stages of pre-rRNA processing, resulting in the simultaneous occurrence of several processing pathways; and (2) the cleavage sites involved in the formation of mature rRNA (L-rRNA in particular) are protected until the generation of their immediate pre-rRNA, thereby specifying a terminal checkpoint before the release of mature ribosomes.

It is not yet clear to what extent the multiplicity of pre-rRNA maturation pathways in rat liver is typical of other eukaryotes. It seems plausible that channeling of pre-rRNA along alternative pathways is connected with the overall rate of ribosome biogenesis in the cell, a more rigid sequence of endonuclease attacks being typical of rapidly growing cells. In any case, the observed multiplicity of pre-rRNA maturation pathways seems to be characteristic for animal cells, whereas a more rigid pattern exists in lower eu karyotes. For example, Saccharomyces cerevisiae temperature-sensitive ts-

0.32 0.33

45 S

1 ~l

5'1 ! or 3'

1 1 1 1 2 3 4

0.81

0.39

41 S

5'.tl=-__ 3' 11 1 2 3 4

FIGURE 10. Most probable model of pre-rRNA processing in rat liver (left) and probabilities of endonuclease cleavage (right) at the separate attack sites in 45 Sand 41 S pre-rRNA numbered 1-4. Endonuclease attacks of 45 S pre-rRNA generate: site 1. 41 S pre-rRNA; site 2. 36 S pre-rRNA + unidentified precursor to 18 S rRNA; site 3. 32 S pre-rRNA + unidentified precursor to 21 S pre-rRNA and 18 S rRNA; site 4. "28 S" rRNA + 39 S RNA. The dotted arrows indicate conversions occurring with very low transfer rates. [Based on data from Dudov et al. (1978).]

40 A. A. Hadjiolov

mutants, defective in pre-rRNA maturation, failed to show a switching on of alternative processing pathways at the nonpermissive temperature (Andrew et aI., 1976; Gorenstein and Warner, 1977; Venkov and Vasileva, 1979). Whether a more flexible sequence of endonuclease attacks in animal cells confers better opportunities for regulation of ribosome biogenesis remains to be elucidated.

4.2.4. Enzyme Mechanisms

It is now evident that endonucleolytic hydrolysis of pre-rRNA phosphodiester bonds is extremely specific. If we consider the pre-rRNA processing scheme (see Figure 9), it is clear that the accurate endonuclease cleavage of only six (out of 10,000-15,000) phosphodiester bonds would be sufficient to ensure the generation of mature S-rRNA, 5.8 S rRNA, and L-rRNA. This concept may be oversimplified, but there is little doubt that the generation and shaping of mature rRNA proceeds with a very high precision. The enzyme mechanisms are still obscure. We do not know whether the substrate is prerRNA or the respective preribosome. We also do not know what structures in the substrate determine a specific cleavage site. Hence we can envisage different possibilities ranging from strictly specific nucleases for each cleavage site to fully nonspecific nucleases acting on cleavage sites specified by the unique structure of pre-rRNA molecules and their complexes with proteins.

Analysis of terminal nucleotides (and sequences) in eukaryotic pre-rRNA and rRNA may provide some information on the enzymes involved in their formation. The obtained results with a broad variety of eukaryotic cells and organisms have been tabulated (Winicov and Perry, 1975; Hadjiolov and Nikolaev, 1976; R. A. Cox, 1977). It is noteworthy that the terminal nucleotides in mature rRNA species appear to be strongly conserved in evolution, thereby suggesting that the nuclease(s) involved in their formation have a specific requirement toward the nucleotides forming their substrate phosphodiester bond (Figure 11).

Studies with more than 10 different eukaryotes show, without exception, that a split of a ... N~U ... bond (site A) forms the 5' end of S-rRNA. The 3' end of S-rRNA is also strongly conserved and results in all cases from the cleavage (site B) of a ... A~N ... bond. In fact, a much longer sequence [ ... AUCAUUAoH] at the 3' end of S-rRNA is invariably found in all eukaryotes studied until now (see Cox, 1977). The second nucleotide in site B has been identified only in Saccharomyces cerevisiae (Skryabin et al., 1979) yielding ... A~A ... as the susceptible phosphodiester bond.

The 5' end of mature L-rRNA is generated by a ... N~U ... cleavage (site C) in lower eukaryotes, whereas attack at ... N~C ... appears to be typical for higher eukaryotes (Sakuma et al., 1976; see Cox, 1977).

The terminal nucleotides (and, in fact, the complete sequence) of several

Biogenesis of Ribosomes

® ~

..... NpU .....

41

@ © t N~U

..... ApN .... NpC •.... • •.•.. UOH

5,8 S rRNA

FIGURE 11. Phosphodiester bonds hydrolyzed by the nucleases shaping the 5' and 3' ends of eukaryotic S-rRNA, L-rRNA, and 5.8 S rRNA. The arrows indicate the sites of nuclease attack.

eukaryotic 5.8 S rRNA are now known (see Erdmann, 1980). With the excep-tion of Saccharomyces having pA ... at the 5' end, all 5.8 S rRNA result from the cleavage (site D) of a ... N~C ... bond. Sequencing studies on the Xen-opus laevis precursor to 5.8 S rRNA show that a ... G~C ... bond is split to yield its 5' end (Ford and Mathieson, 1978; Boseley et al., 1978). The 3' end of 5.8 S rRNA is also conserved, resulting from a cleavage (site E) of a ... U~N ... ( ... C~N ... in Xenopus laevis) bond. In the case of Saccharomyces carlsbergensis this bond is further characterized as ... U~C ... (Dejonge et al., 1978), whereas it is ... C~G ... in Xenopus laevis (Boseley et al., 1978).

The results summarized here indicate that (1) the phosphodiester bond involved in generating the ends of mature rRNA species seems to be of the type ... NPU ... or ... Nbc . .. , where N is likely to be a purine nucleotide; and (2) involvement of a single type of endonuclease (5'-ribonucleotide hydrolase) may be sufficient to generate all mature rRNA species.

The above possibilities are, of course, tentative. Their substantiation requires further studies on the characterization of nucleases involved in prerRNA processing. Whether endonuclease attacks alone would be sufficient to generate directly mature rRNA is also an open question. For example, evidence was obtained favoring a terminal 5'-end processing of L-rRNA in the cytoplasm of hepatoma cells (Kominami et al., 1978). In any case, it is obvious that the nature of the phosphodiester bond at a given cleavage site cannot, in itself, determine the high specificity of nuclease attack typical of pre-rRNA processing.

In fact, we do not know yet whether the entire structure of pre-rRNA alone is sufficient to define a given nuclease cleavage site. The possibility that the structure of pre-rRNA is sufficient to specify its accurate processing is raised by studies with E. coli. Here, the early steps of 30 S pre-rRNA processing are catalyzed by ribonuclease III, an enzyme specific for double-

42 A. A. Hadjiolov

stranded RNA (see Nikolaev et al., 1975). The possible role of doublestranded regions in pre-rRNA in specifying endonuclease cleavage sites is supported by several studies. Hydrolysis of 45 S pre-rRNA from HeLa cells with purified E. coli RNase III yielded several discrete RNA fragments, some of which showed an electrophoretic mobility similar to that of pre-rRNA species found in vivo (Gotoh et al., 1974; Torelli et al., 1977). Inhibition of normal pre-rRNA processing by some intercalating agents (Snyder et aI., 1971; Waltschewa et al .. 1976) also supports the possible involvement of doublestrand specific endonucleases. Such endonucleases were isolated from nuclei or nucleoli of animal cells, and evidence was obtained that they generate intermediate pre-rRNA similar to those found in vivo (Saha and Schlessinger, 1978; Grummt et al .• 1979). Several authors have isolated endonucleases from the nuclei or nucleoli of animal cells splitting primary pre-rRNA (see Winicov and Perry, 1975; Hadjiolov and Nikolaev, 1976). It is noteworthy that some of these endonucleases, i.e., isolated from nucleoli of L cells (Winicov and Perry, 1974, 1975) or Novikoff hepatoma (Prestayko et al .. 1972, 1973) show some preference for uridylate or cytidylate residues, a finding in support of their participation in pre-rRNA processing.

All the above findings indicate the existence in nucleoli of specific endonuclease(s) whose requirements toward the primary and secondary structure of pre-rRNA indicate their role in processing. However, it is likely that the definitive specification of the endonuclease cleavage sites is determined by the conformation of the entire preribosome. In vitro processing of preribosomes (containing 45 S pre-rRNA) with partially purified endonucleases was reported for HeLa cells (Mirault and Scherrer, 1972; K wan et al.. 1974), mouse L cells (Winicov and Perry, 1974), and Novikoff hepatoma (Prestayko et al .. 1973). In all cases partial maturation of the preribosome was observed, which appeared to be closer to the in vivo process than was observed with purified pre-rRNA as substrate.

4.3. Pre ribosomes: Structure and Processing

All pre-rRNAs in the nucleolus are associated with proteins to form preribosomes. Two types of such particles were identified initially in HeLa nuclei: "80 S" containing 45 S pre-rRNA, and "55 S" containing 32 S pre-rRNA (Warner and Soeiro, 1967). Both particles contain also 5 S rRNA. Analogous types of particles were soon identified and characterized to a different extent in studies with Saccharomyces carlsbergensis (Trapman et al .. 1975b), Tetrahymena pyriformis (Leick, 1969), amphibian oocytes (Rogers, 1968), mouse L cells (Liau and Perry, 1969), rat liver (Narayan and Birnsteil, 1969), Novikoff hepatoma (Prestayko et al.. 1972), and several other cells and organisms.

It is now firmly established that the "80 S" preribosome is a precursor of


the "55 S" preribosome, which in turn matures to produce the large ribosome (see Warner, 1974; Craig, 1974). In most studies, separate particles containing other intermediate pre-rRNA could not be identified. However, in mouse L cells, a distinct particle, containing 36 S pre-rRNA was found (Liau and Perry, 1969), whereas in HeLa cells a discrete particle carrying 41 S pre-rRNA was reported (Mirault and Scherrer, 1971). It appears that the "80 S" and "55 S" particles are, in fact, processing particles, containing small amounts of other intermediate pre-rRNA, aside from the major component. It is remarkable that particles containing precursors to S-rRNA usually are not identified in the nucleolus, probably because of leakage during isolation or because of a rapid transfer to the nucleoplasm and cytoplasm. However, mild isolation of HeLa cell nucleoli permitted the identification of a small pool of "40 S" particles, containing 20 S pre-rRNA (Mirault and Scherrer, 1971). In yeasts and other lower eukaryotes, the last maturation step yielding S-rRNA takes place in the cytoplasm, where the respective preribosome has been identified (Udem and Warner, 1973; Trapman et al., 1975b; Pousada et al., 1975). A rapidly labeled ribonucleoprotein particle, sedimenting at about 30 S, was isolated from nucleoli of different animal cells. This particle contained 45 S pre-rRNA and labeled faster than the 45 S pre-rRNA in "80 S" particles, thus suggesting that it may be an earlier-stage preribosome (Bachellerie et al., 1971, 1975). All in all, taking into account the difficulties in the isolation of pure ribonucleoprotein particles, only the "80 S" and "55 S" preribosomes are now characterized to some extent.

The protein complement of preribosomes was studied in several laboratories. The proteins of "80 S" preribosomes are still poorly characterized. These preribosomes constitute only 10-20% of nucleolar preribosomes, and their isolation is obviously more difficult. Studies by 2 D gel electrophoresis of the proteins from total nucleolar ribonucleoproteins from Novikoff hepatoma (Prestayko et al., 1974) or HeLa cells (Phillips and McConkey, 1976) showed that 10-15 S-proteins (out of 30 in cytoplasmic small ribosomes) are presumably associated with "80 S" preribosomes. Direct analysis of the proteins from isolated "80 S" preribosomes of HeLa (Kuter and Rodgers, 1976) or mouse leukemia (Auger-Buendia and Longuet, 1978; Auger-Buendia et al., 1978) cells were published recently. In the study with HeLa cells, S-proteins could not be identified. However, in leukemia cells, the authors identified 31 L-proteins (out of 40), 15 S-proteins (out of 31), and 7 nonribosomal proteins. Their results strongly suggest that although "80 S" preribosomes contain several structural ribosomal proteins, still a large number (S-proteins in particular) are added at later stages of ribosome biogenesis.

The proteins of "55 S" preribosomes have been studied in more detail, including analysis of isolated particles from HeLa cells (Kumar and Warner, 1972; Kumar and Subramanian, 1975; Kuter and Rodgers, 1976), regenerat-

44 A. A. Hadjiolov

ing rat liver (Tsurugi et ai., 1973), mouse leukemia cells (Auger-Buendia and Longuet, 1978), and Saccharomyces carlsbergesis (Kruiswijk et ai., 1978). Important observations were made, which may be summarized as follows:

(1) Several nonribosomal proteins are associated with preribosomes. Their number varies from 10 to 30 in separate studies, and it appears that some are joined during the conversion of "80 S" into "55 S" preribosomes. These proteins turn over more slowly than do ribosomal proteins. They do not leave the nucleolus and are apparently reutilized in preribosome assembly and maturation (see Warner, 1974).

(2) The "55 S" preribosome contains about 30 L-proteins. It is remarkable that the L-proteins in "80 S" and "55 S" particles are practically the same, a single protein being added at this stage (Auger-Buendia and Longuet, 1978).

(3) In contrast, about 10 L-proteins are added at the last maturation step leading to mature large ribosomes. About 16 S-proteins are also added after a putative precursor to the small ribosome is released from the "80 S" preribosome. These considerable changes in the protein complement of ribosomes during the last stages of maturation correlate well with the established more stringent control of pre-rRNA processing at this stage (Dudov et al., 1978).

The bulk of the available evidence demonstrates that during maturation of preribosomes the structural ribosomal proteins are added in a sequential pattern, a large number of proteins joining the preribosomes at late stages. Moreover, some ribosomal proteins seem to enter the ribosome particles after their exit into the cytoplasm. Thus three Saccharomyces cerevisiae L-proteins exchanged with the large ribosome in the absence of new ribosome formation (Warner and Udem, 1972). More recently, evidence for the recycling in the cytoplasm of a few L- and S-proteins was obtained in studies with HeLa (Kumar and Subramanian, 1975; Lastick and McConkey, 1976; Aiello et ai., 1977) and Drosophila (E. Berger, 1977) cells.

In summary, the emerging sequential pattern of involvement of structural ribosomal proteins in the assembly of pre ribosomes and ribosomes opens the way for a deeper understanding of ribosome biogenesis. Elucidation of the role of individual ribosomal and nonribosomal proteins in the assembly and maturation of preribosomes appears to be a promising approach in future studies.

5. REGULA nON

5.1. General Considerations

Ribosomes are needed for protein synthesis, and the control of ribosome biogenesis has to respond to requirements for protein production. Formation of polyribosomes by the interaction of short-lived mRNAs with an excess of stable


ribosomes is clearly not the case in eukaryotes. In fact, continuous formation of ribosomes is taking place throughout the life cycle of eukaryotic cells. Even in resting cells ribosomes are involved in incessant turnover (Loeb et ai., 1965; Hadjiolov, 1966). There are several cases wherein ribosomes display a faster response to external stimuli than is true of mRNA. For example, growth stimulation of resting fibroblasts resulted in a more than twofold increase in protein synthesis and formation of new ribosomes, whereas synthesis of mRNA remained unchanged (Rudland et ai., 1975). This and other similar results indicate that ribosome biogenesis includes mechanismf> that permit rapid changes in its intensity.

The experimental evidence available at present demonstrates that ribosome biogenesis is a complex multistep process. The following major stages in ribosome biogenesis may be outlined:

1. Transcription of rRNA genes to produce primary pre-rRNA, containing S-rRNA, 5.8 S rRNA, and L-rRNA sequences. Transcription of 5 S rRNA genes.

2. Synthesis of ribosomal proteins, involving transcription of r-protein genes, maturation of r-protein pre-mRNA, and translation of r-protein mRNA.

3. Chemical modifications of the primary rDNA transcript. Interaction with ribosomal and nonribosomal proteins to build the primary preribosome. Association of 5 S rRNA with the preribosome.

4. Cleavage of the primary pre ribosome into precursors to the large and small ribosome.

5. Processing of the two preribosomal particles to mature large and small ribosomes, including chemical modification (generation of 5.8 S rRNA; conversion of pre-rRNA into rRNA; "late" methylations of rRNA) plus addition and removal of proteins.

6. Nucleolus - nucleoplasm and nucleoplasm - cytoplasm transfer of large and small ribosomes including some late maturation changes.

7. Involvement of the large and small ribosomes in the ribosome polyribosome cycle and protein synthesis.

8. Degradation of the large and small ribosome.

The precise molecular mechanisms of most of these stages are not yet understood. The problem becomes more complicated when we consider that the major control mechanisms may vary in different eukaryotic cells and organisms or under different physiological or pathological conditions. Different aspects of the regulation of ribosome biogenesis were considered in earlier reviews (Maden, 1971; Craig, 1974; Warner, 1974; Perry, 1976; Hadjiolov and Nikolaev, 1976). Here, I shall consider only some aspects of the problem, where important new information was obtained.

46 A. A. Hadjiolov

5.2. Transcriptional Control

The information on the basic mechanisms of transcription of rRNA genes was considered. Here I shall discuss some aspects of the regulatory mechanisms determining the rate of primary pre-rRNA production. Regulation by changes in the total number of rRNA genes, achieved by their differential replication was already considered (Section 2.1.2; see also Craig, 1974; Spear, 1974; Tartof, 1975). This regulation seems to operate only in some cells (oocytes) and organisms to meet unusually high demands for ribosome production. Usually, the number of rRNA genes is typical for a given cell and, at least in most somatic cells, it remains constant throughout the life cycle (see Birnsteil et al., 1971). In principle, regulation of pre-rRNA synthesis may operate by two mechanisms, changing (1) the number of active rRNA genes, and (2) the rate of transcription of active rRNA genes. These will be considered in turn.

5.2.1. Number of Active rRNA Genes

The total number of rRNA genes in a broad variety of eukaryotic cells and organisms is known (see Birnstiel et aI., 1971). However, evaluation of the number of active rRNA genes is not easily attained. Several studies supply evidence that, normally, only part of the rRNA genes are active. For instance, Drosophila melanogaster lines with a varying total number of rRNA genes produced identical amounts of rRNA (Mohan and Ritossa, 1970). Shortage of rRNA genes was felt only when they were reduced below a critical number, as found in mutants with alterations in the "bobbed" gene mapping in the nucleolus organizer region. In this case the flies grow more slowly, but still have the same number of ribosomes as found in wild-type individuals (Mohan and Ritossa, 1970). Similar findings were reported in studies with various Xenopus laevis mutants (see Reeder, 1974). Numbering of active genes after spreading is possible in principle. This was attempted in a detailed study with Acetabularia mediterranea (Spring et al., 1978). The results showed that, at fully active stages, there are 3500-4800 active rRNA genes per cell, a figure corresponding to a total number of about 3800 genes. Thus, at least in lower eukaryotes, switching on of all rRNA genes is attained in some stages of their life cycle. The gene-spreading technique was also used to study the changes in the number of active rRNA genes during early embryonic stages in Drosophila melanogaster (McKnight and Miller, 1976; McKnight et al., 1978) and Oncopeltusfasciatus (Foe et al., 1976; Foe, 1978) and at different stages of spermiogenesis in Drosophila hydei (Meyer and Hennig, 1974) and of oogenesis in Triturus alpestris (Scheer et al., 1976; Franke et al., 1978). All these studies provided conclusive evidence that on-off switching of rRNA genes is a basic mechanism in the regulation of ribosome biogenesis. As discussed earlier, these


investigations also showed that it is most likely that (1) activation of rRNA genes precedes their actual transcription, and (2) separate rRNA genes may be independently switched on and off. The precise mechanisms of activation are still unknown, but it is clear that nuclear or cytoplasmic factors controlling the uncoiling of r-chromatin are likely to playa crucial role in regulating the number of active rRNA genes.

The number of active rRNA genes in mature somatic cells has still not been evaluated. The role of rRNA gene activation or inactivation, under different physiological or pathological conditions, thus remains a challenge for future studies. That such mechanisms operate as well in somatic cells is indicated by phenomena such as the repression of some nucleolar organizers by "nucleolar dominance" (see Reeder, 1974) or, more specifically, by the observed inhibition of human rRNA synthesis in mouse-human heterokaryons (Eliceiri and Green, 1969; D. A. Miller et al., 1976).

The identification of putative control proteins, nucleic acids, or other compounds involved in restriction of rRNA genes is at an early stage (see Wang and Kostraba, 1978). Several factors, often unexpected, could be involved, examples being given by the reported activation of transcription by ornithine decarboxylase (Manen and Russell, I 977a,b ) or the identification of smallmolecular-weight RNA, covalently bound to DNA, and likely to participate in maintaining tertiary chromosome structure (Pederson and Bhorjee, 1979).

5.2.2. Control of the Rate of Transcription

Active rRNA genes may be transcribed at different rates, depending on putative regulatory factors acting on initiation, elongation, or termination of primary pre-rRNA chains. The following major factors may be envisaged: (1) amount and activity of RNA polymerase A; (2) concentration of substrate nucleoside-5'-triphosphates; (3) conformation of the growing ribonucleoprotein chain; (4) participation of specific factors controlling initiation, elongation, or termination; and (5) steric hindrance in the nucleolus by accumulation of preribosomes. The rate-limiting role of these factors is not yet clear. It should also be kept in mind that their relative importance may vary under different experimental conditions.

Analysis of the results obtained by the gene-spreading technique (see above) demonstrates the prevalence of a general pattern in the activation or inactivation of rRNA genes, i.e., most "open" genes are fully loaded with RNA polymerases and growing fibrils. This fact suggests that normally the rate of transcription for most rRNA genes is not limited by RNA polymerase A, substrates, initiation, or termination control factors. Therefore, at least in these cases, the overall rate of transcription is governed by the number of active rRNA genes. This fact is further supported by studies on the transcription of

48 A. A. Hadjiolov

rRNA genes injected, at more than 100-fold excess, into Xenopus laevis oocyte nuclei (Trendelenburg and Gurdon, 1978; Trendelenburg et al., 1978). The observation of fully loaded transcription units (see Figure 7) shows that once "open" the rRNA gene is transcribed at a maximal rate.

A possible rate-limiting role for some of the above factors may be inferred from electron microscopic studies. Thus, in OncopeltusJasciatus, a denser packing of lateral fibrils at the distal end of some rare genes suggested possible temporary block in transcript release (Foe, 1978). Alternatively, increasing lengths of fibril gradients, upon activation of Drosophila melanogaster rRNA genes, were interpreted as showing that termination control does not determine the packaging of transcribing RNA polymerases (McKnight and Miller, 1976). More often, a lower density of growing fibrils was found in part of the active genes. This is best exemplified in cases of inactivation of Triturus alpestris oocytes (Scheer et al., 1975, 1976), suggesting the participation of initiation control factors in lowering the overall rate of transcription.

Several lines of evidence converge to show that in animal cells RNA polymerase A molecules are in excess over the number required for the estimated transcription rates of rRNA genes. The number of RNA polymerase A molecules per diploid cell is in the range of 2-4 X 104 (Chambon et at., 1972; Cochet-Meilhac et aI., 1974; Cox, 1976; Coupar et aI., 1978). Assuming a polynucleotide chain elongation rate in vivo of 40-80 nucleotides/sec (see Maden, 1971) and a chain length for primary pre-rRNA in the range of 1.25-1.35 X 104 nucleotides (Dabeva et aI., 1976), the cellular polymerases would be able to produce about 3.5-15 X 103 primary pre-rRNA molecules/min and

Type of cell or tissue

Erythroid cells Fibroblasts

Resting Growing

Myoblasts Myofibers L-cel\s HeLa cells Rat liver

Table II Rates of Ribosome Synthesis in Animal Cells·

10-3 X ribosomes/ min/cell

0.22

0.5 1.2 1.2 0.63 4.5 3.1 l.l

Reference

Hunt (1976) Emerson (1971)

Bowman and Emerson (1977) Bowman and Emerson (1977) Brandhorst and McConkey (1974) Wolf and Schlessinger (1977) Dudov et al. (1978)

"Data obtained in experiments measuring the rate of synthesis of ribosomes (or primary pre-rRNA) are included. It is assumed that degradation of newly synthesized pre-rRNA or rRNA is not taking place. The calculations are based on a molecular weight for 28 S + 18 S rRNA = 2.4 X 10'.


presumably the same number of ribosomes. Yet estimated rates of ribosome formation in animal cells are markedly lower (Table II). That RNA polymerase A is not limiting the rate of transcription of rRNA genes is shown also by the finding of active RNA polymerases in anucleolate Xenopus iaevis mutants, where rRNA synthesis is absent (Roeder et ai., 1970). Also, in studies on oogenesis in Amphibia, the practically constant enzyme activity did not correlate with drastic changes in transcription (Roeder, 1974; Hollinger and Smith,1976).

The metabolic stability of RNA polymerase A is also an important factor to be considered. A rapid turnover of nucleolar RNA polymerase was deduced from studies with cycloheximide block of protein synthesis in rat liver (Yu and Feigelson, 1972). However, complete block of transcription of nucleoplasmic genes by a-amanitin did not alter nuclear RNA polymerase A for several hours (Tata et ai., 1972; Hadjiolov et a/., 1974a). Furthermore, the total RNA polymerase A activity in nuclei was not affected by cycloheximide (Benecke et ai., 1973; Schmid and Sekeris, 1973; Onishi et ai., 1977). These and other studies with different eukaryotes (Shields and Tata, 1976; Hildebrandt and Sauer, 1976) permit the conclusion that RNA polymerase A is metabolically stable. Therefore, alterations in the amount of enzyme molecules are unlikely to participate in short-term transcription control.

Formation of ribosomes in eukaryotes is under stringent control by the continuous supply of proteins (see Warner, 1974; Hadjiolov and Nikolaev, 1976). With some, but not all (see below), experimental systems it was found that shortage of proteins results in rapid alterations in the rates of transcription of rRNA genes. Considering the stability of RNA polymerase A, other factors are likely to be involved. In search of such factors, an important observation was the finding that RNA polymerase A exists in two forms-free and chromatin bound (Lampert and Feigelson, 1974; Yu, 1974, 1975; Chesterton et ai., 1975; Matsui et ai., 1976). A regulatory role for putative control factor(s) involved in the association of RNA polymerase A with chromatin was proposed (Yu, 1976), and evidence was obtained that a reduced binding of the enzyme to chromatin may be an early response to cycloheximide block of protein synthesis (Chesterton et ai., 1975; Onishi et ai., 1977). A rapid release of bound RNA polymerase A upon amino acid deprivation or cycloheximide block of protein synthesis and a respective decrease of initiation of transcription was also deduced from studies with yeasts and ascites tumor cells (Gross and Pogo, 1976a,b; Grummt et ai., 1976). The identity of the presumed rapidly turning over protein factor remains to be established among available (see Wang and Kostraba 1978) or still unknown candidates. In a detailed study on liver RNA polymerase A in fed and fasted rats it was concluded that the elongation rate of growing pre-rRNA chains is more tightly coupled to protein synthesis (Coupar et ai., 1978). Thus the site of action of short-lived protein factors remains

50 A. A. Hadjiolov

to be specified. It is noteworthy that the observed maximal packing of RNA polymerases in active rRNA genes suggests that initiation is not rate limiting. Rather, once the gene is "open," the rate of elongation appears to be the major factor controlling overall transcription rates.

Modification of the RNA polymerase A molecule may also affect its activity. Phosphorylation of the enzyme seems the best documented. An inhibition of phosphorylation (or adenylation) of RNA polymerases was proposed to explain their inhibition by the Bacillus thuringiensis exotoxin (Smuckler and Hadjiolov, 1972), later shown to inhibit adenyl ate cyclase (Grahame-Smith et aI., 1975). Phosphorylation of RNA polymerase A by a cyclic AMP-dependent protein kinase was carefully documented (Jungmann et al., 1974; Hirsch and Martelo, 1976; Bell et al., 1976; Buhler et al., 1976). Whether phosphorylation or other possible modifications of RNA polymerase A play some role in the rapid response of the enzyme to changes in protein synthesis remains unknown.

Finally, it should be stressed that in isolated nuclei or nucleoli initiation is blocked, whereas the elongation rate is about IOO-fold lower than it is in vivo (Cox, 1976; Coupar et al., 1978). The loss or rapid exhaustion of initiation or elongation control factor(s), or both, constantly supplied from the cytoplasm, may thus be envisaged.

In most studies, the role of the conformation of growing pre-rRNA chains in modulating transcription rates has not been envisaged. Yet the tight packing of active transcription units in the nucleolus indicates that distortions in the conformation of growing fibrils may influence their elongation rate. Studies with rat liver or He La nuclei demonstrate that added ribosomal proteins are rapidly taken up in the nucleolus (Roth et al., 1976) and stimulate the synthesis of "45 S" pre-rRNA (Bolla et aI., 1977). Because the pool size of free ribosomal proteins in the nucleus seems to be very low (Phillips and McConkey, 1976) the participation of structural ribosomal proteins in the control of transcription rates offers an attractive alternative possibility.

Transcription of 5 S rRNA genes does not appear to be coordinated with that of rRNA genes (Reeder, 1974). Inhibition of pre-rRNA synthesis and processing by the exotoxin of Bacillus thuringiensis (Mackedonski et al., 1972) or 5-fluoroorotate (Hadjiolova et al., 1973) did not alter 5 S rRNA synthesis, although its nucleocytoplasmic flow was halted. Inhibition of mRNA synthesis by a-amanitin (Hadjiolov et al., 1974a) and of protein synthesis by cycloheximide (Hayashi et al., 1977) also did not alter appreciably 5 S rRNA synthesis. Experiments with bobbed Drosophila melanogaster (Weinmann, 1972) and anucleolate Xenopus laevis (L. Miller, 1973) mutants supplied independent evidence for the absence of stringent coordination in the transcription of rRNA and 5 S rRNA genes.

In summary, the mechanisms that control the transcription of rRNA genes play an important role in ribosome biogenesis. Changes in the number


of active rRNA genes seem to constitute a basic response of the cell to variations in the demand for new ribosomes. The switching on and off of rRNA genes is certainly involved in long-term adaptations of the cell as in embryogenesis. Whether such mechanisms operate in more flexible adjustments to changes in translation efficiency is not known. The rate of transcription of active rRNA genes is usually more stable. The amount of RNA polymerase A and nucleotides does not appear to be rate limiting. In some cases, rapidly turning over initiation or elongation factors, or both, may be critical in establishing a stringent control over transcription rates. The loss of such factors could explain the fact that transcription rates in vitro are about lOO-fold lower than in vivo. The correct conformation of the growing, protein-coated, pre-rRNA chain deserves closer attention as a potential factor modulating transcription. Transcription of 5 S rRNA genes is independent and more stable. The supply of 5 S rRNA does not seem to limit either transcription of rRNA genes or ribosome biogenesis.

5.3. Posttranscriptional Control

The large number of studies showing the importance of posttranscriptiona I control mechanisms has been reviewed and different models discussed in detail (Perry, 1973; Warner et al., 1973; Warner, 1974; Hadjiolov and Nikolaev, 1976). Here, I shall consider some aspects of the problem related to the elucidation of critical control sites in ribosome biogenesis.

5.3.1. The Role of Protein Synthesis

The formation of ribosomes is dependent on continuous protein synthesis. This fact, first established with HeLa (Warner et al., 1966) and L (Ennis, 1966) cells, was confirmed with all eukaryotes tested (see Hadjiolov and Nikolaev, 1976). The possibility that the stringent control of ribosome biogenesis operates at the level of transcription was considered. Unexpectedly, in most cases transcription of rRNA genes remained initially unaltered, whereas posttranscriptional mechanisms displayed a faster and deeper response to limited supply of proteins (see Hadjiolov and Nikolaev, 1976). For example, ribosome formation in Saccharomyces cerevisiae is blocked immediately after cycloheximide inhibition of protein synthesis, while synthesis of pre-rRNA continues (DeKloet, 1966; Udem and Warner, 1972). A similar response is obtained with various animal cells upon inhibition of protein synthesis with puromycin (Soeiro et al., 1968), low doses of cycloheximide (MandaI, 1969; Rizzo and Webb, 1972; Farber and Farmar, 1973; Goldblatt et al., 1975; Stoyanova and Hadjiolov, 1979) and by amino acid deprivation (Maden et al., 1969;

52 A. A. Hadjiolov

Vaughan, 1972; Chesterton et al., 1975). What proteins may be involved in the observed stringent posttranscriptional control?

Structural ribosomal proteins join the preribosome in the process of its maturation and are obvious candidates for such a role. The pools of most rproteins in the cell are very low (Wool and Stoffier, 1976; Phillips and McConkey, 1976) and their continuous formation is compulsory for ribosome biogenesis. It is known also that r-proteins are produced by cytoplasmic polyribosomes (Craig and Perry, 1971; Heady and McConkey, 1971) and evidence was obtained that free, rather than membrane-bound polyribosomes are involved (Nabeshima et al., 1975). Studies with yeasts (Mager and Planta, 1976), ascites tumor cells (Hackett et al., 1978), and rat liver (Nabeshima et al., 1979) showed that the mRNAs coding for r-proteins are monocistronic and their translation takes place in small polyribosomes. The mRNAs for r-proteins are relatively stable (Craig and Perry, 1971; Craig et al., 1971; Maisel and McConkey, 1971), but indirect evidence with regenerating rat liver shows that increased transcription of rRNA genes is correlated with enhanced synthesis of mRNAs for r-proteins (B. C. Wu et al., 1977). The information about individual r-protein mRNAs is still limited. A recent study showed that the synthesis of at least 34 S- and L-proteins continues after 75% inhibition of hnRNA synthesis by actinomycin D (Warner, 1977). The newly synthesized r-proteins are rapidly transferred to the nucleus (R. S. Wu and Warner, 1971; Maisel and McConkey, 1971; see Busch et aI., 1978) where they associate with pre-rRNA during its transcription and processing. In fact, r-proteins are taken up and concentrated more than 50-fold in the nucleus and the nucleolus and this involves not only pre-rRNA but probably other nucleolar structures as well (Warner, 1979). The observation that nucleolar and nuclear matrix proteins are distinct (Todorov and Hadjiolov, 1979) suggests that the nucleolar matrix may play an active role. It is also remarkable that, when not involved in ribosome structure, r-proteins are unstable and are rapidly degraded (Warner, 1977).

Summarizing all the above findings it may be safely concluded that the continuous synthesis of r-proteins and their supply to the nucleolus is a major factor involved in the stringent posttranscriptional control of ribosome biogenesis. Whether the supply of some nonribosomal proteins is also rate limiting remains to be established.

5.3.2. The Role of the Pre-rRNA Structure

Pre-rRNA has a unique primary structure and conformation, and its rRNA sequences are extensively modified. This fact raises the question about the role of pre-rRNA structure in ribosome assembly and processing. Progress has been disappointingly slow. There is no positive evidence about the role of


pseudouridylation. Accurate methylation seems to playa rate-limiting role in ribosome formation and enhances the degradation of undermethylated prerRNA (Vaughan et al.. 1967; Liau et al.. 1976; Wolf and Schlessinger, 1977). However, undermethylation of pre-rRNA, induced by cycloleucine, did not prevent the formation of mature ribosomes and their involvement in polyribosomes (Caboche and Bachellerie, 1977). To what extent methylation is critical for pre-rRNA processing in vivo still remains an open question. In any case, it seems that pre-rRNA methylases are metabolically stable and thus unlikely to participate in stringent control mechanisms. Block of protein synthesis by puromycin or cycloheximide did not alter the methylation pattern of pre-rRNA (Tamaoki and Lane, 1968; Shulman et al .• 1977).

Along an independent line, the effect of various nucleoside analogues, incorporated into pre-rRNA chains was studied. Generally, transcription of rRNA genes remains unaltered, while formation of ribosomes is blocked. Incorporation of analogues, like toyocamycin (Tavitian et al.. 1968, 1969) or 5-fluoroorotate (Cihak and Pitot, 1970; Wilkinson et al.. 1971; Hadjiolova et al.. 1973; Hadjiolov et al.. 1974b; Alam and Shires, 1977) did not inhibit the synthesis of pre-mRNA and its processing to mRNA, or the formation of tRNA and 5 S rRNA, whereas ribosome biogenesis was rapidly stopped. Studies with 5-fluoroorotate in mice liver (Hadjiolova et al .. 1973) or 5-fluorouridine in Novikoff hepatoma (Wilkinson et al .. 1975) demonstrated that the block occurs at the last nucleolar stages of pre-rRNA maturation. A similar block at the last steps of ribosome formation was observed in studies on the action of toyocamycin in Saccharomyces cerevisiae (Venkov et al.. 1977), Novikoff hepatoma (Weiss and Pitot, 1974) and mouse leukemia (AugerBuendia et al .. 1978) cells. The synthesis of proteins and the assembly of primary preribosomes was not appreciably altered either by 5-fluorouridine (E. Berger, 1977) or by toyocamycin (Auger-Buendia et al .. 1978). However, with both analogues, accumulation of 36 S pre-rRNA was observed (Hadjiolov and Hadjiolova, 1979; Auger-Buendia et al .. 1978), thus suggesting that alterations in the conformation of preribosomes may induce the channeling of pre-rRNA along alternative processing pathways and the block in ribosome formation.

These and many similar findings (see Hadjiolov and Nikolaev, 1976) strongly suggest that a correct structure of primary pre-rRNA is critical for the accurate assembly and processing of preribosomes.

5.3.3. Critical Control Sites

The importance of posttranscriptional control in ribosome biogenesis is now firmly established. However, the available information about the sites and mechanisms of this posttranscriptional control is still scarce and often controversial. The main reason seems to be the necessity for extensive quantitative

54 A. A. Hadjiolov

and detailed tracer kinetics studies on the nuclear and cytoplasmic conversions of preribosomes and ribosomes. Such studies are already very difficult with steady-state cell systems, whereas even greater difficulties are encountered with unbalanced populations of cells in culture or in the whole animal. The separate stages in ribosome biogenesis were outlined (see Section 5.1). Here I shall try to discriminate those among them which seem to be usually critical in adapting the overall process to the needs of the cell. It is hoped that the elucidation of such critical control sites will help to unravel the underlying molecular regulatory mechanisms.

a. Turnover of Ribosomes. This is the last stage in ribosome biogenesis and it may be a major factor involved in its regulation. The turnover of ribosomes was first observed in adult liver (Loeb et al., 1965; Hadjiolov, 1966; Hirsch and Hiatt, 1966), but it is characteristic of all resting animal cells and tissues (Hogan and Korner, 1968; Boyadjiev and Hadjiolov, 1968; Emerson, 1971; Weber, 1971; Abelson et al., 1974; Kolodny, 1975; Nissen-Meyer and Eikhom, 1976a,b; Scott, 1977; Bowman and Emerson, 1977; Melvin and Keir, 1978). In the case of rat liver, operating under steady-state conditions, both ribosomal particles turn over at essentially the same rate (Hadjiolov, 1966; Tsurugi et al., 1974; Eliceiri, 1976). It is plausible that turnover of ribosomes plays an important role in the control of ribosome biogenesis. This possibility is supported by observations showing that in fasting or protein-deprived animals the turnover rate of ribosomes is markedly increased, half-life values in the range of 50-120 hours being reported under different feeding conditions (Hirsch and Hiatt, 1966; Nordgren and Stenram, 1972; Gaetani et at., 1977). Enhanced cytoplasmic degradation of ribosomes may serve as a signal for changes in transcriptional mechanisms (i.e., inhibition of RNA polymerase A; Coupar et at., 1978) or posttranscriptional mechanisms controlling ribosome production. It is known that decreased protein synthesis results in the accumulation of single ribosomes. Accordingly, several authors have proposed that the pool of single ribosomes supplies signals for both their degradation and for deceleration of ribosome biogenesis (Rizzo and Webb, 1968; Henshaw et al., 1973; Perry, 1973).

A similar situation exists in cultured animal cells. Intensive turnover of ribosomes is switched on in resting cells (Abelson et al., 1974; Bowman and Emerson, 1977). Interestingly, in some cases, the turnover rate of the large ribosome appears to be markedly higher than that of the small particle (Abelson et al., 1974; Kolodny, 1975; Nissen-Mayer and Eikhom, 1976b). Although the significance of this dichotomy in the fate of the two ribosomes is not yet clear, there is little doubt that the turnover of ribosomes may be considered as an important critical control site in ribosome biogenesis, operating in nongrowing eukaryotic cells.


b. Intranuclear Degradation ("Wastage") of Preribosomes and Ribosomes. Several experimental findings converge to show that the continuous supply of r-proteins is a major factor adapting ribosome production to the efficiency of cytoplasmic protein synthesis. This control is very fast and versatile and operates largely posttranscriptionally. In many cases, shortage of rproteins still allows processing of primary pre-rRNA (and preribosomes), but formation of mature ribosomes is halted almost immediately. Thus, a key critical control site in ribosome biogenesis operates at the last maturation step leading to mature ribosomes.

The strongly decreased rates of formation of mature ribosomes, although the synthesis of primary pre-rRNA remains unchanged, may be caused by (1) degradation of excess preribosomes or ribosomes, and/or (2) decreased rates of processing of preribosomes to mature ribosomes.

In a series of studies with resting and phytohemagglutinin-stimulated lymphocytes, Cooper (Cooper, 1969, 1970, 1973; Cooper and Gibson, 1971) provided evidence that upon limited protein synthesis, "wastage" of excess rRNA (largely 18 S rRNA) takes place in the nucleus. This conclusion was based largely on the fact that the ratio of labeled 18 S to 28 S rRNA in the cell was markedly lower than the expected equimolar values. As this relationship was attributable mainly to cytoplasmic rRNA, it was concluded that "wastage" of excess S-rRNA takes place in the nucleus. To preserve the equimolar 18 S to 28 S rRNA ratio, "wastage" of excess L-rRNA in the cytoplasm was postulated to take place at a later stage. The possibility of a posttranscriptional degradation of excess rRNA was considered also as a regulatory mechanism in resting fibroblasts (Abelson et al., 1974) and differentiating myoblasts (Clissold and Cole, 1973; Bowman and Emerson, 1977).

Although a likely phenomenon, further evidence for the extent, intracellular location and even the existence of "wastage" of rRNA is needed before its role in ribosome biogenesis can be evaluated. For example, unequal labeling of L-rRNA and S-rRNA may reflect the known polarity in the labeling of primary pre-rRNA during transcription (see Hadjiolov, 1967). Accordingly, at the very low transcription rates in some resting cells, a lower labeling of SrRNA (located at the 5' end of primary pre-rRNA) is expected (Emerson, 1971). Also, the pools of large and small ribosomes in the cytoplasm of cultured cells may not be equimolar (Nissen-Meyer and Eikhom, 1976a,b), and the observed excess of small ribosomes could explain the apparently slower turnover of this particle.

Channeling of preribosomes along alternative pre-rRNA processing pathways provides another possibility to explain the stringent control over ribosome production caused by shortage of proteins. As shown recently, inhibition of rat liver protein synthesis by low doses of cycloheximide induces rapid alterations

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in pre-rRNA processing (Stoyanova and Hadjiolov, 1979). Soon after inhibition of protein synthesis (Figure 12) the production of 18 S rRNA is abolished and that of 28 S rRNA reduced to one-half the level in controls. This dichotomy in the production of the two ribosomes is correlated with a block in the formation of 41 Sand 21 S pre-rRNA. Generation of 36 Sand 32 S pre-rRNA is still possible, but formation of 28 S rRNA is decreased. A similar channeling of pre-rRNA along alternative processing pathways was also observed in resting and phytohemagglutinin-stimulated lymphocytes (Purtell and Anthony, 1975).

The above observations indicate that the cell may have the capacity to accumulate some unfinished preribosomes pending the supply of critical r-proteins. It is possible that under these conditions the large ribosome displays better survival capability, thereby creating a dichotomy in the production of the two ribosomes. Further limitations in protein supply would switch on degradation of defective preribosomes and eventually slow down transcription of rRNA genes (see also Warner, 1974).

c. Migration of Ribosomes. Another critical site in ribosome biogenesis may be related to the control over the flow of ribosomes along the pathway nucleolus -+ nucleoplasm -+ cytoplasm. Analysis of compartmentation of ribosomes in the cell meets with considerable technical difficulties. However, quantitative estimates in hepatocytes (Table III) show that the concentration of ribosomes in the nucleolus is about 30-fold higher than in the nucleoplasm (Hadjiolov et al., 1978). Therefore, a gradient-driven release of ribosomes from the nucleolus is expected. That this may not be the case is suggested by findings showing that the release of ribosomes from the nucleolus is related to contin-

Table III Amount and Concentration of Ribosomes in Different

Compartments of the Hepatocyte"

V Ribosomes Ribosomes per Compartment (I'm') (N) I'm' X 10-'

Nucleus 280 9.2 X 10' 0.33 Nucleoli 4.6 3.2 X 10' 7.00 Nucleoplasm 280 6.0 X 10' 0.21

Cytoplasm 4000 7.0 X 106 1.75 Free ribosomes 4000 3.5 X 10' 0.09 Monosomes 4000 7.0 X 10' 0.18 Polysomes 4000 6.0 X 106 1.50

"The figures are based on estimates of the amount of 28 Sand 18 S rRNA and are expressed on a per-cell basis (HadjioloY et al .• 1978; Dabeva et al., 1978). V, volume of compartment; N. number of ribosomes.

58 A. A. Hadjiolov

uous transcription of rRNA genes (see Hadjiolov and Nikolaev, 1976). For example, inhibition of transcription with camptothecin did not alter processing of primary pre-rRNA, but blocked the release of L-rRNA from the nucleus (Kumar and Wu, 1973). This effect could be due to alterations in nucleolar chromatin structure caused by the drug. When transcription was blocked by D-galactosamine-induced depletion of UTP, processing of primary pre-rRNA and nucleocytoplasmic transfer of both S-rRNA and L-rRNA proceeded apparently unhampered at least at the initial stages of drug action (Gajdardjieva et al., 1977). Therefore, maturation of preribosomes and release of mature ribosomes from the nucleolus is not always coupled with transcription of rRNA genes. However, particles containing L-rRNA, but defective in their protein complement, seem to be retained in the nucleolus (Soeiro et al., 1968; Willems et aI., 1969; Lonn and Edstrom, 1977b). The role of additional factors, like flow-through capacity of nuclear pores or cytoplasmic compartmentation of ribosomes may be rather complex and their importance in posttranscriptional control mechanisms deserves closer attention. In any case, the restricted mobility of the large ribosome in the cytoplasm (Edstrom and Lonn, 1976; Lonn and Estrom, 1976,1977 a) is a good example of possible posttranscriptional controls related to compartmentation phenomena.

In summary, posttranscriptional control mechanisms play an important role in causing the immediate response of ribosome biogenesis to changes in protein synthesis. A major driving force in this control seems to be the continuous supply of r-proteins to the nucleus. The last step leading to formation of mature ribosomes in the nucleolus appears to be under most stringent control. Temporary storage and degradation ("wastage") of defective preribosomes and ribosomes in the nucleus modulate ribosome production, but the extent and molecular mechanisms of these phenomena remain obscure. Turnover of ribosomes in nongrowing cells plays a major control role in maintaining adequate levels of ribosomes in the cell.

6. CONCLUDING REMARKS

It is clear from this review that in the last few years considerable progress in our understanding of ribosome biogenesis in eukaryotes has been achieved. The molecular mechanisms of an ever-increasing number of aspects of the whole process may now be outlined. They reveal a common pattern of sequential transformations of ribonucleoprotein structures leading to mature ribosomes and thus ensuring the machinery for gene expression.

Ribosome biogenesis begins with transcription of rRNA genes. The organization and structure of the multiple rRNA repeating units seems to guarantee an excess of rRNA genes in the cell and their precise location in the


genome. It is now clear that switching on and off of rRNA genes is a major mechanism controlling ribosome formation during the life cycle of the cell. What are the factors that trigger the opening of new rRNA genes remains an intriguing problem. It is apparent that a better understanding of active and inactive r-chromatin and its interactions with the highly specialized RNA polymerase A will provide attractive targets for future research. In this respect, the likely possibility that ribosome formation cannot begin before transcription of pre-rRNA chains (modified and complexed with ribosomal and nonribosomal proteins) is completed provides important clues to our understanding of transcriptional regulatory mechanisms.

Once formed, the primary preribosome is involved in a complex sequence of maturation steps. The major part takes place in the nucleolus and their elucidation will certainly contribute to understand the rapid physiological and pathological responses of this organelle. The established sequential addition of r-proteins stresses the importance of studies on the role of individual proteins in the maturation of preribosomes. The very low pool sizes of r-proteins make their continuous supply to the nucleolus a major factor in posttranscriptional controls of ribosome biogenesis. Flexibility in the basically rigid sequential pattern of nuclease attacks provides possibilities for degradation or storage of unfinished preribosomes. How the cell achieves the operation of its final checkpoint before only ribosomes with the correct structure are released remains a challenge for future studies. The observed dichotomy in the output of large and small ribosomes is also intriguing. There is little doubt that the fine adaptation of ribosome production to efficiency of protein synthesis is under stringent posttranscriptional control, but the factors and mechanisms involved are still unknown.

The amount of ribosomes in the cell is adapted to the needs of protein synthesis. In most higher eukaryotic cells, operating under more or less steadystate conditions, the switching on of ribosome turnover seems to play an important regulatory role. The molecular mechanisms of ribosome turnover are still unknown and their role in cell function and growth remains to be clarified. In any case, the phenomena of nuclear and cytoplasmic compartmentation of ribosomes seem to be more important than generally realized.

In broader terms ribosome biogenesis offers one of the best understood models of gene expression in eukaryotes. It is now evident that eukaryotes are basically distinct from prokaryotes in their gene expression mechanisms. Important posttranscriptional, but pretranslational mechanisms have evolved in evolution. It seems appropriate to delimit these mechanisms as modulation of genetic information (Hadjiolov and Nikolaev, 1976). The scheme in Figure 13 illustrates the fact that the phenomena of processing of pre-RN A molecules (Perry, 1976) occupy a central position in gene expression in eukaryotes. There is little doubt now that the phenomena of gene splitting and splicing of pre-

60 A. A. Hadjiolov

/ pre-t RNA ---~. tANA 0 om;" ooid,

DNA •

~ pre-mRNA ---+t mRNA ~ Protein

pre-rRNA ---•• 'RNA~ (preribosome) (ribosome)

ITRANSCRI PTION I I MODULATION I ITRANSLATIO N I FIGURE 13. Scheme of the basic stages of gene expression in eukaryotes.

mRNA molecules playa fundamental genetic role. Considered from this angle, ribosome biogenesis is a suggestive example of modulation. It shows that the gene transcript (modified by posttranscriptional enzyme reactions) associates with specific proteins to form unique ribonucleoprotein structures. Conformation of preribosomes is thus a major factor specifying not only the sequential pattern of their maturation, but allows the inclusion of important posttranscriptional regulatory mechanisms. Elucidation of the role of similar phenomena in the modulation of pre-mRNA will certainly constitute a leading trend in the future development of molecular biology. It is likely that further studies on ribosome biogenesis will provide an explanation of the molecular structure and function of the nucleolus. It is hoped that studies on the modulation of premRNA will help us understand the role of the nucleus in eukaryotic cells.

ACKNOWLEDGMENTS

The author is indebted to his colleagues Dr. K. V. Hadjiolova, Dr. K. Dudov, and Dr. L. B. Dolapchiev for their help in preparing the manuscript.

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78 A. A. Hadjiolov

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Chapter 2

The Role of Ribonucleic Acids in the Organization and Functioning of Ribosomes of E. coli

A. A. Bogdanov, A. M. Kopylov, and I. N. Shatsky A. N. Belozersky Laboratory of Molecular Biology and Bioorganic Chemistry Moscow State University Moscow 117234, U.S.S.R.

1. INTRODUCTION

The study of ribosomal ribonucleic acids (rRNA) has considerably intensified in recent years. The reason for this is not only that rRNA is the major structural component of ribosomal particles, comprising almost two-thirds of their mass and determining the location of the proteins in the ribosome, but also that there is growing evidence for the participation of RNA in the organization of the functional centers of the ribosome. There is scarcely any doubt now that rRNAs are directly involved in the interactions with mRNA, tRNA, and protein-synthesis factors. It has long been believed that these ribosomal centers are composed predominantly of protein subunits and, as a consequence, structural studies on ribosomes have for many years centered on the protein components. As a result, we have a vast number of data on the primary and macromolecular structure of the ribosomal proteins and on their localization in the ribosomal subparticles. But this abundance of data cannot be used immediately for elucidating the structural organization of ribosomes and the mechanism of their functioning, because, until very recently, our knowledge of the structure of RNA in ribosomes has been extremely poor.

Our ideas about the structure of rRNA (and all single-stranded RNAs for that matter) have been greatly influenced by the work performed in the laboratories of P. Doty and A. S. Spirin in the late 1950s and early 1960s

81

82 A. A. Bogdanov et al.

(Doty et al., 1959; Spirin et a/., 1959; see also Spirin, 1964 for a review on this earlier work). In terms of the Doty-Spirin model, rRNA is built of rather short double-stranded fragments in which double helices are formed as the result of complementary interactions between neighboring RNA segments and are connected by short single-stranded portions of the polynucleotide chain. The existence of a large number of short helical regions enables the RNA molecule to acquire an ordered and, under certain conditions, rather rigid structure. The structural parameters of the double-stranded fragments of RNA were determined by the fundamental work of Arnott et al. (1968).

Although we do not know the mode of packing of the double-stranded fragments in rRNA macromolecules, the brilliant work of Rich, Klug, and their collaborators (Kim et al., 1974; Robertus et al., 1974), who determined the three-dimensional structure of tRNA, has contributed a host of ideas on the organization of the tertiary structure of single-stranded RNA.

The discovery of the extraordinary conformational lability of high-molecular-weight rRNAs and their ability to respond to changes in ionic strength by altering their structure from being completely unfolded to rather compact, was another important outcome of the early studies. This property is the consequence of the poly electrolytic nature of RNA on the one hand, and of the peculiarities of their secondary structure on the other (Spirin, 1964).

It is noteworthy that the ordered character of certain structural elements in rRNAs as well as their conformational lability are retained in rRNA of ribosomes and are prerequisites of one of their most important properties, i.e., the compactness of the structure in combination with conformational mobility, which is displayed both under changing external conditions and, apparently, in the functioning of the ribosome. Moreover, we shall try to demonstrate here that the formation of the ordered and compact structure of rRNAs in ribosomes occurs because of their ability to undergo conformational transitions, and conversely, that the existence of the superbly organized tertiary structure of RNAs in ribosomes determines their ability to undergo conformational rearrangements.

This chapter analyzes our current views on the macromolecular structure of rRNA in ribosomes, on the intraribosomal interactions that involve rRNA, and on the role of these interactions in the structure and functioning of the ribosomal subunits.

Several reviews on the structure and function of ribosomes have been published recently (Kurland, 1977a,b; Brimacombe et al., 1978; Brimacombe, 1978). Therefore, we shall concentrate on the questions that have not been described adequately in these works. We have also been somewhat self-indulgent in choosing from the enormous number of reported data those that are closer to our own interests.

Structure and Functions of Ribosomal RNA

2. THE SECONDARY STRUCTURES OF RIBOSOMAL RNA IN RIBOSOMES

83

It is known that the 16 S RNA, which comprises the RNA component of the small subunit in E. coli ribosomes, consists of about 1540 nucleotide residues, whereas the large ribosomal subunit contains the 23 S RNA molecule, which is twice as long as the 16 S RNA, and a low-molecular-weight RNA, the 5 S RNA. The total primary structure of the 5 S RNA from E. coli ribosomes (Brownlee et ai., 1967), as well as the total nucleotide sequences of many 5 S RNAs from other organisms, have been known for some time (reviewed by Erdmann, 1976). The nucleotide sequence of 16 S RNA has been studied in Ebel's laboratory (reviewed by Fellner, 1974), and this long-term work has recently been completed (Carbon et aI., 1978). At the same time, the total primary structure of the 16 S RNA molecule was deduced from the complete nucleotide sequence of its gene, which has been determined by Noller and his collaborators (Brosius et ai., 1978). That the sequences derived in the two independent studies are practically identical testifies to their reliability.

The nucleotide sequences of rather long segments of the 23 S RNA from E. coli ribosomes have been determined by classic RNA sequencing methods (Branlant et ai., 1976), and the complete sequence for this RNA containing 2904 nucleotides has recently been obtained by DNA sequencing of the corresponding gene (J. Brosius, T. Dull, and H. Noller, personal communication).

The hypothetical models of the 16 Sand 23 S secondary structures published up to now are based on older sequence data and are not in complete agreement with the results of certain chemical and enzymatic studies that have been carried out to identify the single-stranded regions in the RNA molecules (for references, see Brosius et ai., 1978). It is known from early physicochemical analysis of rRNA secondary structures that about 65% of the nucleotide residues are base-paired and that the double-stranded regions are enriched with GC pairs (Cox, 1966; Cotter and Gratzer, 1969). However, until now only in the case of the rather short 3'-terminal fragment of the 16 S RNA has a direct physical method, viz. nuclear magnetic resonance (NMR) spectroscopy, been used to elucidate the secondary structure of this large rRNA molecule (Baan et ai., 1977).

Although the rRNA molecules retain most of their double-helical regions in going from solution to ribosome structure, significant changes in their secondary structure are brought about by the binding of ribosomal proteins. One can detect these alterations by a comparison of the circular dichroism (CD) spectra of RNA in the ribosomes and in the isolated form in the wavelength region of the main positive band (240-300 nm). The amplitude of the first positive CD band is 15-20% lower in the 30 S subunit than in the free 16 S


RNA (Shatsky et al., 1971a). The differences in the magnitude of this band must be caused exclusively by a change in the conformation of RNA, as the CD of ribosomal proteins at these wavelengths is negligible. It should be noted that analogous changes in CD spectra have been observed during the reconstitution of native 30 S subunits from 16 S RNA and 30 S proteins; in particular, the characteristic CD alterations accompany activation of the reconstitution intermediate particles (RI particles) (Kopylov et al., 1974). It is noteworthy that the spectra of the 16 S RNA and denatured 30 S subunits coincide (Shatsky et aI., 1971b).

Analogous alterations have been observed in the CD spectra of fragments of the 16 Sand 23 S RNA that contain specific binding sites for proteins S4 and L24 after interaction with these proteins (Chichkova et al., 1974; Tritton and Crothers, 1976).

In an attempt to account for the perturbations in the CD spectra, the difference spectra of the 16 S RNA in various conformational states were analyzed, and it was suggested that these alterations were caused by a decrease in both the number of double-helical regions and base-stacking interactions in the 16 S RNA (Drigina, 1975).

The interaction of the 5 S RNA with protein LI8 also produces specific changes in the 240-300-nm portion of the CD spectrum (Bear et aI., 1977; Spierer et al., 1978). In this case, however, the amplitude of the first positive CD band was 20-25% higher in the RNA-protein complex than in free 5 S RNA, and this effect was interpreted as being caused by the perturbation of a structural regularity within one of the double-stranded segments of the 5 S RNA (Spierer et al., 1978).

Differences in secondary structure between free rRNA and rRNA incorporated into the ribosome can be also detected by comparison of their melting curves and hypochromic effects. Although the differences in shape of the melting curves of the 30 S subunits and 16 S RNA in the presence of Mg2+ are significant, the hypochromicity of the 16 S RNA in 30 S subunits measured in solvents at low Mg2+ concentration is only about 3% lower than that of the isolated 16 S RNA (Shatsky et al., 1971b). It should be borne in mind, however, that this small effect could represent the melting of several dozens of base pairs in a RNA molecule with a length of 1540 nucleotide residues. In addition, it was shown in experiments with ribosomal particles irreversibly fixed with cross-linking reagents that the hypochromicity of the 16 S RNA in intact 30 S subunits was apparently overestimated, and the real difference in hypochromicity between free and intraribosomal 16 S RNA was more than 5% (Rezapkin et aI., 1977). More pronounced differences in the melting curves of the isolated 16 S RNA and 30 S subunits have been reported by Araco, Belli, and their co-workers (Araco et al., 1975; Belli et al., 1976). It is interesting that

Structure and Functions of Ribosomal RNA 85

specific binding of individual protein S4 with the 16 S RNA also produces a decrease in the uv absorbance of the RNA (Seals and Champney, 1976).

Nevertheless, a decrease in the number of nucleotide residues involved in base pairing or in stacking interactions as a result of the protein binding to rRNA does not necessarily mean that the RNA structure becomes disordered. On the contrary, such a rearrangement of RNA conformation is likely a necessary step in the formation of its specific intra ribosomal tertiary structure.

3. COMPACT FOLDING OF RNA IN RIBOSOMAL SUBUNITS

The tertiary structure of rRNA in ribosomes (as well as the intraribosomal secondary structure of the RNA) is predetermined in large part by the secondary and tertiary structure of free RNA in the isolated state (see also subsequent sections). However, under conditions optimal for functioning of ribosomes in vitro (Le., at 20-40 0 C and a MgH concentration of about 5 mM), RNA chains are much more tightly folded in ribosomal subunits than in the free state. This fundamental property of rRNA was first noted when the intrinsic viscosities (Spirin and Gavrilova, 1971) and electrophoretic mobilities in polyacrylamide gels (Mikhailova and Bogdanov, 1970) of free RNA and subunits were compared. It was also observed that rRNA in ribosomal subunits denatured in a more cooperative way than free rRNA, suggesting that in subunits rRNA molecules were packed in a more orderly fashion (Miall and Walker, 1969; Shatsky et al., 1971a).

Additional experimental evidence that ribosomal subunits are more compact than their constituent rRNA was provided by the laser light-scattering technique, which yields accurate values of the translational diffusion constant D, a sensitive indicator of variations in macromolecular size and shape. The possibility of discriminating between increases in the folding of a molecule and increases in its mass constitutes the obvious advantage of this approach. The values of D obtained for 30 Sand 50 S subunits are 10% greater than those for 16 Sand 23 S, respectively, despite the higher molecular weight of the subunits (Bogdanov et al., 1978). In general, it is impossible to distinguish between changes in size and changes in as symmetry from the diffusion constants of macromolecular aggregates, such as ribosomal subunits. However, it is reasonable to suggest that asymmetry of the 16 S RNA does not significantly change when incorporated into the 30 S subunit. Therefore, the diffusion constant measurements actually demonstrate that the attachment of ribosomal proteins to rRNA constrains the latter molecules to adopt a more compact tertiary structure.

These findings are in good agreement with measurements of the radii of


gyration of rRNA and of the volumes that rRNA occupies within ribosomal subunits (Serdyuk and Grenader, 1977; Serdyuk, 1978). For example, the volumes of 23 Sand 16 S RNA within the 50 Sand 30 S subunits, respectively, calculated from neutron-scattering data, appeared to be only 1.8 times those of the dry volumes of these RNAs. This ratio corresponds to very tight packing of the hydrated RNA molecules within the subunits (Serdyuk, 1978).

The ability of the rRNA chain to fold into compact structures is an inherent property of rRNA. Systematic analysis of the sedimentation constants of 16 S RNA and 30 S subunits as a function of the [Mg2+] /[K +] ratio shows, however, that the 16 S RNA attains a conformation as compact as that of the 30 S subunit only at a very high ("nonphysiological") concentration of Mg2+. At the same time, at a [Mg2+]/[K+] ratio optimal for ribosome functioning in vitro, the differences in the compactness of free 16 S RNA and 30 S subunits are maximal. The suggestion was made that under "physiological" conditions rRNA within ribosomes had a "stressed" conformation (Potapov and Bogdanov, 1977). This conformation is partially maintained by magnesium ions (see Spitnik-Elson and Elson, 1976, for review). But, as mentioned above, magnesium ions alone are not sufficient to hold this conformation. Ribosomal proteins play an important role in maintaining the "stressed" RNA conformation. Removal of certain proteins from subunits of ribosomes or alteration of a protein conformation within ribosomes would therefore cause unfolding of the structure of ribosomes. Indeed, unfolding of the 30 S subunit has been observed as the result of oxidation of its proteins with monoperphthalic acid (E. Skripkin, personal communication).

During the functional cycle, ribosomal subunits apparently undergo conformational changes (see section on functional role of rRNA). The intriguing question here is whether "the stressing" of rRNA molecules within ribosomes is connected with the structural flexibility of the subunits. This problem is discussed in more detail in subsequent sections.

4. DOMAIN ORGANIZATION OF RIBOSOMAL SUBUNITS

Ribosomal proteins not only maintain the compact conformation of rRNA within the ribosome, but they also seem to play an active role in the organization of the internal RNA structure in ribosomes. Prior to discussing this important question, however, it is necessary to introduce the concept of the organization of ribosomal subunits from structurally independent ribonucleoprotein segments (domains). Strictly speaking, the existence of ribonucleoprotein (RNP) domains has been documented only in the case of the 30 S subunit, which has received the most study. There are good reasons to believe, however, that 50 S subunits also contain RNP domains (e.g., Ktihlbrandt and Garrett,


1978; Spitnik-Elson et al., 1978). The stable complex of 5 S RNA with 50 S subunit proteins L5, LI8, and L25 that can be integrated into the large subunit (Yu and Wittmann, 1973) can be also considered a distinctive RNP domain.

The subdivision of the 30 S subunit into domains has been demonstrated by two independent methods. First, mild digestion of 30 S subunits with RNases produces two large RNP fragments (RNP I and RNP 11). RNP I consists of a RNA fragment consisting of approximately 900 nucleotides, originating from the 5' end of the 16 S RNA and proteins S4, S5, S6, S8, S15, S16j17, S18, and S20. An RNA fragment of 450-500 nucleotides from the 3' proximal region of the 16 S RNA and proteins S7, S9, S10, S14, and S19 have been identified in RNP II (see Brimacombe et al., 1976, for review). In addition, it was shown in these studies that the I 50-nucleotide 3'-terminal fragment of the 16 S RNA was not involved in strong RNA-protein interactions (see also Section 7). Second, the careful investigation of the 30S subunit reconstitution system, carried out by Zimmermann and his collaborators (reviewed by Zimmermann, 1974), suggests that the binding regions for three groups of 30 S proteins delineate distinct structural domains in the 5'-terminal, middle, and 3'-terminal thirds of the 16 S RNA. The first group contains proteins S4 (the main protein), S16, S17, and S20; the second group consists of proteins S8 and S15 (the main proteins), and S6 and S18; and proteins S7 (the main protein), S9, S13, and S19 comprise the third group. The interaction of the main protein from each group with the corresponding region of the 16 S RNA governs the association of other proteins from each group with the 16 S RNA. At the same time, the RNA-protein interactions within one domain are relatively independent of the association with the 16 S RNA of proteins from other domain. All these results are summarized in Figure 1, which demonstrates that RNP I is divided into two parts or two domains (RNP I' and RNP I"). RNP I' is formed from the 5' third of the 16 S RNA molecule and proteins of the S4 group, whereas RNP I" consists of a comparatively short RNA fragment from the middle third of the 16 S RNA and proteins of the S8jS15 group.

Thus ribosomal subunits consist of large RNA-protein segments (domains) stable enough to exist autonomously and can be reconstituted from their RNA and protein components independently of other parts of the subunits.

Inside the domains there are strong RNA-protein and RNA-RNA interactions. It is important to note that not only neighboring RNA regions, but also quite distant ones, are involved in the interactions (Figure 1). Indeed, limited hydrolysis of the S4-16 S RNA complex or of free 16 S RNA in solution at high Mg2+ concentration with RNase A covalently bound with Sepharose yielded large RNA fragments with a total molecular weight of about 130,000. The nucleotide sequences of RNA fragments isolated both from the complex and from free 16 S RNA were identical (C. Ehresmann et a/., 1977b; Ungew-


Cobra venom nuclea&e hydrolysi&

-l- ,J" '" ",,J,, ,J., ,J., reduced in70S

.t, U ,J.,J.'JUU enhanced in70S ~in70S

T1RNase hydrolysis reduced in70S

-l- '" U Un-ketoxe\ modification red~70S

'" L -l-H"H'HclF QRG M B I' r l c· c th~ 0 I ' • 5~ 11111 1111 I II II I III I I o I

500 I

S4 ----=------------:::L

1i'.w. J. :J, U m 0' 0 E' K P P' E A J

I II II I I f-3' 1000

I

RNPu

1500 I

S8.M5 __ ~S7~ ________ _

~<® L~O_~~S4~===~=8=s=6=15=S1=8===~=S7===S===r~S13 519

FIGURE 1. Summary of data on location of binding sites of main proteins of RNP domains on the 16 S RNA and accessibility of the 16 S RNA in the 30 S subunit. The scale indicates number of nucleotides from the 5' end of RNA. Capital letters refer to sections generally used in 16 S RNA sequencing work (Fellner, 1974). The regions of the 16 S RNA corresponding to different RNP domains and associated with the main proteins of domains are indicated by bold lines (Zimmermann, 1974; Brimacombe et al., 1976). The dotted line indicates the RNA fragment in sections D'O, which forms a tertiary interaction with S4-RNA (C. Ehresmann et al., 1977b; Ungewickell et al., 1977). Thin solid lines represent cross-linked protein pairs and indicate their close proximity in 30 S subunits (SHiffler and Wittmann, 1977). Arrows indicate the positions of T I RNase (Santer and Shane, 1977) and kethoxal (Chapman and Noller, 1977) reactive sites, representing single-stranded RNA regions exposed in 30 S subunits. The sites of hydrolysis of the 16 S RNA in 30 S subunits with cobra venom nuclease (S. K. Vasilenko, C. Ehresmann, and J.-P. Ebel, personal communication) corresponding to exposed double-stranded fragments are also shown. The alterations in accessibility of exposed RNA regions after association of 30 Sand 50 S subunits are shown.

ickell et al., 1977b). Under denaturating conditions, the large RNA fragments dissociated into noncontiguous subfragments with some sequence regions separated by about 400 nucleotides (see Garrett et al., 1977, for review). Thus the tertiary structure of the RNA portion of the RNP I' domain is maintained by specific MgH -dependent RNA-RNA interactions. Large RNA fragments, enriched with guanine nucleotides, could also be produced from the 16 S RNA-S4 complex by exhaustive RNase A hydrolysis. It was suggested that interactions between guanylic acid residues were important for maintaining the tertiary structure of this region of the 16 S RNA (Chichkova et al., 1974). It is interesting that a 500-nucleotide RNA fragment with strong RNA-RNA


interactions has been isolated from the 5'-terminal portion of the 23 S RNA (Sloof et al., 1978), thought to be involved in the formation of an RNP domain with L24 as the main protein (Kiihlbrandt and Garrett, 1978).

The capacity of proteins S8 and S15 to interact with a "denatured" form of the 16 S RNA indicates that the RNA part of the RNP I" domain also has very stable conformation in the isolated state (Muto and Zimmermann, 1978). It is not surprising, because the binding sites for S8 and S 15 appear to be organized exclusively from rather long double-stranded RNA fragments (Zimmermann et al., 1975). In addition, the protein-binding sites in this region appear to be very highly conserved in prokaryotic 16 RNAs (Thurlow and Zimmermann, 1978).

On the other hand, the RNA segment corresponding to tbe S7 domain (RNP II) is highly degraded under conditions in which the S4-RNA is completely stable (Muto et al., 1974; Zimmermann et al., 1975). The compact folding of the RNA fragment in this domain is apparently maintained by ribosomal proteins. Indeed, the association of protein S7 with the 16 S RNA stabilize the RNA chain against unfolding at low MgH concentration and promote the formation of a more compact tertiary structure at high MgH concentration as revealed by the D values of free RNA and its complexes with the protein (Bogdanov et aI., 1978).

Although the internal tertiary structure of the RNA portion of the RNP I' domain is stable in the absence of protein S4, influence of this protein on the overall 16 S RNA conformation is pronounced. For example, the accessibility of the 16 S RNA-S4 complex to complementary binding with hexanucleotides is 40% lower than that of the RNA alone (Kopylov et al., 1975; see also Section 7).

It is customary to assume that the strong influence of proteins S4, S7, and S15 on the 16 S RNA structure is related to their elongated conformation in the 30 S subunit, which allows them to interact simultaneously with quite distant RNA regions. However, the concept of the highly extended conformation of these proteins is open to question at the present time. First, it was found that proteins S4, S7, and S15 in isolated form possessed a pronounced secondary structure which must be stabilized by a hydrophobic core (Spirin et al., 1979). Second, measurements of the radii of gyration (Rg) of proteins S4 and S7, isolated by a very mild procedure, have shown that Rg values for these proteins are relatively small and typical of compact globular proteins (Serdyuk et al., 1978). Third, stereochemical analysis of the complete amino acid sequences of some of these proteins also supports the idea that their conformation is globular in solution (Lim et al., 1978). Although the results of these studies are inconsistent with certain other physical measurements on these proteins (e.g., Paradies and Franz, 1976; Osterberg et al., 1977; Giri and Franz, 1978), they


demonstrate unambiguously that the polypeptide chains of these proteins are at least capable of folding into compact globular conformations in solution. In addition, the results of immunoelectron-microscopic studies on ribosomal protein topography, which provide the most convincing arguments in favor of an asymmetric, elongated structure for many ribosomal proteins, need further verification. For example, two groups of investigators have claimed that protein S4 revealed multiple antibody binding sites at widely separated points of the 30 S subunit surface (reviewed by SWiller and Wittman, 1977). It was reported recently, however, that in the 30 S subunit this protein is not accessible for binding with highly purified antibodies (Winkelmann and Kahan, 1978).

If the main proteins of the RNP domains are not very extended in subunits (protein S8, for instance, is undoubtedly globular), their influence on the 16 S RNA structure cannot be a result of the local rearrangements of proteinbinding sites. Rather, these effects have to be attributed to interdomain interaction, which will be discussed in the next section.

Direct protein-protein interactions probably do not playa significant role in the organization of the domain structure as, after the cross-linking of all the neighboring proteins in ribosomal subunits by polyaldehydes of up to 25 A chain length, fixed particles unfold on removal of MgH ions in the same manner as native ones (Rezapkin et al., 1977; also see Kurland, 1974, 1977a, for discussion). It would be wrong, however, to argue that protein-protein contacts do not exist in ribosomes. Specific interactions between isolated proteins have been observed in solution (Rohde et al., 1975; Aune, 1977) and the N-terminal region of protein S4, which is not involved in RNA-protein interactions, is essential for binding of several proteins within the 30 S subunit (Changchien and Craven, 1976). In addition, protein-protein interactions can be very important for ribosome functioning. We would like to emphasize here only that this type of intra ribosomal interactions is not so important for the organization of the overall structure of ribosomes as are RNA-protein and RNA-RNA interactions.

It has to be noted that existence of the relatively independent structural regions in ribosomal particles is consistent with various known properties of ribosomes, for instance, with stepwise unfolding (reviewed by Spirin, 1974) and stepwise melting (Fox et al., 1978) of their subunits. The pr~sence of certain structural domains in the 30 Sand 50 S subunits was also proved by investigation of unfolding of subunits cross-linked with reagents of different length (Rezapkin et al., 1977). Finally, the separate structural regions in ribosomal subunits can be observed by electron microscopy (Vasiliev, 1974; see also Lake, 1976, for references). At the present time, it is hard to say, however, how these structural segments of ribosomes are related to the RNP domains described in this section.


5. ROLE OF RIBOSOMAL PROTEINS IN THE ORGANIZATION OF RNA TERTIARY STRUCTURE WITHIN RIBOSOMAL SUBUNITS

The domain structure of rRNA is therefore determined either by the preexisting stable conformation of large rRNA segments, or it is formed as a result of RNA-protein interactions. How, then, does the interaction between the domains, which is the last stage of the organization of the compact native structure of ribosomal subunits, occur? The first factor to be taken into consideration is that the formation of this structure involves very few proteins. In the case of the 30 S subunit, these proteins have been identified as the major proteins of the domains, i.e., S4, S8, SIS, and S7 (Kopylov et aI., 1976). That this group of proteins has a special role can be inferred from the following facts. First, the interaction of only these four proteins with the 16 S RNA transforms its secondary structure into the one that is characteristic of an intact 30 S subunit (Drigina, 1975). Second, the 16 S RNA in RNP particles, which contain only proteins S4, S8, SIS, and S7, binds many fewer dye molecules and is much more resistant to various RNases than is free RNA (Kopylov et ai., 1976). Third, exposed regions of free 16 S RNA capable of binding with complementary hexanucleotides are sterically hindered in the complex of S4, S8, SIS, and S7 with the 16 S RNA to almost the same degree as in the 30 S subunit (Kopylov et ai., 1975). These effects are so strong that they cannot be explained simply by direct screening of the RNA sites by physical contact with the bound proteins. It was suggested that these proteins stimulate certain regions of RNA to participate in intra ribosomal interactions that result in the formation of particles almost as compact as 30 S subunits. This hypothesis was proved valid by electron microscopy and laser light scattering. In the first case, it was shown that the complex of the 16 S RNA with S4, S7, S8, and SIS appeared to be very similar to the 30 S subunit both in size and morphology (Vasiliev et ai., 1977a; also see Section 7). In the latter case it was established that this complex had a maximal D value exceeding that for the 30 S subunit (Bogdanov et ai., 1978).

A group of six proteins that are prominent in the organization of a compact RNA structure has also been identified in the 50 S subunit. These proteins are L3, L4, L13, L17, L22, and L29. In association with the 23 S RNA, they protect large RNA regions, such as the 13 Sand 18 S fragments from Tl RNase hydrolysis (Branlant et aI., 1977). By contrast, the 23 S RNA-protein complex lacking L4, L22, and L29 is much more sensitive to RNase Tl treatment. One can suggest that these three proteins stimulate interdomain RNARNA interactions between distant RNA regions (see also Figure 2). The binding of L3, L4, L13, L17, L22, and L29 with the 23 S RNA is thought to bring


I ABC D E F G ~, I JKL 1M N 0 I P Q R STU V W X V Z

5' 3'

~ ! L 24 :500 i 100 1, 00 i 10?0 I: 5S E~A : 500 : : i i: 9 I I I I I II L18 I I I I II I ! lil, l22, (L29?) ,rn--L13 __ ntJ.!£..J 'L3 1' L25 I "l....J' I

L_f;========================~ _____________ ~ FIGURE 2. Location of protein binding sites on the 23 S RNA. Numbers denote distances in nucleotide residues from the 5' and 3' ends of the molecule. Uppercase letters refer to sections of the 23 S RNA. Protected fragments recovered from T, RNase digest of 23 S RNA-protein complexes are shown by dashed lines. The tentative location of the 5 S RNA binding site on the 23 S RNA is also indicated. Dashed lines indicate RNA fragment protected from RNase hydrolysis with the same protein. [Based on the data of Branlant et al. (1977).]

together the 5'- and 3'-terminal regions of RNA, resulting in the formation of a stable complex between two complementary octanucleotide sequences (Branlant et al., 1976).

The 3'- and 5'-end regions of 16 S RNA are also in close proximity in the 30 S subunit. This follows from the analysis of 30 S protein topography (see Figure 1), from RNA-protein cross-linking data (Moller et al., 1977) and from the fact that the binding of proteins of the 5' proximal domain (RNP I') with 16 S RNA induces the rearrangement of its 3' proximal region and makes the latter accessible to enzymatic methylation (Thammana and Held, 1974). Considering that the 3'- and 5'-terminal nucleotide sequences of the 5 S RNA undoubtedly form a double-stranded stem (see Erdmann, 1976, for review; Wagner and Garrett, 1978), one can conclude that the close opposition of rRNA termini is a common property of rRNA structure in ribosomes. This is apparently important for interactions between RNP domains.

Up to this point, we have emphasized that under physiological conditions ribosomal proteins help to maintain the tightly folded RNA conformation. The intriguing question involves the actual role of proteins in the creation of this conformation. To attempt an answer to this question, let us consider some data that have been obtained from studies on the reconstitution of ribosomal subunits. It is known that rRNA samples extracted from ribosomes represent a set of conformers (see Hochkeppel and Craven, 1976b, for references). The conformational heterogeneity of rRNA can be shown by gel electrophoresis (e.g., Ungewickell et ai., 1977a). The efficiency of subunit reconstitution depends, to a great extent, on the proportion of various conformers in the rRNA preparation, some of which can be considered denatured forms of rRNA molecules. However, certain forms of inactive rRNA can recover the ability to interact with ribosomal proteins when heated at 40°C in the presence


of MgH (Zimmermann, 1974; Muto and Zimmermann, 1978). In addition, the capacity of rRNA to interact with individual proteins has been found to depend on the procedure used for the extraction of rRNA from ribosomes (Hochkeppel and Craven, 1976a; Hochkeppel et al., 1976).

Taking into account this property of rRNA molecules, one can suggest two alternative mechanisms for the participation of ribosomal proteins in the organization of rRNA tertiary structure: (1) proteins change the initial RNA conformation and thereby induce new RNA-RNA interactions, which lead to the tight packing of rRNA molecules within RNP particles; and (2) from an equilibrium mixture of conformers, proteins choose the rRNA molecules, the conformation of which is identical (or very similar) to that of RNA within subunits, bind to them and displace the equilibrium toward rRNA molecules with a proper conformation. At present it is hard to give preference to either of these mechanisms, although the first seems more probable to us.

Indeed, the existence of conformational transitions in rRNA structure during ribosomal subunit reconstitution is beyond question. This is documented in part by data already discussed, including alterations in rRNA CD spectra and the interrelationship of the RNA-binding capacity of different proteins (see Sections 2 and 3). Such transitions are also evident from the increase in the accessibility of rRNA to nucleases (Mackie and Zimmermann, 1978) and to processing enzymes (methylases) (Thammana and Held, 1974; Dahlberg et al., 1977) that result from the binding of certain proteins whose binding sites are distant from the sites of modification.

Furthermore, the reconstitution of ribosomal subunits proceeds at a reasonable rate only at relatively high temperatures. The binding of ribosomal proteins to rRNA at temperatures of 0-20°C gives an inactive intermediate RNP particles containing an incomplete set of proteins (reviewed by Nomura and Held, 1974). The intermediates can be activated at higher temperature (40°C); however, this process is followed by alterations in secondary and tertiary structure of both rRNA (Kopylov et al., 1974; Potapov et al., 1973) and proteins (Lemieux et aI., 1974). Under conditions optimal for reconstitution of the 30 S subunit, the tertiary structure of free RNA is partially unfolded (Schulte et al., 1974), and the ordered elements of RNA secondary structure (hairpin loops) are quite flexible (Potapov and Bogdanov, 1977). Therefore, the number of rRNA molecules with a compact structure must be very small in the reconstitution mixture. A rise in flexibility of rRNA molecules, on the other hand, facilitates changes in their conformation. Ribosomal proteins apparently exploit this situation and transform the RNA conformation into compact form even at high temperatures (Potapov and Bogdanov, 1977).

In conclusion, we would like to emphasize that whatever the real mecha-


nism of the compact folding of RNA molecules within ribosomes, it must be based on the inherent conformational flexibility of rRNA structure.

6. DIRECT PARTICIPATION OF RIBOSOMAL RNA IN RIBOSOME FUNCTIONING

The first experimental evidence favoring the involvement of rRNA in the functional centers of the ribosomes was obtained in the pioneering work of Budker and co-workers (Budker et aI., 1972; Bochkareva et aI., 1973). These workers synthesized a peptidyl derivative of tRNA containing a chemically reactive label, showed that it could be used as a substrate for the donor site (P site) of the peptidyl transferase of the 70 S ribosomes and then found that the label reacted predominantly with the 23 S RNA molecules. They also described the efficient affinity labeling of the 23 S RNA with chemically reactive analogues of mRNA (Girshovich et al., 1974).

The affinity and photoaffinity labeling of the 5 S, 16 S, and 23 S RNA in ribosomes were subsequently observed by many authors in experiments with various derivatives of aminoacyl- and peptidyl-tRNA, mRNA, and inhibitors of protein synthesis (reviewed by Pelligrini and Cantor, 1977; Towbin and Elson, 1978). In addition, early studies on the functional importance of rRNA demonstrated that the binding of tRNA to the A site protected six or seven guanine residues in the 16 S RNA component of the 30 S subunit from chemical modification by kethoxal (3-ethoxy-butanon-2-al-l) (Thomas et aI., 1975).

Neither affinity labeling nor chemical modification experiments, however, can answer the question of whether rRNA is directly involved in interactions with tRNA, mRNA, and other constituents of the protein biosynthesis apparatus. Although affinity labeling permits the identification of components that are in close proximity to the activated group, they need not necessarily be in direct contact with the tRNA, mRNA, etc., bearing this group. On the other hand, chemical reagents that are supposed to be specific for RNA modification must also modify ribosomal proteins, because the reactivity of the latter is usually higher than that of RNA bases. Consequently the results of rRNA chemical modification within ribosomes are ambiguous as well.

Nevertheless, the direct interaction of tRNA with rRNA does exist in functioning ribosomes. This has been shown in experiments in which 70 S ribosomes with certain N-acetylaminoacyl-tRNAs bound noncovalently at the P site were irradiated with light of 300-350 nm (Of eng and et aI., 1979; Zimmermann et al., 1979; Prince et al., 1979). Under these conditions the tRNA molecules became covalently attached exclusively to the 3' third of the 16 S RNA chain. From a comparison of active and inactive tRNA sequences, it was


suggested that the reactive nucleotides are the modified uracil residues [5-methoxyuridine (m05U), 5-carboxymethoxyuridine (cm05U), and 5-methylamino-methyl-2-thiouridine (mam5slu)] that occur in the "wobble" position of various tRNA anticodons. Because the cross-linking was codon-dependent, it was concluded that tRNA, mRNA, and 16 S RNA were all in very close proximity at the decoding center of 70 S ribosomes when peptidyl-tRNA occupied the P site. Other affinity-labeling data suggest that the 5' portion of the 16 S RNA is also located close to the decoding center (Wagner et al., 1976).

The binding of aminoacyl-tRNA at the A site of the peptidyl transferase center is believed to involve the interaction between the common TlfCG sequence of tRNA and a complementary sequence in the other RNA component of ribosomes, namely the 5 S RNA (see Erdmann, 1976; and Of eng and, 1977, for reviews). This proposal is based on the observation that the oligonucleotide T lfCG strongly inhibits binding of the complex of phe-tRN A and elongation factor EF-Tu to the ribosomal A site (Of eng and and Henes, 1969; Richter et al., 1973). However, this hypothesis contradicts certain other findings: the TlfC loop of tRNA is not normally exposed when tRNA is in the free state, for instance, whereas the residues of guanylic acid in the appropriate region of the 5 S RNA are not accessible to kethoxal modification when the 5 S RNA is incorporated to the 50 S subunit (Noller and Herr, 1974). To overcome these difficulties it has been suggested that unfolding occurs on ribosomes (Erdmann, 1976) and, in the case of tRNA, evidence that the tlfCG sequence in the tRNA becomes exposed upon codon-anticodon interactions has recently been obtained (see Schwartz et at., 1978, for discussion). Further conflicting results have been obtained by R. Liihrmann (personal communication), who showed that the inhibitory effect of TlfCG depends very much on the quality of the oligonucleotide preparations. Highly purified TlfCG samples did not show any inhibitory effects on EF-Tu and polyU-dependent phe-tRNA binding to 70 S ribosomes. Therefore, much more study is required in order to elucidate the mechanism of tRNA binding at the ribosome A site.

Several functional activities of the 30 S subunit of ribosomes are obviously related to the 3'-terminal segment of the 16 S RNA. Its functional importance has been discovered through investigations of the mode of protein synthesis inhibition in E. coli cells by the antibiotic colicin E3. This inhibitor and cloacin DG 13 happen to be the specific nucleases that produce a single cleavage near the 3' end of 16 S RNA in 70 S ribosomes (reviewed by Nomura et at., 1974; Baan et at., 1976). As a result, a small fragment of about 50 nucleotides containing 3' end of the 16 S RNA is produced. The appearance of this single nick in the 16 S RNA chain inhibits both the functioning of initiation factor IFI and the translocation of peptidyl-tRNA from the A to the P site (Baan et al., 1976,1978a,b).


Another type of modification of the 3'-terminal segment of the 16 S RNA, i.e., the absence of methylation of two successive adenylic acid residues-24 and 25 bases from the 3' terminus-makes E. coli cells resistant to kasugamycin (Helser et aI., 1972).

A very different situation has been found in 23 S RNA-methylation of 23 S RNA molecules in 50 S subunits of Staphylococcus aureus (Lai et al., 1973a,b) and Streptomyces azureus (Cundlife, 1978) results in ribosomemediated resistance to erythromycin and thiostrepton, respectively.

The relationship of ribosomal kasugamycin sensitivity to 16 S RNA methylation is not well understood. Interestingly, the antibiotic does not interact directly with the two dimethyl adenine residues (Am~), as they are not exposed to solution and are not accessible for specific antibodies in the native 30 S subunit and 70 S ribosome (Thammana and Cantor, 1978). Nonetheless, the methylated part of 16 S RNA must be able to exert an indirect influence on the kasugamycin-binding site. It must be mentioned here that the conformation of this part of the 16 S RNA is quite flexible and is affected by proteins from the 5'-proximal and central domains of the 30 S subunit (Thammana and Held, 1974).

It has been known for some time that the 3'-terminal segment of 16 S RNA is involved in mRNA recognition by ribosomes. According to the hypothesis of Shine and Dalgarno, the 16 S RNA in 30 S subunits forms a specific complex with mRNA owing to complementary interaction between a polypyrimidine sequence near the 3' end of the 16 S RNA and polypurine stretch located at the 5' side of the initiation codon in the initiation region of most mRNAs (Figure 3) (Shine and Dalgarno, 1975). The first direct evidence of this model came from the work of Steitz and lakes (1975), who isolated a double-stranded RNA-RNA hybrid consisting of the colicin fragment from the 3' end of the 16 S RNA and the A protein initiation region from bacteriophage R17 RNA. Since then, the hypothesis of Shine and Dalgarno has been treated in great detail in several reviews (e.g., Kurland, 1977a,b; Brimacombe et al., 1978; Brimacombe, 1978). However, we would like to mention two very recent confirmations of this mechanism.

First, it was found that the oligonucleotide AGAGGAGGUoH, which is complementary to the 3' end of the 16 S RNA, strongly inhibits the formation of the initiation complex between Qf3 RNA and 70 S ribosomes, but does not affect the binding of the AUG codon to ribosomes (Taniguchi and Weissmann, 1978). Second, the analysis of a mutation that changes the preinitiation sequence in the mRNA for the gene 0.3 protein of bacteriophage T7 shows that a decrease in complementarity between the 3'-terminal sequence of the 16 S RNA and the preinitiation sequence of mRNA strongly depresses the synthesis of the 0.3 protein. This mutation can be suppressed by a second mutation


FIGURE 3. Location of functional sites on the 3'-terminal segment of the 16 S RNA. The arrow indicates the position of hydrolysis of the 16 S RNA in 70 S ribosomes with colicin E3. Solid box indicates the RNA sequence involved in the interaction with mRNA initiation sites. Dashed box indicates the trinucleotide sequence, which is probably involved in the interaction with nonsense codons. The complementarity of the 3' ends of the 16 Sand 23 S RNA (smaller base symbols) is also shown (see text for details). The secondary structure of the 3'-terminal segment of the 16 S RNA is given in accordance with NMR data (Baan et al., 1977). Ksg, the methylated sequence of the 16 S RNA responsible for kasugamycin-sensitivity of E. coli ribosomes.

in this region, which restores both the complementarity of the Shine-Dalgarno complex and the synthesis of the 0.3 protein (Dunn et al., 1978).

An alternative hypothesis has been put forward by Van Knippenberg (1975), who proposed the formation of a double-stranded complex between the preinitiation sequence of mRNA and a complementary sequence located near the 5' end of 16 S RNA. So far as we are aware, no experimental proof for this proposal has been published.

It has also been suggested that the 3' end of the 16 S RNA participates in the termination of protein synthesis and in the association of 30 Sand 50 S subunits. The first suggestion is based on the presence of the UUA sequence at the 3' end of the 16 S RNA, which is complementary to all three nonsense codons (Shine and Dalgarno, 1975). The second hypothesis has been advanced on the basis of the pronounced complementarity between the 3' ends of the 16 Sand 23 S RNAs shown in Figure 3. Attempts to check the latter hypothesis by means of psorlens, which cross-link the bases in double-helical nucleic acids (Shen and Hearst, 1976; Isaacs et al., 1977), have been of no avail (A. A.


Bogdanov, unpublished results). At the same time, treatment of 70 S ribosomes by bis(2-chloroethyl)amine (Ulmer et aI., 1978), results in cross-linking of the 16 Sand 23 S RNAs.

There are good reasons to believe that rRNA interacts with protein synthesis factors. It is particularly clear in the case of the initiation factor (IF) 3. The IF3 binding sites are apparently located on the 16 S RNA. Indeed the modification of guanine residues by kethoxal and the oxidation of adenine residues with monoperphthalic acid strongly inhibit the binding of IF3 to the 30 S subunit (Pon and Gualerzi, 1976). This binding also decreases in the presence of RNA-specific ligands and after limited degradation of the 16 S RNA in 30 S subunits witl: RNases. On the other hand, the selective modification of ribosomal proteins, as well as the limited tryptic hydrolysis of 30 S subunits, did change the capacity of the ribosomes to bind IF3 (Gualerzi and Pon, 1973; Pon and Gualerzi, 1976). 30 S subunits lacking more than half their proteins bind IF3 to the same extend as do native ones (Gualerzi and Pon, 1973; Laughrea et aI., 1978). There is some evidence that the IF3 binding site is located near the center of the 16 S RNA chain (M. Grunberg-Manago and J.-P. Ebel, personal communication). IF3 has been also cross-linked in a low yield to the oxidized 3' end of the 16 S RNA (Van Duin et al., 1975). It is unlikely, however, that any strong binding sites of IF3 are located in this region of the 16 S RNA, as removal of the colicin fragment does not change IF3-30 S subunit interactions (Laughrea et aI., 1978).

Until now, interactions between rRNA and other initiation, elongation, and termination factors in ribosomes have not been studied, although there are some indications that IFI and IF2 (Brimacombe et al., 1978), as well as EF-G (Koteliansky et aI., 1977; Sander et al., 1978), bind to rRNA.

Taking all these data together, it may be said that rRNA molecules are involved in functional interactions with most, if not all, components of the protein-synthesis apparatus. It is not excluded, however, that the functional role of rRNA in ribosomes goes beyond the scope of the participation in the arrangement of tRNA, mRNA, and protein biosynthesis factors on a ribosome surface.

Indeed, as was originally suggested by Spirin and co-workers (Lerman et al., 1966; Gavrilova et al., 1966), interactions between macromolecules within ribosomes are of a cooperative nature. This is manifest in the cooperative RNA-protein interactions that occur during reconstitution of ribosomal subunits in vitro (see Kurland, 1974, for discussion). This is also seen in the cooperative character of structural transformations, which ribosomal subunits undergo when the environmental conditions are changed (reviewed by Spirin, 1974; Spitnik-Elson and Elson, 1976). Long-range cooperative effects are also involved in the association of ribosomal subunits, since formation of 70 S ribosomes results not only in the shielding of certain parts of the ribosome, but also


in higher reactivity of some sites in their RNA (for 30 S subunits, it is shown in Figure 1) and in many proteins (see, e.g., Ginzburg and Zamir, 1975). Finally, it is in terms of the cooperativity of the ribosomal particle as a whole that one can explain why the binding of a small ligand with a ribosome causes (or prevents) a change in its entire structure. Suffice it to mention that the binding of antibiotics to the small ribosomal subunit makes the proteins of the large subunit more likely to undergo chemical modification (Martinez et ai., 1978). Also, the binding of streptomycin to the 30 S subunit stabilizes its whole structure, making it more stable in solutions with a low concentration of Mg2+ ions (reviewed by Spitnik-Elson and Elson, 1976). It is therefore not surprising that significant changes in the overall structure of the ribosome are observed when the latter interacts with protein factors. For example, initiation factor IF3 increases the accessibility of many proteins of 30 Sand 50 S subunits to modifying agents (Michalski et ai., 1976; Michalski et ai., 1978) and sharply changes the pattern of the proteins of the 30 S subunit, which can be crosslinked with 16 S RNA by uv irradiation (Broude et ai., 1978).

It is evident that the cooperativity of the interaction between the components of ribosome, like any other fundamental feature of its structure, must be associated with the mechanism of its functioning in protein biosynthesis.

On the basis of what was said above about the role of various types of interactions within the ribosome in the organization of its structure, it may be argued that the cooperativity of the ribosome is determined by its RNA component or, to be more precise, by the specific manner in which the RNA macromolecules are organized within the ribosome. Thus the major functional role of rRNA most likely consists of ensuring the conformational mobility of the ribosome as a unit, and of the transmission of allosteric effects throughout the whole ribosome.

7. TOPOGRAPHY OF rRNA IN RIBOSOMES

Two major tendencies can be distinguished in the investigations of the topological aspects of RNA organization in ribosomes: (1) the elucidation of the general principles of the spatial relationships between rRNA and proteins in the ribosomal particle, and (2) the identification ofrRNA fragments located either inside or on the surface of the distinct morphological parts of small and large ribosomal subunits. In an effort to solve these problems, a number of physical and biochemical approaches have been employed.

X-ray and neutron-scattering techniques have been used in an attempt to define some general regularities in the packing of rRNA and proteins in ribosomal subunits in solution. Despite their low structural resolution, these methods provide reliable information on the arrangement of ribosomal components


(see Serdyuk, 1978, for review). Earlier studies suggested that the distribution of rRNA and proteins in the 50 S subunit is not uniform with proteins located mainly on the particle surface and the 23 S RNA concentrated closer to the center of the subunit (Serdyuk et aI., 1970). This inference was confirmed by the use of the neutron-scattering technique in solvents composed of H20-D20 mixtures (Stuhrmann et al., 1976; Serdyuk, 1978), as well as by an approach combining light, X-ray, and neutron scattering (Serdyuk and Grenader, 1975, 1977), which allows direct and independent measurement of rRNA and protein parameters in ribosomes. The distribution of RNA and protein in the 30 S subunit has been found to be similar, but not identical, to that of the 50 S subunit (Beaudry et al., 1976). The 16 S RNA also forms the central core of the particle, but its shape seems to be more asymmetrical than that of the 23 S RNA (Serdyuk, 1978).

As indicated above, ordered segments of RNA macromolecules are compactly packed within ribosomal subunits. A working model of tight RNA folding has been proposed by Potapov and Bogdanov (1974), in which the doublehelical RNA segments are packed in a parallel manner and are held together by very specific magnesium ion-phosphate bonds. It may now be possible to determine directly the organization of ordered segments of the rRNA in ribosomes by means of scattering techniques. The characteristic maxima of the Xray-scattering curves of ribosomal particles in the region of medium scattering angles (corresponding to Bragg distances of 4.5-2.0 nm), which have been observed by many authors (see Serdyuk and Grenader, 1977, for references), appear to be caused by the RNA component of ribosomes (Serdyuk, 1978; Spirin et al., 1979). The elucidation of the nature of these maxima may, therefore, clear up the real spatial arrangement of ordered portions of the rRNA.

Because ribosomal particles are not amenable to high-resolution X-ray crystallography, the fine details of the RNA topography within ribosomes can be investigated only by indirect and time-consuming approaches. Identification of protein binding sites is of particular importance here because the study of ribosomal protein topography is far ahead of that of rRNA, and from the known sequence of these sites along the RNA macromolecule, the pattern of polynucleotide chains folding within ribosomal subunits may be deduced.

The most important results in this area of research have been obtained by Zimmermann, Garrett, Brimacombe, and their collaborators (reviewed by Brimacombe, 1978). A recent development in this field is the use of covalent RNA-protein cross-linking, which has several advantages. When rRNA is cross-linked to a protein by uv irradiation, the oligonucleotides, which are in physical contact with the protein, could be identified. This has been done with the complexes of the 16 S RNA with S4, S20 (B. Ehresmann et al., 1977), and S7 (Zwieb and Brimacombe, 1978). Using bifunctional cross-linking reagents a more extended rRNA area surrounding the given protein, including


RNA regions which are not involved in direct contact or strong interactions with proteins can be distinguished (Moller et al., 1977; Ulmer et aI., 1978). Unfortunately, very few of the ribosomal protein binding sites identified so far consist of short oligonucleotide sequences. In most cases protein binding sites have been isolated within rather large regions of the rRNA (see Brimacombe, 1978, for review). The resolution of the methods used in protein topography studies is also not very high. Moreover the results of different groups employing immunoelectron microscopy-considered to be the most powerful technique for the elucidation of protein spatial organization in ribosomes-are in disagreement (compare, e.g., Tischendorf et ai., 1975, with Lake and Kahan, 1975). As mentioned previously, the reliability of the observations which indicated that many ribosomal proteins are elongated both in the isolated form and within subunits, is presently open to question. In other words, much additional data are still necessary in order to combine the results of the topographic studies on RNA and protein into a harmonious scheme.

The investigation of rRNA segments which are exposed in the subunits is a point of great interest. Indeed, these "open" RNA regions are very likely to be involved in the interactions of ribosome functional centers with tRNA, mRNA, initiation factors, and so forth, as well as in subunit association. There are many recent reports concerning the determination of regions of the 16 S RNA that are accessible to hydrolysis with different nucleases. These data are partly summarized in Figure 1. They show, in particular, that the 3'-terminal region of 16 S RNA comprises the largest unprotected part of this molecule within the 30 S particle (Santer and Santer, 1972; Santer and Shane, 1977). These results are in a good agreement with the high functional activity of this portion of the 16 S RNA (see Section 6). RNase hydrolysis, as well as chemical modification of the guanine residues by kethoxal, also appear to be very useful for the identification of the 16 S RNA regions located at the interface between the 30 Sand 50 S subunits (Santer and Shane, 1977; Chapman and Noller, 1977).

Valuable information about the exposed rRNA regions in ribosomes may also be obtained by isolation and sequencing of oligonucleotides released into solution after limited RNase hydrolysis under the conditions in which the protein composition and sedimentation constant of a ribosomal subunit are not changed. This approach have been used to identify exposed oligonucleotide sequence in the 16 S RNA (Teterina et aI., 1978a,b), most of them are located in the 3'-terminal portion of the RNA (N. L. Teterina, unpublished results).

The binding of complementary oligonucleotides also appears to be useful for the investigation of rRNA topography in ribosomes. In particular, it has been shown by this technique that the 5' end of the 16 S RNA is also located on the surface of both the 30 S subunit and the 70 S ribosome (Skripkin et ai., 1979) and is therefore available for functional interactions.

102

a

o

c

~

b

A. A. Bogdanov et al.

FIGURE 4. Electron micrographs of (a) 168 RNA; (b) 168 RNA-protein 84 complex; (c) 168 RNA-protein 84, 87, 88, and 815 complex; and (d) native 30 8 subunits. [Presented on the basis of the data from Vasiliev (1974) and from Vasiliev et al. (1977a,b, 1978).] The schematic representations of images are shown at the top of each set of micrographs.


The only approach that allows one to observe directly the localization of certain ribosomal components within their subunits is electron microscopy (EM). Immuno-EM is especially promising in this respect. Nonetheless, conventional EM techniques, which have contributed so much to the determination of the size and form of the ribosomal subunits, can also be applied to the elucidation of the topography of ribosomal components. Thus Vasiliev et al. (Vasiliev et al., 1977a,b, 1978) used shadow casting to study lyophilized RNP particles with various sets of proteins, beginning with intact RNP particles and ending with 16 S complexes with protein S4 alone. In the same series of experiments, free 16 S RNA was also studied. Under the experimental conditions used, four proteins, i.e., S4, S7, S8, and S 15, were sufficient to maintain the RNP particles in the form of intact 30 S subunits (Vasiliev et al., 1977a). Even in the complex of 16 S RNA with S4 alone, an appreciable proportion (about 20%) of the particles strongly resemble 30 S subunits (Vasiliev et al., 1977b) (Figure 4).

From these observations it was concluded that the major morphological peculiarities of the 30 S subunits are inherent in the very structure of 16 S RNA. This was also convincingly demonstrated by experiments with free 16 S RNA. Under special conditions that favored compact binding, the 16 S RNA acquire a V- or V-like conformation, the dimensions of which are very similar to the 30 S subunit (Figures 4 and 5b) (Vasiliev et al., 1978). As was concluded above, ribosomal proteins change the rRNA conformation during reconstitution of ribosomal subunits and maintain it in a compact state under the "physiological" conditions. Taking into account the results of Vasiliev and his colleagues (Vasiliev et al., 1977a,b, 1978) it must be admitted that alterations in rRNA structure proceed with the overall mode of folding of this chain into a V(Y)-like structure being retained.

The next step in the EM study of rRNA spatial organization in ribosomes is to identify rRNA regions located in certain morphological parts of subunits. The immuno-EM technique may provide particularly useful data here.

In some special cases the antibodies specific to modified RNA bases can be used for localization of rRNA regions, as has been done by Politz and Glitz (1977) in the case of two N 6,N6-dimethyladenosine residues of the 16 S RNA (Figure 5a).

A more general approach has been recently developed by Shatsky et al. (1979). It is based on the covalent binding of low-molecular-weight ligands containing the residue of phenyl-iJ-D-lactoside hapten to certain points of RNA (Shatsky et al., 1978) and the localization of the binding site of the antibody specific to this hapten by EM. The 3' end of the 16 S RNA has been localized on the 30 S subunit with rather high accuracy using this approach (Shatsky et al., 1979) (Figure 5a).

104

" ,

({

1 ... 1 ... , .. -... , ,

I 2 I \ I --~

a

A. A. Bogdanov et al.

b FIGURE 5. (a) Location of the 3' end (circle 1) (Shatsky et al., 1979) and Am! Am! sequence (circle 2) (Politz and Glitz, 1977) of the 16 S RNA in the 30 S subunit model of Vasiliev (1974) (see text). (b) Proposed location of the 16 S RNA molecule (cross-hatched area) and the S4-RNA (shaded area) in the 30 S subunit model of Vasiliev (1974). [Presented on the basis of the data from Vasiliev et al. (1978) and from Garrett et al. (1977), respectively.]

The study of rRNA topography in ribosomes is clearly at an early stage of development. However, the great significance of the rRNA spatial arrangement in ribosome subunits in understanding of overall ribosome structure and functioning, on the one hand, and recent appearance of new physical and biochemical techniques in ribosomology, on the other hand, allow one to hope that this problem will be solved in the not too distant future.

ACKNOWLEDGMENTS

We thank Professor A. S. Spirin and Professor R. A. Zimmermann for their critical reading of the manuscript and stimulating discussion of unpublished data. We also express our gratitude to Professor R. A. Zimmermann for his help in preparation of the manuscript. We are grateful to Dr. V. D. Vasiliev for generously providing us with electron micrographs. We are greatly indebted to Dr. R. Brimacombe, Dr. C. R. Cantor, Dr. H. Noller, and Dr. B. Ehresmann for sending us preprints describing their unpublished work.

Structure and Functions of Ribosomal RNA

8. ADDENDUM: A PRELIMINARY MODEL FOR THE SECONDARY STRUCTURE OF 16 S RIBOSOMAL RNA*

105

Significant progress was recently made on the elucidation of the secondary and tertiary structure of the 16 S RNA of E. coli.

First, Ross and Brimacombe (1979) have identified the greater part of the sequences in the 16 S RNA, which form double-stranded structures. The approach they used involves digestion of RNA with the single-stranded specific nuclease SI and separation of the products on a two-dimensional gel electrophoresis system, the first gel dimension being run under nondenaturating conditions and the second under denaturating conditions. RNA sequences that participated in the formation of paired segments become dissociated and appear as pairs or families of fragments in the second dimension. After determination of the nucleotide sequences of these fragments the secondary structure of more than half the 16 S RNA molecule has been elucidated. As well as numerous short double-stranded fragments formed with neighboring RNA segments, two long-range interacting regions of the 16 S RNA have been discovered.

Second, intramolecular cross-linking of the 16 S RNA has been studied by means of the double-stranded specific reagent hydroxymethyltrimethylpsora len both in the free state (Wollenzien et al., 1979) and within the 30 S ribosomal subunits (Thammana et al .. 1979). After rigorous denaturation of the cross-linked 16 S RNA, large loops ranging in size from 200 bases to essentially fully circular molecules have been detected in unfolded RNA chains by electron-microscopic examination. Determination of the absolute orientation of these loops in the 16 S RNA molecules permits identification of RNA segments, involved in the long-range interactions, with an accuracy of between ± 25 and ± 50 bases. It is important to note that most of the cross-links discovered in the free 16 S RNA have also been found in the RNA within the 30 S particle although the reactivity of the 16 S RNA in the subunit was much less than that of the free RNA (Thammana et al., 1979).

These data are sufficient to permit us to propose a preliminary model of the secondary structure of the 16 S RNA, which also describes tertiary interactions within its molecule (Figure 6). It is based mainly on data obtained by Ross and Brimacombe (1979) using RNA solutions containing Mg2+ ions ("magnesium fragments"). The structure of several short hairpin loops has been proposed on the basis of the principle of the maximum base pairing.

The tentative model presented in Figure 6 is in a good agreement with

*By E. A. Skripkin and A. A. Bogdanov.


FIGURE 6. A preliminary model for the secondary structure of the 16 S ribosomal RNA from E. coli. [Model based mainly on data of Ross and Brimacombe (1979)-see text.] The model is presented in the form of three segments (a, b, and c) that correspond to three RNP domains in the 30 S particle (see the text). Arrows indicate sites of cleavage of the 16 S RNA by RNase TI (within 30 S subunits) (Santer and Santer, 1972; Santer and Shane, 1977), nuclease SI (Ross and Brimacombe, 1979), and cobra venom nuclease (within 30 S subunits) (S. K. Vasilenko, personal commumcation; Carbon et al., 1979). K, kethoxal-reactive guanine residues of the 16 S RNA in active 30 S subunits (Herr et al., 1979). PS, complementary segments in the 16 S RNA areas cross-linked by uv irradiation of the complex of the 30 S subunit with psoralen (Thammana et al., 1979). S7, nucleotide residues, which can be covalently cross-linked with protein S7 by uv irradiation (Zwieb and Brimacombe, 1979). Oligo, RNA sequences accessible for complex formation with complementary synthetic oligodeoxyribonucleotides (Skripkin et al., 1980; E. A. Skripkin and E. Krynetsky, unpublished results). Sequences that can be released from compact 30 S subunits with mild RNase A or TI treatment are indicated by solid lines with arrows.


FIGURE 6. (Continued)

most of the experimental data discussed in this chapter. For example, one can see the highly structured regions (positions 70-570, 570- 870, and 980- 1320) corresponding to the three RNP domains described above in detail. Several interdomain interactions can also be seen. It is interesting that the 3'-end segment of the third domain appears to interact with the 3' end of the second domain, and both of them are in rather close proximity of the 3' end of the first domain.

This model is also consistent with most of data on the chemical modification and nuclease digestion of the 16 S RNA in the 30 S subunit: most guan-


ine residues modified with single-stranded specific reagent and most of the cleavage sites produced with single-stranded specific nucleases are located outside the double-helical segments of RNA. At the same time, the cuts produced with cobra venom nuclease, which is specific to highly ordered RNA segments, are almost exclusively inside or near the ends of the double-stranded regions. It is interesting that uridine residue number 1239, which can be cross-linked with protein S7 by uv irradiation (Moller et al., 1978; Zwieb and Brimacombe, 1979) appear in the single-stranded segments of the third domain. The 16 S RNA sequences 787-792 and 816-821, which are essential for association of 30 Sand 50 S subunits and protected in 70 S subunits from modification with kethoxal by 50 S subunits, are also in single-stranded form. It is in agreement with the suggestion that this region of 16 S RNA can be involved in base pairing with the corresponding complementary regions of the 23 S RNA (Herr and Noller, 1979; Herr et al., 1979).

The data on complementary binding of synthetic oligonucleotides (Skripkin et aI., 1980) and the release of oligonucleotides from the surface of the 30 S subunit (N. L. Teterina, personal communication) demonstrating the singlestranded conformation of the 5'-end proximal area and of several 3'-end proximal regions of the 16 S RNA are also in agreement with the proposed model.

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Structure and Functions of Ribosomal RNA III

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Chapter 3

Aspects of the Role of the Endoplasmic Reticulum in Protein Synthesis

Asbjorn M. Svardal and Ian F. Pryrne Cell Biology Research Group Department of Biochemistry The Preclinical Institute University of Bergen Arstadveien 19,5000 Bergen, Norway

1. INTRODUCTION

The endoplasmic reticulum is an organelle discovered using the electron microscope (EM) and subsequently found to be present in nearly all cells of higher plants and animals (Claude, 1970; Palade, 1955b, 1956; Palade and Porter, 1954; Porter, 1953; Porter et al., 1945; Porter and Palade, 1957). Ever since 1945, when Porter and co-workers first noticed this organelle, there has been a growing interest concerning the elucidation of the role of the endoplasmic reticulum in cellular processes, such as drug metabolism and control of protein synthesis. Eukaryotic cells specialized for the secretion of proteins contain a large fraction of their ribosomes bound to membranes of the endoplasmic reticulum (Birbeck and Mercer, 1961; Palade, 1956). Secretory polypeptides synthesized by bound ribosomes are vectorially transferred to the lumen of endoplasmic reticulum cisternae as a first step in the process leading to their extracellular discharge (Redman et al., 1966; Redman and Sabatini, 1966).

Other polypeptides destined for transport to inner compartments of other cellular organelles, e.g., lysosomes, Golgi complex, peroxisomes, and mitochondria, and those that form the tightly bound class of integral membrane proteins or are deposited asymmetrically toward the noncytoplasmic face of membranes, together with some viral proteins, may also be synthesized on bound polysomes. On the basis of the available information (mostly morphological), it has been suggested that a basic general secretory pathway exists in all

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118 Asbjl)rn M. Svardal and Ian F. Pryme

eukaryotic cells (Palade, 1975). This pathway can be divided into six successive steps: (1) synthesis of proteins on polysomes attached to the endoplasmic reticulum, i.e., membrane-bound polysomes; (2) segregation of the synthesized proteins in the cisternal space of the rough endoplasmic reticulum; (3) intracellular transport through the rough and smooth endoplasmic reticulum to the Golgi apparatus; (4) concentration of secretory products in condensing vacuoles formed by the Golgi apparatus; (5) intracellular storage of secretory proteins in secretion granules, which represent the ultimate products of the concentration process occurring in the condensing vacuoles; and (6) discharge of the secretory proteins from the cell following fusion of the membrane of these secretory granules with the plasmalemma.

The current situation evokes the necessity of obtaining detailed and comprehensive data on the chemistry and function of the different membranes of the secretory pathway and on their interactions. With this type of information our understanding of the basic organization and function of the ER in eukaryotic cells could be further advanced. This review summarizes the present state of knowledge concerning the role of the endoplasmic reticulum in protein synthesis.

2. MEMBRANES OF THE ENDOPLASMIC RETICULUM

The early electron micrographs of cells (Porter, 1953; Porter et al., 1945) displayed a complex membrane network permeating much of the cell body, but that did not extend into the clear ectoplasmic zone at the periphery of intact cells. This observation of the organelle led to the term endoplasmic reticulum. With the development of suitable techniques for the study of cells in ultrathin sections (Palade and Porter, 1954), the endoplasmic reticulum was shown to be comprised of tubular, vesicular, and cisternal structures. As the resolution of the electron micrographs improved, it was noted that in certain areas the outer surfaces of the membranes of the reticulum were often studded with small dense granules of uniform size (Palade, 1955b). These granules are now known as ribosomes. It was thus recognized that the cytoplasmic membranes can be either of the granular (rough-RER) or agranular (smooth-SER) type.

Watson (1955) observed that the agranular and the granular reticulum and the nuclear envelope were interconnected. The Golgi complex, being agranular, was originally included in the agranular part of endoplasmic reticulum. Even though the SER and Golgi apparatus are intimately related (Claude, 1970; Lane and Swales, 1976), the Golgi complex is now considered a separate system (Fawcett, 1969).

The reticulum may have a characteristic or random intracellular distri-

The ER and Protein Synthesis 119

bution and be either abundant or scarce (Fawcett, 1969; Threadgold, 1976). There is therefore very great diversity in the form and ultrastructure of the endoplasmic reticulum, depending on cell type, phase of cell cycle, stage of growth and/or differentiation, and function. For example, in the cells of the adrenal gland and the interstitial cells of the testis, the SER is believed to be involved in the synthesis of steroid hormones, whereas in muscle cells the SER is called sarcoplasmic reticulum and is involved in muscle contraction and relaxation (Threadgold, 1976). The cells having the most extensively developed RER are pancreatic exocrine cells, plasma cells, and liver cells, and these are specialized to perform primarily one function, i.e., to synthesize and secrete a limited number of proteins (Birbeck and Mercer, 1961; Palade, 1956; Potter, 1967; Rolleston, 1974).

The ER is also characterized by its property of possessing a high degree of adaptation to varied physiological and experimental conditions. For example, an extensive reorganization of the liver ER takes place in the newborn rat (Dallner et a/., 1966a,b) reflecting an adaption to extrauterine life. The liver ER also responds to phenobarbital, carcinogens and other xenobiotics by increasing its volume and surface area (Staubli et al., 1969). In addition, changes occur with respect to the specific activity of several enzymes (Orrenius et al., 1965). After ether or halothane anesthesia of rats there occurs a proliferation of SER and an induction of microsomal drug-metabolizing enzymes (Ross and Cardell, 1978).

The adaptability and functional importance of this organelle has resulted in it being the object of quite considerable attention, ranging from isolation and subfractionation to compositional and structural studies. From such studies it became apparent that the ER membranes can act as selective barriers by which the entrance of certain materials into the inner space (lumen) of the ER is controlled. The importance of this is exemplified by the process of secretion whereby specific molecules (e.g., insulin, albumin, immunoglobin) are released from the cell.

It appears that the membranes of the ER can be instrumental in regulatory mechanisms and that the system as a whole should bear a high degree of adaptability in order to meet these requirements.

2.1. The Composition and Structure of the Endoplasmic Reticulum Membranes

Contamination of the ER fraction with other membranes (e.g., nuclear, plasma, mitochondrial, lysosomal, and peroxisomal membranes) (Depierre and Dallner, 1976) has made it difficult to perform an exact chemical analysis of the ER. Nevertheless, progress has been made and a characteristic assay is given in Table 1.

120 Asbjilrn M. Svardal and Ian F. Pryme

Table I Percent Composition of Stripped

Endoplasmic Reticulum Membranes·

Protein Lipid Cholesterol RNA Sialic acid

"After Threadgold (1976).

55 (± 10%) 40 (± 10%) 4.5 (± 1.5%) Trace Trace

About 70% of the total lipid consists of phospholipids of which choline phosphoglyceride and ethanolamine phosphoglyceride account for the major part (Meldolesi and Cova, 1972; Meldolesi et al., 1971a-c). The functional importance of the lipids is not entirely clear, but it is recognized that they exert significant effects on the activity of ER enzymes and are important in establishing many of the fundamental properties of the membranes (Ernster and Orrenius, 1973; Jones and Mills, 1973; Laitinen et al., 1975). The high protein content is accounted for by a combination of structural proteins and a high concentration of enzymes (Jones and Mills, 1973). The protein/phospholipid ratio of ER membranes is variable in preparations obtained from different cell types, and this may reflect differences in metabolic activity (Geuze et al., 1977). The RNA found in the membranes may represent contaminating ribosomes, absorbed tRNA, or "membrane RNA" (Pitot, 1964; Tata, 1967).

Much attention has been devoted to the elucidation of differences in properties between rough and smooth microsomes. A comparison of these is shown in Table II.

Compared to plasma membranes the ER membranes are considerably thinner (8-10 nm and 5 nm, respectively) (Geuze et al., 1977). They also differ on two other points. First, the ER membrane is considered to be symmetrical, i.e., the inner and outer protein components are of equal width, whereas the plasma membrane is asymmetrical. Second, the central lipid component of ER membranes, being crossed by dense septa displays a globular structure, the opposite ends of which are fused to the inner and outer protein components; in plasma membranes this globular pattern is not normally observed (Geuze et al., 1977; Threadgold, 1976). It has been speculated that these lipoprotein globules could become confluent to form a simple bimolecular leaflet, spreading out over large areas (Geuze et al., 1977). Such phase changes could have physiological significance, allowing for dynamic changes in the permeability of the membranes.

The ER and Protein Synthesis

TableD Comparison of Rough and Smooth Microsomes

Analysis RNA content Cholesterol Dolichol phosphate Galactose, sialic acid

Property

Net negative surface charge Affinity for monovalent cations Equilibrium density Tendency to aggregate Appearance of electron transport enzymes after birth or

induction Membrane-bound ribosomes 60 S subunit binding sites Ribophorins I and II Protein-synthetic ability Presence of secretory polypeptides after labeling in vivo

Rough microsomes

Very high Low High Low High High High Low

Soon Present Very many Present Good Early

2.2. Functional Aspects of the Endoplasmic Reticulum

121

Smooth microsomes

Negligible High Low High Low Low Low High

Later Absent Few Absent Poor Later

In cells that secrete considerable amounts of protein, such as plasma cells, liver cells, and pancreatic cells, there is relatively more RER than SER (Palade, 1956, 1975). Although the RER is clearly specialized for protein synthesis, there are at present little or no data suggesting other qualitative functional differences between rough and smooth ER (Depierre and Dallner, 1975). For example, no enzyme activity associated with the membrane of the ER has been shown to be localized exclusively in rough or smooth microsomes, and quantitatively the enzymatic pattern of these two subfractions (with the exception of activities associated with ribosomes) is almost identical (Depierre and Dallner, 1975).

The RERjSER ratio varies considerably from cell type to cell type, and this is related to the particular cell's primary function (Threadgold, 1976). In addition, there are many examples of alteration in the function (and the RERj SER ratio) of the ER, caused by genetical changes or dietary or various other environmental conditions (Siekevitz, 1976). Consequently it is to be emphasized that the membranes of the endoplasmic reticulum are highly dynamically organized (Ernster and Orrenius, 1973).

122 Asbjijrn M. Svardal and Ian F. Pryme

To illustrate this dynamic organization the variety of enzymes associated with the ER membranes of liver cells can be considered. These include oxidoreductases, transferases, and hydrolases (Ernster and Orrenius, 1973). These ER enzyme systems playa major role in cholesterol biosynthesis (Dugan and Porter, 1976; Geuze et al., 1977; Goad, 1970), phospholipid metabolism (Higgins, 1976; Jungawala and Dawson, 1970; Lands and Crawford, 1976), and in the biosynthesis of lipoproteins (Marsh, 1971), glycoproteins (Molnar, 1976), and glycolipids (Lennarz and Scher, 1972). Enzyme systems in liver ER also play other important roles, e.g., glycogen metabolism (Cardell, 1977).

Two enzyme systems associated with the ER are of particular interest. The first system is composed of at least two protein components-cytochrome P-450 and NADH-cytochrome P-450 reductase-and it uses electrons from NADPH in the oxidation of xenobiotics, steroids, and fatty acids (Conney, 1967).

The second electron-transport chain is composed of at least three protein components-cytochrome bs, NADH-cytochrome bs reductase, and fatty acylCoA desaturase-and it uses electrons from NADH in the desaturation of fatty acids (Oshino and Omura, 1973; Strittmatter et al., 1974).

Because of the central position of these two multienzyme complexes in liver function, much work has recently been devoted to the question of whether the enzymes are distributed at random along the ER, or whether particular enzymes are concentrated in different regions of the reticulum, with a consequent functional specialization of such regions. As far as the rough and smooth segments of the reticulum are concerned, it now appears that these are closely similar in terms of relative enzyme composition (Depierre and Dallner, 1975). An important exception, however, is the case of an enzyme or an enzyme system in the stage of active induction, in which there may be a disproportion in the enzyme contents of the two segments. Thus it is found, for example, that during the drug-induced synthesis of the hydroxylating enzyme system, (the NADPH-linked electron-transport system) the concentration of its enzyme components is higher in the rough than in the smooth microsomes during an early stage of induction (Orrenius, 1965; see Table II), whereas the converse relationship holds at a later stage (Orrenius, 1965; Remmer and Merker, 1963). Confirmation of this indication of functional heterogeneity of different parts of the endoplasmic reticulum, based on an uneven distribution of enzymes associated with the membrane, has been one of the chief aims of subfractionation of microsomes in recent years.

As mentioned above, an extensive reorganization of liver ER can take place, for example, in the newborn rat. A striking enzymatic change is increased NADPH- and NADH-linked electron-transport activities (Dallner et al., 1966a,b). In all cases, the increase is caused by enhanced rates of enzyme synthesis, as indicated by turnover measurements, and is accompanied by a gross reorganization of the ER, as demonstrated by the appearance of


smooth-surfaced profiles resulting probably from an outgrowth of new membranes from the rough-surfaced segments of the ER.

It is now well established that administration of drugs or other substrates of the monooxygenase system induces a greatly increased catalytic capacity of this system in liver microsomes. The induction process leads to a selective and parallel increase in the levels of the flavoprotein and cytochrome components of the monooxygenase system, leaving other microsomal enzyme activities largely unaffected (Ernster and Orrenius, 1965; Orrenius et at., 1965). It involves increased rates of both protein and RNA synthesis, as indicated by its sensitivity to puromycin and actinomycin D (Conney and Gilman, 1963; Gelboin and Blackburn, 1963; Orrenius et at., 1965). It is likely that this phenomenon is a reflection of a posttranscriptional (Tomkins et aI., 1969, 1972) or transcriptional (Nebert and Gelboin, 1970) control of induced enzyme synthesis.

A striking phenomenon accompanying drug-induced enzyme synthesis is the proliferation of ER membranes (Feuer et at., 1977; Grasso et at., 1977; Higgins, 1976; Orrenius et aI., 1965). From the available evidence it appears that the induced enzyme synthesis begins in the rough-surfaced areas of the ER, but leads eventually to an accumulation of smooth-surfaced profiles, rich in components of the monooxygenase system (Ernster and Orrenius, 1965; Orrenius, 1965). Thus the synthesis of new membranes seems to be geared to the formation of new enzymes. The mechanism for this synthesis is not clear, but it may involve an outgrowth and budding off of smooth-surfaced membrane profiles from the rough-surfaced ER at the sites where the enzyme synthesis takes place (Ernster and Orrenius, 1973). Some caution is warranted, however, before completely accepting this conclusion, because evidence has been presented that enzymes required for phospholipid synthesis reside in SER membranes, and these enzymes increase in amount in response to phenobarbital treatment (Higgins, 1974, 1976; Higgins and Barrnett, 1972). Thus SER is capable of forming at least the phospholipid component of its membranes without direct participation of RER. How the protein complexes are formed and inserted into SER membranes is not entirely clear.

It has been stated that images showing continuity of SER with RER are more abundant during SER induction than in unstimulated cells (Jones and Mills, 1973). Morphological changes in RER during SER induction have been reported by several investigators (Bolender and Weibel, 1973; Dallner et at., 1966a,b; Staubli et at., 1969). Increased numbers of "groups" of ribosomes in profiles of RER during SER induction have been described, and such groups may represent polysomes that are active in the synthesis of membrane components. In addition, regions of RER membranes with no or few ribosomes occur with increased frequency during periods of SER induction. It has been suggested that such areas may indicate the location of products of recent membrane synthesis (Dallner et at., 1966a).


It seems appropriate at this point to state in broad terms a general concept for the function of SER that is compatible with most of the available data. Although exceptions have occurred and others may yet be found, the following concepts apply to most cell types investigated thus far and offer guidelines for interpreting the function of SER in all cells (Cardell, 1977). An inducer of SER interacts with a cell and stimulates the formation of all types of RNA. The newly formed mRNA directs the synthesis of the appropriate proteins (structural and constitutive enzymes of the ER membranes) at the level of ribosomes attached to RER. The proteins are inserted into forming ER structures, resulting in the production of additional ER membranes containing specific components. The newly synthesized components appear in SER membranes after transformation of RER to SER. By such a mechanism the cell responds to an inducer by forming new SER membranes that are quantitatively different from the old membranes. As long as the inducer stimulates the cell, the levels of SER remain elevated. If the inducer is removed or its concentration in the blood is lowered, the production of SER decreases and the rate of degradation of its membranes increases (Depierre and Ernster, 1976). A cell utilizing this mechanism is able to control the quantity and quality of its SER membranes in response to fluctuating levels of inducing substances. This makes the endoplasmic reticulum a highly dynamic structure with a correspondingly high adaptive capacity toward various physiological and experimental conditions (Cardell, 1977; Ernster and Orrenius, 1973).

Carcinogens bind to a variety of macromolecules including DNA, RNA, and especially proteins (Arcos and Argus, 1974; Berenblum, 1974), and also bind to components in different subcellular fractions, e.g., nuclei, mitochondria, microsomes, and cytosol. The binding of carcinogens to specific macromolecules and organelles may be of relevance for transport, metabolic activation, detoxification, and other functions. Direct binding of carcinogens occurs at sites in the ER that are in close proximity to, or are perhaps identical with, those known to bind steroids, phenolic antioxidants, inhibitors of microsomal hydroxylases, and retinol. There is strong evidence suggesting that carcinogens must bind to ER sites that are regulated or modified by the influence of steroids or drugs (Kisilevsky and Weiler, 1976; Litwack and Morey, 1970; Sunshine et al., 1971; Williams and Rabin, 1969, 1971; Williams et aI., 1972). Steroid hormones are not only involved in carcinogenjER interactions (Arcos et al., 1975; Huggins et al., 1962), but also in binding ribosomes to membranes (Rabin et al., 1970; Sunshine et al., 1971; Williams and Rabin, 1971). The events occurring in ER membranes during malignant neoplasia have been reviewed by Apffel (1978).

Morphological alteration and biochemical changes in the ER in cancer cells and during the process of carcinogenesis are well documented (Apffel, 1978; Franke, 1977; Parkin and Brunning, 1978; Wallach, 1975). Both ultra-


structural and biochemical studies have indicated that there occurs a detachment of ribosomes from the RER, whereas the number of free ribosomes appearing scattered in the cytoplasm increases. This observation is not only a feature of changes occurring during the process of carcinogenesis but is also typical of both spontaneous animal tumors (Dalton et al., 1961; Kajikawa, 1959; Shubin and Tikhomirova, 1959) and experimentally induced tumors in mice [ascites tumors: Daniels et al. (1968), Probst (1962); hepatoma: Trotter (1963); breast adenocarcinoma: Shubin and Tikhomirova (1959); leukemia: Graffi et al. (1959); plasmacytoma: Dalton et al. (1961)], in rats [hepatoma: Moyer et al. (1970), Porter and Bruni (1959), Swoboda and Higginson (1968); rhabdomyoblastoma: Chentsov (1960); sarcoma: Shubin and Tikhomirova (1959)], in hamsters [reticulosarcoma: Vogel (1962); sarcoma: Jacubov et al. (1975)], and in monkey kidney cells [Merkow et al. (1970)]. Similar findings have also been reported in a wide range of human tumors [bladder carcinoma: Bini and VitaliMazza (1960); chronic lymphoid leukemia, Hodgkin's, and lymphosarcoma: Gigante et al. (1961); endometrial carcinoma: Wilsson, (1962); glioblastoma, medulloblastoma, and medulloepithelioma of the brain: Tani and Higashi (1975); HeLa cells: Epstein (1961); uveal sarcoma: Francois et al. (1959), Rabaey and Lagasse (1959)]. The actual significance ofthese findings is not yet understood, although it is interesting to note that a similar tendency is also seen in embryonal cells (Dallner et al., 1966a; Tedde, 1973; Tujlmura, 1958).

Ribosomes can be artifically removed from RER by a variety of treatments (e.g., low detergent concentrations, LiCI or KCl treatment). Under appropriate conditions the ribosomes will reassociate with stripped RER membranes. Although released ribosomes from tumor cells will bind to stripped RER from normal cells, such ribosomes will not attach to stripped ER from tumor cells (Berdinskikh et at., 1975; Gravela et al., 1975; Hochberg et al., 1972; Kisilevsky and Weiler, 1976; Moyer et al., 1970; Staubli and Loustalot, 1962; Villa-Trevino et aI., 1964). At present. it is uncertain whether these observations are a result of changes in composition/structure of ribosomes or ER membranes, or perhaps a combination of factors may be involved.

3. PROTEIN SYNTHESIS

3.1. Protein Synthesis by Free and Membrane-Bound Poly somes

Cytoplasmic ribosomes are present in many tissues, both free in the cytosol and attached to the endoplasmic reticulum. The percentage of the total ribosome population that is bound to the endoplasmic reticulum varies considerably from cell type to cell type (Shires et al., 1974). This distribution, however, is not a fixed parameter and can undergo substantial variation in the same


cell. A few examples where alterations in free/membrane-bound polysomes have been shown to occur illustrate this ability of cells to adjust to a change in conditions. Nonsecretory rabbit mammary gland was found to have 25% of total ribosomes bound to the endoplasmic reticulum, whereas 96 hr after prolactin stimulation the total increased to 70% (Gaye and Denamur, 1970). As most proteins secreted from the liver are synthesized on membrane-bound polysomes (Ganoza and Williams, 1969; Redman, 1968, 1969), whereas proteins for intracellular use are synthesized on free polysomes (Ganoza and Williams, 1969), one might expect to observe a shift of protein synthesis from export proteins to "cellular" proteins in regenerating rat liver. Despite much controversy during the past few years Loeb and Yeung (1978) have provided extremely good evidence indicating that such a shift does, in fact, occur. They showed that 48 hr after partial hepatectomy, when liver cell proliferation is about maximal, a striking difference in the distribution of ribosomes occurred, being manifested by an increase of about 40% in the proportion of free ribosomes. Based on mass of regenerating liver tissue, the absolute concentration of hepatic ribosomes rose to nearly double that in normal liver cells. These results clearly indicate that liver cells can utilize free and membrane-bound polysomes to varying extents, depending on the specific requirements of the tissues.

Chronic alcohol consumption also has an effect on the relative distribution of polysomes in rat liver within free and membrane-bound populations. The in vitro protein-synthesizing abilities of free and membrane-bound polysomes isolated from young rats after alcohol administration have been studied by Khawaja and Lindholm (1978). They found that there was a 125% increase in the amino acid incorporating ability of free polysomes, whereas there was a 25% reduction in that of membrane-bound polysomes. Alcoholic hepatomegaly in the liver has been attributed to an accumulation of secretory products in the tissues arising because of impaired transport (Baraona et aI., 1975). Khawaja and Lindholm (1978) interpret their results to indicate that the enhanced protein synthesis by free ribosomes is some form of compensatory phenomenon, perhaps a result of decreased synthesis of export proteins or because of ineffective transport mechanisms. Khawaja et al. (1978) have also demonstrated a selective inhibition of the protein synthesizing ability of cerebral membranebound ribosomes following the administration of alcohol. Chu et al. (1978) have investigated the protein synthesizing activities of free and membranebound ribosomes during the development of chick liver. Using in vitro studies, they were able to show that the activity in membrane-bound polysomes increased twofold between days 8 and 18 of development, the total ribosome concentration, however, remaining unchanged. The degree of segregation of ribosomes into free and membrane-bound classes thus depends to a large degree on the specific characteristics of the cell in question and the actual func-


tions (e.g., controlled by physiological and/or biochemical parameters), which a population of cells are engaged in at any particular point in time.

Substantial evidence has accumulated suggesting that in addition to synthesizing secretory proteins, membrane-bound polysomes are engaged in the synthesis of structural proteins of the endoplasmic reticulum (e.g., Ragnotti et al., 1969; Rolleston, 1974). Recent investigations, however, indicate that the situation may be a good deal more complex than was first anticipated (see Section 3.3).

It should be emphasized that a secretory protein normally synthesized on membrane-bound polysomes, may, under abnormal conditions, be synthesized on free polysomes. Thus in a hepatoma cell line where albumin was shown to be synthesized, but not secreted, synthesis was shown to be greater in the free than in the membrane-bound polysome fraction (5:1 per mg RNA) (Uenoyama and Ono, 1972).

3.2. Secreted Proteins May Be Synthesized Solely on Membrane-Bound Polyribosomes

The first direct evidence that secretory proteins are synthesized by membrane-bound ribosomes came from in vivo labeling experiments (Siekevitz and Palade, 1960). Following injection of guinea pigs with [I4C] leucine, the specific activity of nascent a-chymotrypsinogen in the pancreas after 1 min was found to be greatest in the bound ribosomes fraction. This observation was the basis for enumerable studies which have led us to our present knowledge concerning the understanding of the process of secretion.

Although the pancreas was the focus of attention in the early days, other studies have concerned proteins secreted by a variety of different organs and cells. In all cases, secreted proteins have been found to be synthesized primarily in the bound polyribosome fraction. Some examples are the following: immunoglobulin synthesized in plasma cells (Choi et al., 1971; Cioli and Lennox, 1973; Pryme et al., 1973; Sherr and Uhr, 1970); serum protein in the rat (Ganoza and Williams, 1969; Redman, 1968, 1969); albumin in rat liver (Hicks et al., 1969; Takagi and Ogata, 1968; Takagi et al., 1969); thyroglobulin in porcine thyroid (Di Lauro et aI., 1975) and sheep thyroid (V assart, 1972); collagen in chick embryo connective tissue (Diegelmann et al., 1973); rat growth hormone (Bancroft et al., 1973); kidney glycoproteins (Priestley et al., 1969); mammary gland milk protein (Gaye and Denamur, 1970; Gaye et al., 1973a,b); egg white proteins in the oviduct of avian species (O'Malley et al., 1975); invertase in yeast (Holley et al., 1973); and cellulase, an extracellular cell-wall enzyme in higher plants (Verma et al., 1975).

The reason for the necessity of translating mRNAs that code for secretory proteins in membrane-bound polysomes rather than on free polysomes has now

128 Asbjllrn M. Svardal and Ian F. Pryme

become apparent. All the secretory proteins mentioned above have a common destination, i.e., they are finally released from the cell to the external environment. Before this event can occur, however, they have to cross a membrane barrier consisting of a hydrophobic lipid bilayer. There is now considerable evidence to support the original suggestion by Blobel and Sabatini (1971), showing that secretory proteins are synthesized with a characteristic peptide at the N-terminus of the molecule. This peptide has been termed the "signal" peptide and is metabolically short-lived, being present only on incomplete nascent chains. Its function is merely to direct the growing nascent chain from an initial cytoplasmic site through the hydrophobic lipid bilayer of the endoplasmic reticulum into the lumen of the membrane, thus accomplishing the first step in ensuring a final secretion of the completed molecule from the cell. It seems to be becoming increasingly apparent that not only secretory proteins, but other proteins as well that have to traverse the endoplasmic reticulum, also possess a similar signal sequence. Aspects of the signal hypothesis are discussed further in Section 4.2.

Because glycosylating enzymes are membrane-bound, it was originally thought that translation of secretory protein mRNAs must occur in association with membranes. However, the fact that secretory proteins need not necessarily be glycosylated, yet are efficiently secreted, has led Melchers (quoted by Campbell and Blobel, 1976) to conclude that the role of glycosylation in the secretion of proteins is not yet fully understood.

Wherever tested, the synthesis of secretory proteins in normally functioning tissue has always been found to occur on membrane-bound polysomes, although the situation in tissue that cannot be regarded as normal may be otherwise (see Section 3.1).

3.3. Proteins Synthesized on Membrane-Bound and/or Free Poly somes

HeLa cells, although they do not secrete more than 2% of the protein they synthesize, have 15% of their ribosomal population attached to the endoplasmic reticulum (Rosbash and Penman, 1971a). This discovery suggested that not all membrane-bound ribosomes are involved in the synthesis of secretory protein and much evidence is now available showing that in addition to an engagement in the synthesis of secretory proteins membrane-bound polysomes also synthesise, for example, soluble tissue proteins and proteins to be directed into various membrane-containing organelles.

It was originally suggested that soluble tissue proteins are synthesized exclusively on free ribosomes (Ganoza and Williams, 1969). Although this is probably the case for the majority of such proteins there are several well-documented exceptions. Both tyrosine aminotransferase (Chuah and Oliver, 1971) and serine dehydratase (Ikehara and Pitot, 1973), which are found in the


cytosolic compartment of the cell, are synthesized on membrane-bound polysomes. Linder et al. (1977) compared the in vitro synthesis of ferritin by rat liver free and membrane-bound polysomes and found that both had a similar capacity to synthesize ferritin. Eukaryotic cell ribosomal proteins are synthesized in the cytoplasm (Craig and Perry, 1971; Heady and McConkey, 1970) before they are transferred to the nucleus, where they combine with precursor ribosomal RNA. In order to facilitate transport into the nucleus, it would be reasonable to assume that ribosomal proteins may be synthesized on membrane-bound polysomes attached to the nuclear envelope. Mager and Planta (1975) investigated this process in yeast and found that polysomes engaged in the synthesis of ribosomal proteins were, in fact, distributed randomly in both free and membrane-bound polysome popUlations. Approximately 25% of the soluble protein in adult mammalian brain is tubulin, which is a major component of microtubules. Floor et al. (1976) have shown that tubulin synthesis occurs to the same relative extent on both free and membrane-bound polysomes. In rabbit reticulocytes at least 90% of the ribosomes are free in the cytoplasm, while the remainder are found in loose association with the plasma membrane (Lodish and Small, 1975). Bulova and Burkea (1970) provided evidence that the bound class of polysomes was essentially involved in the synthesis of nonglobin proteins. Other studies, however, have shown that alphaand beta-globin mRNA is translated primarily in the membrane-bound fraction (Lodish and Small, 1975; Morrison and Lingrel, 1975; Woodward et al., 1973).

A visualization of the possible positioning of the protein components of the endoplasmic reticulum is shown schematically in Figure 1 (Sabatini and Kreibich, 1976). Because the passage of a completed and folded protein across membranes is not plausible thermodynamically, Sabatini and Kreibich (1976) postulated that integral and peripheral proteins of the luminal faces, as well as transmembrane proteins of all membranes are synthesized by bound polysomes exclusively, as only these can effect the vectorial discharge of polypeptide chains across the phospholipid barrier directly into the lumen of the cisternae of the endoplasmic reticulum. They also presented a model (Figure 2) that shows how this may occur (Sabatini and Kreibich, 1976). There is considerable evidence to support this hypothesis. A prominent role of bound ribosomes in the synthesis of endoplasmic reticulum membrane proteins is supported by the finding that a large fraction of puromycin-released nascent polypeptide chains from membrane-bound ribosomes is not directly discharged into the lumen of the microsomal vesicles, but remains associated with the endoplasmic reticulum membranes (Sauer and Burrow, 1972). The C-terminal ends of these are partially exposed on the outer face of the vesicles and were found to be accessible to added proteases (Sabatini et al., 1972; Kreibich and Sabatini, 1973).

The intracellular site of synthesis of membrane proteins has been the sub-


cytoplasmic side

luminal side

FIGURE 1. Possible dispositions of ER membrane proteins. (a) Luminal protem, {OJ lUmmalface peripheral membrane protein, (c) luminal-face integral membrane protein, (d) transmembrane protein, (e) cytoplasmic-face integral membrane protein, (f) cytoplasmic-face peripheral membrane protein, (g) intramembranous protein (hypothetical). [Reprinted from Sabatini and Kreibich (1976) with permission of the authors and Plenum Publishing Corporation, New York.]

rie-~COOHNH2

~.~.j ~

to" ________ ''''''0'' ."',',, f6' permanent residents p -----II.~ of the ER

NH2 ------... diverted to other membrane bound compartments

FIGURE 2. Possible fates of newly synthesized polypeptides transferred to the luminal side of the endoplasmic reticulum. [Reprinted from Sabatini and Kreibich (1976) with permission of the authors and Plenum Publishing Corporation, New York.]


ject of much controversy. A number of reasons, both technical and physiological, may explain why this may be so. For example, there is evidence that in some cases the formation of initiation complexes, destined to become membrane-bound polysomes, occurs in the cytoplasm (Blobel and Dobberstein, 1975b; Harrison et al., 1974a). Thus, although such initiation complexes would normally complete translation only in association with rough ER in vivo, any assay procedure involving the completion of translation in vitro would indicate that these polysomes are present in the free ribosome fraction.

Peripheral and integral proteins (Singer, 1974; Singer and Nicolson, 1972) of the cytoplasmic face of all membranes do not need to cross membrane barriers and thus can potentially be synthesized on either free or bound ribosomes (in the same compartment). Figure 3 shows schematically how this may take place (Sabatini and Kreibich, 1976). Synthesis on free ribosomes would imply incorporation of the proteins from the cell sap into the recipient membrane face (Bretscher, 1973; Lodish and Small, 1975). This, in turn, implies a specific recognition between the protein and the membrane to which it is destined. Such recognition is likely to be mediated by an integral protein in the

cytoplasmic

proteins

FIGURE 3. Possible fates of newly synthesized polypeptides released from membrane-bound polysomes (but not discharged into the lumen of the endoplasmic reticulum) or free polysomes. [Reprinted from Sabatini and Kreibich (1976) with permission of the authors and Plenum Publishing Corporation, New York.]


receiving membrane and the newly attached protein would then become a peripheral membrane protein (see Figure 1). Further incorporation of the protein deeper into the membrane framework may necessitate cleavage of the newly synthesized polypeptide chain by membrane-associated proteases such as that described for some erythrocyte membrane proteins (Lodish and Small, 1975). Rothman and Lenard (1977) have considered the possible methods for biosynthesis of membrane proteins (and lipids) and their ultimate insertion into membranes. Blobel (1978) has recently discussed some of the current problems concerning known, or possible mechanisms involved in the intracellular compartmentation of protein molecules after their release from ribosomes.

Parry (1978) envisages that membrane-bound polysomes may participate in the formation of membrane proteins in three different ways: (1) membrane polypeptides are incorporated into the luminal face of the endoplasmic reticulum and are then transferred by membrane flow to the plasma membrane; (2) membrane polypeptides appear on the "inside" of vesicles which, following transport to the plasma membrane, are incorporated into the membrane structure such that the proteins then appear on the "outside" of the plasma membrane; and (3) membrane polypeptides synthesized on membrane-bound polysomes are not luminally discharged but are instead released to a cytoplasmic pool, to which free polysomes may also contribute, and these would then be incorporated into the cytoplasmic side of the plasma membrane. One can thus imagine a form of compartmentalization of mRNA for membrane proteins: those destined to appear on the external face of the plasma membrane would be solely synthesized on membrane-bound polysomes and the polypeptides would be discharged through the endoplasmic reticulum (presumably requiring a signal sequence) to the luminal face, whilst those polypeptides to be incorporated into the internal, cytoplasmic face could be potentially synthesized on membrane-bound polysomes, but discharged to the cytoplasm (signal sequence not required), or on free polysomes.

The viral envelope of vesicular stomatitis virus contains two virus-specific proteins-the M protein, which is found on the cytoplasmic face of the plasma membrane of the infected host cell and later appears on the inside of the viral envelope; and a glycoprotein (G), which is located on the outer face of the plasma membrane of the host and subsequently forms the viral spikes. The G protein has recently been shown to be a transmembrane protein (see Figures 1 and 2), its carboxy terminus being exposed on the cytoplasmic face of the endoplasmic reticulum (Katz and Lodish, 1979). Based on the discussion in the previous paragraph one would predict that protein M could be synthesized on either membrane-bound or free polysomes (or both classes), whereas the synthesis of glycoprotein G should occur exclusively on membrane-bound polysomes. Evidence that this is, in fact, the true situation has been provided by Grubman et al. (1974, 1975), Morrison and Lodish (1975), and Toneguzzo


and Ghosh (1975). The synthesis of two species of membrane proteins in intact rabbit reticulocytes (Lodish 1973) that are ultimately localized on the cytoplasmic face of the plasma membrane has been studied by Lodish and Small (1975). These proteins are synthesized on free ribosomes (Lodish and Desalu, 1973) analogous to the situation described above for vesicular stomatitis virus cytoplasmic face membrane proteins. Sindbis virus consists of three proteins and a genomic RNA. Two glycoproteins (E1 and E2) are integral proteins of the viral membrane envelope, whereas the third, a capsid protein (C), is found in association with the viral 42 S RNA (Bonatti et al .. 1979). The 42 S RNA is processed to a 26 S species, which is almost exclusively translated in membrane-bound polysomes (Martire et al.. 1977; Wirth et al .. 1977) resulting in the production of a single protein. Following proteolysis the three proteins E1,

E2, and C are generated, the first two being integrated into the endoplasmic reticulum, and protein C apparently discharged into the cytosol (Bonatti et al .• 1979).

In studies on the biogenesis of plasmalemma glycoproteins, Bergeron et af. (1975) have studied the intracellular site of 5'-nucleotidase synthesis. Free and membrane-bound polysomes were isolated from mouse liver, the mRNA was extracted and injected into oocytes. Specific antiserum was used to determine the degree of synthesis of 5'-nucleotidase. The results showed that the membrane-bound polysome fraction contained approximately 17 times more mRNA coding for 5'-nucleotidase than in the free polysome fraction and suggested that either small amounts of the enzyme are synthesized on free polysomes in vivo or that the mRNA in transit from the nucleus to the site of translation in the rough endoplasmic reticulum arises artifactually in the free polysome fraction.

Ragnotti et af. (1969), using in vitro experiments, showed that the synthesis of the microsomal enzyme NADPH-cytochrome c reductase occurs on both free and membrane-bound polysomes. Lowe and Hallinan (1973), however, provided evidence that the enzyme was preferentially synthesized on free polysomes. Harano and Omura (1977a), using specific antibodies, were able to precipitate nascent polypeptide chains labeled in vivo both on free and membrane-bound polysomes, indicating that synthesis occurs on both sets of polysomes.

Negishi et al. (1975) demonstrated that NADPH-cytochrome c reductase, after completion of synthesis on membrane-bound polysomes, was exposed on the surface of the endoplasmic reticulum toward the cytoplasm, since its activity could be abolished by the addition of proteases. Harano and Omura (1978) have further investigated the synthesis of this enzyme on membrane-bound polysomes and found that it occurred exclusively on the loosely bound polysomes and not on the tightly bound class. Cytochrome bs, another microsomal enzyme, is apparently synthesized only on membrane-bound poly-

134 Asbjorn M. Svardal and Ian F. Pry me

somes (Harano and Omura, 1977a). Its synthesis, furthermore, appears to be restricted to the loosely bound class (Harano and Omura, 1978). This enzyme is also discharged from membrane-bound polysomes on to the cytoplasmic surface of the endoplasmic reticulum (Harano and Omura, 1977b).

Early studies on the location of the synthesis of cytochrome P-450 suggested that the free polysomes represented the major site (Ichikawa and Mason, 1973, 1974). Craft et al. (1975) showed that these results could be explained by a contamination of the free polysomes with membranes. Correia and Meyer (1975) provided evidence that P-450 synthesis occurs in the endoplasmic reticulum. Using an antibody raised in rabbits against purified cytochrome P-450 from phenobarbital-treated rats, Negishi et al. (1976) were able to demonstrate convincingly that the cytochrome is indeed synthesized predominantly on membrane-bound polysomes. In addition, Fujii-Kuriyama et al. (1979) have shown that the synthesis occurs preferentially on the tightly bound class of membrane-bound polysomes. They observed that nascent cytochrome P-450 peptide chains released from polysomes by puromycin/high salt treatment were found to be qualitatively associated with the endoplasmic reticulum. Treatment of these membranes with trypsin/chymotrypsin resulted in an approximate loss of 90% of the nascent peptides, suggesting that following synthesis the cytochrome is most likely integrated directly into the cytoplasmic face of the endoplasmic reticulum. That the whole molecule, or at least part of it, is exposed toward the cytoplasm has been indicated by immunoelectron microscopy (Matsuura et al., 1978, 1979), immunochemical techniques (Thomas et al., 1978; Welton et al., 1975), and enzymatic iodination (Welton and Aust, 1974). Fujii-Kuriyama et al. (1979) concluded that cytochrome P-450 first appears in the rough endoplasmic reticulum and is then translocated to the smooth endoplasmic reticulum by lateral movement in the plane of the membrane, reaching an even distribution between the two types of membrane within about 1 hr. Cytochrome P-450, however, is not only found on the cytoplasmic face of the endoplasmic reticulum, but also on the luminal side (Depierre and Ernster, 1977; Nilsson and Dallner, 1977). Craft et al. (1978) demonstrated the in vitro synthesis of apocytochrome P-450 in light rough endoplasmic reticulum from rat liver, and further found that when microsomes were treated with trypsin after completion of in vitro protein synthesis, immunoprecipitable P-450 was still obtainable from the vesicles following deoxycholate treatment in the presence of protease inhibitor (Craft et al. 1979a). Craft et al. (1979b) have thus been able to conclude that at least part of the newly synthesized cytochrome P-450 is translocated into the lumen of the rough endoplasmic reticulum before further being transported into other components of the reticulum and then ultimately incorporated into appropriate membrane sites.

The possibility that mitochondrial proteins may be synthesized on mem-


brane-bound polysomes has received much attention. An apparent continuity between the endoplasmic reticulum and the outer mitochondrial membrane has been observed (Franke and Kartenbeck, 1971; Morn~ et al., 1971), and an integration of proteins synthesized on membrane-bound polysomes into the mitochondrial outer membrane could thus be envisaged. Mitochondrial proteins, following completion of synthesis, may later appear in vesicles that would then fuse with the outer mitochondrial membrane, and the contents released into the mitochondria (Chua and Schmidt, 1979). Another possibility would be that the synthesis of mitochondrial proteins could occur on ribosomes attached to the outer surface of mitochondria such that the products could then be directed immediately into the organelle without any intermediate transport steps being necessary (Butow et al., 1975; Kellems and Butow, 1972, 1974; Kellems et al., 1974, 1975). One could also envisage that the synthesis of proteins destined to be incorporated into organelles could occur on free poly somes (Cashmore et al., 1978; Dobberstein et al., 1977; Roy et al., 1977), the completed molecules then recognizing specific receptors at the outer surface of the appropriate organelle.

The site of synthesis of rat liver mitochondrial cytochrome c has been the subject of some controversy. Early work (reviewed by Schatz and Mason, 1974) indicated that the endoplasmic reticulum was the likely site. Using in vitro experiments, Gonzalez-Cadavid and Cordova (I974) found that cytochrome c was apparently synthesized by both free and membrane-bound polysomes. Recently Robbi et al. (I 978a,b) have clearly shown that the cytochrome, in fact, is not synthesized on the rough endoplasmic reticulum. The appearance of cytochrome c in microsomes has becn attributed to be a contaminant arising from an in vitro release of the protein from mitochondria, which then enters a strong relationship with ribosomes or microsomal membranes or both. Robbi et al. (1978a) suggest that cytochrome c is synthesized by ribosomes associated with the outer surface of mitochondria or alternatively on free polysomes.

There is evidence suggesting that enzymes present in the mitochondrial matrix are synthesized by membrane-bound polysomes. Thus Bingham and Campbell (I972) identified synthesis of malate dehydrogenase in rat liver microsomes, and both Godinot and Lardy (1973) and Kawajiri et al. (1977) concluded that glutamate dehydrogenase is synthesized by membrane-bound polysomes. Kawajiri et al. (1977) found that the enzyme is located on the cytoplasmic side of the endoplasmic reticulum after completion of synthesis, because (I) activity was abolished upon the addition of proteases, (2) the protein was accessible to antibody binding, and (3) it could be washed from its site in the membrane by buffers of moderately high ionic strength.

Shore and Tata (I977a) have prepared rough microsomes, rapidly sedimenting endoplasmic reticulum (RSER), and free polysomes from rat liver,


and mRNA isolated from these fractions was translated in an in vitro system. The products of translation were immunoprecipitated using antibodies against mitoplast proteins and electrophoretic studies indicated that rough microsomes, and not RSER or free polysomes, were involved in the synthesis of mitochondrial proteins (Shore and Tata, 1977b).

Chuah and Schmidt (1979) have comprehensively reviewed the current knowledge concerning the subcellular locations of the sites of synthesis of mitochondrial and chloroplast proteins. Their review also covers aspects of the transport of these proteins into the respective organelles.

Elder and Morre (1976) have found that a number of intrinsic membrane proteins in rat liver are common to various membrane-containing fractions: smooth microsomes, rough microsomes, nuclear membranes, Golgi membranes, and plasma membranes, whereas the latter two membranes types exhibited proteins not found in smooth or rough microsomes. The precipitation of poly somes with antibodies against integral membrane proteins led to the observation that proteins common to all membranes were synthesized predominantly on membrane-bound polysomes, whereas those proteins found only in plasma and Golgi membranes were specifically synthesized on polysomes associated with the Golgi fraction.

3.4. Summary

For cells with a well-defined rough ER, the evidence supports the concept that distinct classes of proteins are synthesized by membrane-bound polysomes. These include proteins destined for export or for transport to other cellular organelles and those membrane proteins that form the tightly bound class of intrinsic proteins or that are deposited asymmetrically toward the noncytoplasmic face of membranes (see Figures 1-3.). A common feature of all these proteins is that they require transfer either deep into or across membrane lipid bilayers.

When synthesis does occur in the bound ribosome fraction, the result is the compartmentalization or segregation of the newly formed proteins. Compartmentalization may be advantageous for a variety of cell functions, but the potential changes that the rough ER can undergo in certain cells may conceivably lead to modification or even suspension of these functions. This is especially evident in liver tissue, where the amount and organization of ER may change in order to cope with an alteration in environmental conditions, particularly in response to dietary fluctuations and when enhanced detoxification processes become necessary. Under such conditions it seems probable that there can be fluctuations in the synthesis of a given protein by either bound or free ribosomes (Lowe and Hallinan, 1973).

As mentioned earlier, the RERjSER ratio can vary greatly during growth


and differentiation of both secretory and predominantly nonsecretory tissues. This indicates the important functional role of the ER in synthesizing cell structural proteins. In young differentiating muscle, choroidal epithelial cells, and brain cortical cells, for example, a major fraction of ribosomes are membrane bound, but this fraction diminishes as differentiation is completed (Birge and Doolin, 1974; Greenstein, 1972). Although stimulation of liver growth by drugs, hormones, or partial hepatectomy results in a proliferation of RER, this is usually accompanied only by minor changes in the synthesis of secreted proteins compared with total liver protein production (Schreiber et al., 1971).

4. POLYRIBOSOME-MEMBRANE INTERACTIONS

There is considerable evidence to suggest that more than one component of the polyribosome complex can interact with the membrane of the rough endoplasmic reticulum. Great attention has been directed toward an elucidation of the mechanism(s) involved in selecting specific messenger RNAs for translation by membrane-bound ribosomes. Differences in type of association between polysomes and membrane could account for differential translational control for the mRNA involved.

Three mechanisms have been described suggesting how polyribosomes may interact with the membrane: through the 60 S subunit, the growing nascent chain, and/or the mRNA itself. These mechanisms will now be briefly considered.

4.1. The Physical Nature of Binding between 60 S Subunit and Membrane

Sabatini et al. (1966) found that when liver membrane-bound ribosomes were treated with increasing concentrations of EDT A, the small ribosomal subunits were selectively detached at low concentrations of the chelating agent. At higher concentrations all the small subunits were released, together with some large subunits. They thus proposed that the ribosomes were bound to the membrane by their large subunits, and this view was supported by electron microscopic studies. It is presumed that chelating agents interfere with ribosome binding by breaking divalent cation bridges between ribosomal subunits or subunits and membrane (McIntosh and O'Toole, 1976).

High concentrations of monovalent ions have also been found to detach membrane-bound ribosomes and are presumably able to do so by interfering with the integrity of divalent ion bridges between interacting macromolecules (Blobel and Sabatini, 1971).

Under conditions of high KCl concentration (e.g., 500 mM) and in the

138 Asbjorn M. Svardal and Ian F. Pryme

presence of Mg2+ only partial detachment occurs, but addition of 1 mM puromycin followed by an incubation at room temperature brings about the release of up to 85% of the ribosomes from liver rough microsomes (Adelman et al., 1973). The released nascent chain remained associated with the membrane. At sufficiently low concentrations of KCl (e.g., 25 mM) there was little detachment of ribosomes from the vesicular surface, even in the presence of sufficient puromycin to react with most of the nascent chains. These findings, summarized in Figure 4 (Adelman et al .. 1973), suggest that ribosomes may be bound either by ionic forces alone (detached by high KCl concentration) or by ionic forces and the nascent polypeptide chain acting in concert (detached only by high concentration of KCl when puromycin is present). These concepts have been supported by the work of Lande et al. (1975) with human diploid fibroblasts and Harrison et al. (197 4a) with mouse myeloma cells.

The question then arises as to whether it is the RNA or protein moiety

Low KCI + Puromycin

High KCI + PuromYCin

High KCI

+ Puromycin

FIGURE 4. Schematic model of possible mechanisms of interaction between ribosomes and membranes in rat liver microsomes and of the mode of action of puromycin or high KCl concentration or both. [Reprinted from Adelman et al. (J 973) with permission of the authors and the Journal of Cell Biology.]


(or both) of the 60 S subunit that is involved in ribosome attachment. Demonstration of heterogeneity in the protein composition of free and bound ribosomes has indicated that proteins are of importance (Ramsey and Steele, 1977; Blobel and Dobberstein, 1975a). On the other hand, Scott-Burden et al. (1972) found that addition of nucleic acids interfered with binding of ribosomes to stripped rough microsomes in vitro. This, together with suggestions that regions of ribosomal RNA are partially exposed on the surface of the ribosome (Cox and Godwin, 1975; Wittman, 1976), suggests that the RNA moiety can be important.

Several lines of evidence have indicated that integral membrane proteins of the rough endoplasmic reticulum play an important role in ribosomal binding (Borgese et al., 1974). The ability of stripped rough endoplasmic reticulum (puromycin/KCI treatment) to rebind ribosomes in vitro was abolished as a result of performing mild proteolysis or heat denaturation. On the other hand, neither removal of peripheral membrane proteins nor treatment with phospholipase had any effect on the reattachment of ribosomes to stripped rough microsomes. Use of the nonionic detergent Kyro EOB (a polyethoxyalkylether) has proved of value in identification of specific proteins involved in ribosome binding. This detergent was first used by Birckbichler and Pryme (1973) for separating free and membrane-bound polysomes from suspension-cultured cells; at suitable concentrations of the detergent, the plasma membrane was sufficiently solubilized to allow release of free polysomes, and while polysomes attached to the endoplasmic reticulum remained membrane bound, they were released from the endoplasmic reticulum only upon the addition of the ionic detergent sodium deoxycholate. Kreibich et al. (1978a) showed that Kyro EOB solubilized by far most of the membrane proteins and phospholipids from the endoplasmic reticulum without affecting the association of two polypeptides with ribosomes. These two polypeptides were shown to be integral membrane proteins and to have molecular weights of 63,000 and 65,000. They have been termed ribophorins I and II. Following the partial solubilization of rough endoplasmic reticulum membranes by Kyro EOB, the two proteins in the remaining membraneous material were found to exist in a fixed stoichiometric ratio with respect to number of ribosomes present (Kreibich et al., 1978c). Both proteins are apparently exposed on the surface of rat liver microsomes, since they could be iodinated by a lactoperoxidase catalysed reaction (Kreibich et al., 1978b). That the ribophorins are in direct association with 60 S ribosomal subunits was demonstrated by cross-linking experiments (Kreibich et al., 1978b). A preferential cross-linking of 60 S subunits to microsomal membranes could be achieved by using extremely low glutaraldehyde concentrations (0.005-0.02%), and puromycin/KCI treatment was then unable to cause release of ribosomes from membranes. When low concentrations of methyl-4-mercaptobutyrimidate were used, the cross-linking of ribophorins to ribosomes

140 Asbjl)m M. Svardal and Ian F. Pryme

was shown to be reversible. The ribophorins were released together with ribosomes when membranes were solubilized by 1 % sodium deoxycholate and could then be separated from the ribosomes by disruption of cross-links by reduction. Nascent polypeptide chains are apparently not involved in the association between ribophorins and ribosomal proteins of the 60 S ribosomal subunit. As the binding capacity of rough endoplasmic reticulum stripped of ribosomes is unaffected by either high salt or EDT A treatment, it can be assumed that proteins of the binding sites are intrinsic membrane components interacting directly with the microsomal phospholipid bilayer (Kreibich et ai., 1978a; Sabatini and Kreibich, 1976).

It is almost certain that ribosomal binding sites have other functions than merely providing attachment of 60 S ribosomal subunits to the cytoplasmic face of the rough endoplasmic reticulum. They are probably organized into functional complexes responsible for accepting and processing selected classes of polypeptides manufactured on membrane-bound polysomes. It is therefore supposed that a multitude of functions are associated with the ribosomal binding sites and the ribosome-membrane junctions. At present the most obvious are (1) binding of large ribosomal subunits, (2) the recognition and cleavage of the hydrophobic amino acid sequence at the amino-terminal end of the nascent chain polypeptide chains (signal peptide, see Section 4.2), (3) the transfer of nascent polypeptide chains across the membrane to the luminal face, and (4) posttranslational modifications (see Sections 4.2 and 5.2) necessary to determine the ultimate destination of polypeptides that cross the membrane (Blobel and Dobberstein, 1975a,b; Ojakian et ai., 1977).

It is well known that lipids and membrane proteins can exhibit lateral mobility in the plane of the membrane (see Singer, 1974; Singer and Nicolson, 1972), and a similar mobility of ribosomes bound to the rough endoplasmic reticulum has been shown to occur under experimental conditions (Ojakian et aI., 1977). The mRNA of bound polysomes was cleaved by ribonuclease treatment of salt-washed rough microsomes, thereby allowing for a potential lateral movement of individual bound ribosomes. When microsomes were treated with RNase at 4 0 C and were then maintained at this temperature until fixation for electron microscopy, freeze-etch and thin-section studies showed that the ribosomes retained a pattern of homogeneous scatter on the membrane surface. This was in contrast to the situation observed after incubating the RNasetreated microsomes at 24 0 C for 30 min before fixation; this temperature is above the thermotropic phase transition of the microsomal phospholipids. Under these conditions, large tightly packed aggregates of ribosomes were observed. It was thus evident that 60 S ribosomes together with their attached binding sites, other membrane-associated proteins, and nascent polypeptide chains, were capable of performing lateral displacement within the membrane. As this movement was temperature dependent, it can be assumed that it is controlled by the fluidity of the membrane.


In addition to the above experimental evidence suggesting a mobility of ribosomes/ribosomal binding sites, there are results from electron-microscopic studies that suggest that this mobility also occurs in vivo. Grazing sections of rough endoplasmic reticulum membranes have shown characteristic spiral patterns that have been interpreted to represent polysome configurations (Baglio and Farber, 1965; Palade, 1955a, 1975). When initiation of protein synthesis was inhibited by either Verrucarin A or ethionine (Adesnik et al., 1976; Baglio and Farber, 1965), these structures disappeared. They reappeared, however, upon removal of the inhibitor (Adesnik et aI., 1976). The biological role of the potential mobility of ribosomal binding sites has been suggested to be an efficient provision of new binding sites in the region of the free 5' end of mRNAs translated in association with the rough endoplasmic reticulum (Ojkaian et al., 1977).

The actual dynamics of movement of the components of membrane-bound polysomes during protein synthesis are poorly understood. One could imagine that either a movement of ribosomes (plus binding sites) within the lateral plane of the membrane along a fixed mRNA molecule (see Section 4.3) or the displacement of mRNA along a series of immobilized ribosomes would both satisfy the requirements. A further possibility can be envisaged should both the ribosomes and mRNA be bound to the membrane and fixed relative to one another; in this case a long untranslated region of the messenger molecule close to the site of its binding to the membrane would be required (Sabatini and Kreibich, 1976). It has been suggested that such an arrangement would give rise to spiral patterns of polysome aggregates observed in the electron microscope (Baglio and Farber, 1965; Palade, 1955a, 1975). The various models for the translation of mRNAs in bound polysomes have been considered in detail by Sabatini and Kreibich (1976).

As opposed to the situation in rat liver (Figure 5a), a major fraction (30-60%) of ribosomes present in the bound polysomes population in cultured cells (e.g., HeLa, mouse sarcoma, ascites tumor cells) is anchored indirectly to the membrane via a "dangling" messenger (i.e., not all ribosomes on the mRNA are attached to the membrane, see Figure 5b). These are released following mild digestion of polysomes with ribonunclease (Faiferman et al., 1973; Mechler and Vassalli, 1975; Rosbash and Penman, 1971a). In addition (Bleiberg et al., 1972), the unattached ribosomes can be released by any treatment causing polysome disaggregation (e.g., puromycin, EDT A). Accordingly, two classes of membrane-bound ribosomes have been defined in cultured cells-loosely bound and tightly bound (Rosbash and Penman, 1971a,b). In myeloma cells, the ribosomes liberated following treatment of rough microsomes with ribonuclease (loosely bound fraction) contain nascent polypeptide chains found on average to be two to three times shorter than chains associated with the tightly bound ribosomes (Mechler and Vassalli, 1975). This implies that only recently initiated 60 S subunits are liberated from intact polysomes following disaggre-


5' a

5'

~ b

5'

0--FIGURE 5. Polyribosome-membrane interactions. Polysomes may become membrane-bound by mechanisms involving (I) interactions between sites in the membrane and 60 S ribosomal subunits, (2) interactions between nascent polypeptides and membrane receptors, and (3) binding of a region at the 3' end of the mRNA to a specific membrane site (not shown). Situations: (a) Described in rat, where 60 S subunits may first interact directly with the membrane before being later anchored by growing nascent polypeptides entering the lumen of the endoplasmic reticulum (evidence: ribonuclease treatment of isolated rough microsomes removes relatively few ribosomes). (b) Described for cultured myeloma cells, suggests that functioning 60 S ribosomes at the 5' end only become membrane-bound when their associated nascent chains are long enough to establish interaction with membrane receptor proteins (evidence: before attachment occurs large amounts of ribosomes are released upon mild digestion of the microsomes with ribonuclease). (c) Based on indirect evidence, indicates how polypeptides synthesized on membrane-bound polysomes could be released free to the cytoplasm rather than be discharged into the lumen of the endoplasmic reticulum (cf. Figure 3). [Reprinted from Shore and Tata (1977c) with the permission of the authors and Biochim. Biophys. Acta.]

gation, i.e., both classes are present in the same polysome complex (Figure 5b). This finding is not consistent with the original suggestion that loosely bound and tightly bound ribosomes may represent different functional groups in HeLa cells (Rosbash and Penman, 1971a,b).

Svardal and Pryme (l980a) have shown that under specific experimental conditions shearing forces generated during ultracentrifugation of light rough microsomes from MPC-ll cells on discontinuous gradients, can result in the


artifactual production of two classes of polysomes. One class consists of ribosomes associated with the 5' end of the mRNA (short polypeptide chains), which has been cleaved from the 3' end of the molecule and thus loses its association with the membrane, whilst the ribosomes on the 3' end remain associated with the ER (longer polypeptide chains). The polysomes appearing on the cleaved mRNA fragments obviously cannot be termed loosely bound and tightly bound, as they do not represent different polysome classes; in fact, they originate from the same polysome fraction.

Loosely and tightly bound ribosomes have also been defined on the basis of whether or not attached ribosomes are further anchored to the membrane by a growing nascent chain (Adelman et al., 1973; Harrison et al., 1974a). By this definition, loosely bound ribosomes represent the fraction released by high concentrations of KCI alone, whereas tightly bound ribosomes require prior treatment with puromycin. Studies have been performed to examine possible functional differences between the two classes. Harrison et al. (197 4b) using myeloma cells showed that immunoglobin mRNA was present in both KCIsensitive and KCI-resistant polysome populations. These authors suggested that loosely bound ribosomes are precursors of the tightly bound fraction. In contrast, the possibility exists that the poly (A) segment at the 3' end of the most mRNAs may playa selective role in the segregation of mRNA into the two fractions. Zauderer et at. (1973), using myeloma cells, reported that histone mRNA [which lacks this poly (A) segment] is found only in the loosely bound fraction of the membrane-bound polysomes. In rat liver the synthesis of albumin is apparently restricted to tightly bound ribosomes (Tanaka and Ogata, 1972). In all the studies that purport to show functional differences between tightly and loosely bound ribosomes it is very difficult to rule out the possibility that the loosely bound fraction arises artifactually by contamination of rough microsomes with free polysomes, and this indeed has been proposed to be the case (Ramsey and Steele, 1977). At least at low ionic strength in vitro, rough microsomes are well known to bind free ribosomes nonspecifically (Borgese et al., 1974). Nevertheless recent studies with rat liver (Shore and Harris, 1977) suggest that most, if not all, polypeptides synthesized on purified rough microsomes in vitro are vectorially discharged either deep into or across the membrane.

4.2. Interaction between Nascent Polypeptides and Membranes

As already discussed, the presence of the nascent polypeptide does not appear to be necessary for the integrity of the ribosome-membrane complex at low monovalent cation concentration, but does add stability at higher concentrations.

Mild proteolysis of rough microsomes causes an extensive removal of pro-


teins and ribosomes from the outer face of the membrane (Sabatini and Blobel, 1970), but did not lead to complete digestion of nascent chains. Instead, sufficiently long nascent polypeptides were cleaved into two main classes of fragments, both of which were largely protected from proteolysis. Intraribosomal fragments of approximately 39 amino acids in length, located near the carboxyterminal end of the nascent polypeptides, were removed together with the detached ribosomes. The amino-terminal portions of the nascent chains were retained in the denuded microsomes. Puromycin releases nascent chains from bound ribosomes so that attachment of nascent chain (i.e., the peptidyl tRNA) to the large subunit is mediated only through its tRNA moiety (Shore and Tata, 1977c). The carboxy-terminal sequences of the nascent chains which are at anyone time buried within the large subunit do not themselves strongly interact with ribosomal components.

With respect to the mechanism whereby nascent polypeptide chains are transferred across the membrane, Blobel and Sabatini (1971) formulated the so-called "signal" hypothesis, which has received support from several recent studies (Blobel, 1977; Blobel and Dobberstein, 1975a,b; Devillers-Thiery et aI., 1975). It has been found that the primary translation products in heterologous cell-free systems of a variety of messengers coding for secreted proteins are larger than either the final product secreted in vivo or the product that is proteolytically processed in vivo in the smooth endoplasmic reticulum-Golgi system. Examples of these include immunoglobulin light chain (Blobel and Dobberstein, 1975b; Milstein et aI., 1972; Schechter and Burstein, 1976), the milk whey protein a-lactalbumin (Craig et aI., 1976), insulin (Chan et ai., 1976; Duguid et ai., 1976), parathyroid hormone (Habener et ai., 1976), and pancreatic secretory proteins (Devillers-Thiery et ai., 1975). An extra N-terminal amino acid sequence (the signal) is found on these precursors. Several signal peptides have been sequenced, and their amino acid compositions are thus known (e.g., see Leader, 1979). All exhibit a similar content of hydrophobic residues, although they otherwise differ in length (from about 15-30 amino acids) and are significantly different with regard to amino acid composition. The signal peptide sequences have been termed "pre" such that they are not to be confused with "pro" sequences which represent intermediate protein precursors (e.g., proalbumin, proinsulin). Thus it is proposed that after initiation of protein synthesis, the growing hydrophobic signal sequence is channeled through the 60 S subunit (Blobel and Dobberstein, 1975b). When 10-40 amino acid residues of the nascent chain have emerged from the ribosome, attachment occurs between the signal sequence and specific receptor proteins present in the membrane, and the rest of the polypeptide chain will naturally follow as translation proceeds. In the case of the light chain precursor enzymatic cleavage (signalase or clippase) of the signal sequence is apparently achieved before synthesis of the polypeptide has been completed (Blobel and


Dobberstein, 1975b; Milstein et al., 1972), implying an interaction between enzyme and nascent polypeptide chain, As the portion of the nascent chain which extends into the luminal space assumes a bulky, tertiary structure, passage across the membrane becomes virtually undirectional and will provide an effective form of anchorage of membrane-bound polysomes to the endoplasmic reticulum (Adelman et al., 1973; Sabatini et al., 1972; Lampen, 1974). Details of the signal hypothesis are summarized in Figure 6.

Wickner (1979) suggests a second function for a hydrophobic leader sequence at the N-terminal end of the polypeptide chain, and this function is incorporated in a model termed the membrane-trigger hypothesis. In this model the hydrophobic sequence (and also possibly other regions of the protein) recognizes either protein or lipids in the already existing membrane structure, enabling binding to occur. Such binding may also be inaugurated by specific physical properties of predetermined areas within the lipid bilayer. Once bound, the protein sequence would fold upon interaction with the lipid elements in such a manner that the hydrophobic residues of the polypeptide would be exposed to the fatty acyl chains of the lipid bilayer. Once the protein is correctly orientated within the membrane, the hydrophobic sequence can be cleaved and the protein would assume a permanent role as a membrane protein. As in the case of the signal hypothesis, the hydrophobic amino acid sequence may well be removed before the protein is completely synthesized. It is thus likely that a hydrophobic N-terminal sequence may not only be necessary to direct polypeptide chains through the ER into the lumen (e.g., secretory proteins, luminal proteins), but also to lead them into predetermined sites

5' 7 o 3'

'RIBOSOME RECEPTOR PROTE IN

FIGURE 6. Details of the signal hypothesis shown in a schematic form. [Reprinted from Campbell and Blobel (1976) with permission of the authors and FEBS Letters.]


within the membrane structure (e.g., various types of membrane proteins; see Figure 1).

In addition to an interaction between the hydrophobic signal sequence and specific receptor proteins in the ER membrane, other forms of association between growing peptide chains and the membrane can be envisaged. Many proteins undergo structural modification in the ER, and at least some of these are modified before the polypeptide chain has been completely synthesized on membrane-bound polysomes. The synthesis of collagen has been studied by Lazarides et al. (1971), using stationary phase 3T6 cells, and by Diegelmann et al. (1973), using 14-day-old chick embryo tibiae. Both reports conclude that prolyl hydroxylase is a membrane-bound enzyme capable of hydroxylating proline residues in nascent collagen polypeptides still attached to membranebound polysomes. Because the degree of hydroxylation of released, completed collagen chains was not higher than on nascent collagen peptides (Lazarides et al .. 1971), an interaction between the latter and the prolyl hydroxylase in the ER can be inferred. An enzyme that catalyzes sulphydryl-disulfide interchange in proteins has been isolated from beef liver microsomes (De Lorenzo et al .. 1966) and is apparently engaged in the process whereby the portion of the nascent polypeptide chain that has entered the lumen of the ER attains a specific three-dimensional conformation. One can again envisage a direct association between membrane-bound enzyme and growing polypeptide chain. Another well-studied process is glycosylation (see Section 5.2), in which the incorporation of sugar residues into nascent polypeptide chains on membranebound polysomes has been thoroughly documented. These studies indicate a further form of interaction between nascent polypeptide chain and a membrane-bound enzyme, in this case a glycosyl transferase.

The sequence of events suggested in the signal hypothesis may not be restricted to secretory proteins, but it may apply to the synthesis of all proteins that have to be transferred across a membrane (Blobel and Dobberstein, 197 5b) (e.g., endoplasmic reticulum proteins, mitochondrial, lysosomal, plasmalemmal proteins). The question then arises as to whether these are released to the membrane, to the intraluminal space, or free to the cytosol.

A great many investigations have demonstrated that not all functioning ribosomes associated with the rough endoplasmic reticulum contain polypeptide chains that interact with the membrane. One could visualize that 60 S ribosomes at the 5' end of mRNA are bound to the membrane, but contain nascent polypeptide chains below a critical length to allow interaction to occur with the membrane (Figure 5b). A different situation may exist for myeloma cells grown in culture wherein 60 S subunits at the 5' end of mRNA may not initially be membrane bound and only become so after a sufficiently long sequence of amino acids emerges from the subunit, enabling membrane binding to occur. Attachment is thus afforded by both direct association of 60 S


subunits with ribophorins and an interaction of the signal sequence with membrane receptors. Circumstantial evidence exists suggesting that primarily nonsecretory tissue contains classes of membrane-bound polysomes whose growing nascent chains may not interact strongly with membranes (Andrews and Tata, 1971; Tata, 1971; Zauderer et al., 1973), perhaps because of a lack of the signal sequence at the N-terminus end of the nascent polypeptide. Upon completion of synthesis, therefore, the proteins would be expected to be released free to the cytoplasm (Figure 5c) or perhaps to remain associated with the cytoplasmic face of the endoplasmic reticulum. In the latter case proteins may be synthesized directly into predetermined, preexisting sites on the endoplasmic reticulum.

4.3. Direct Interaction between Messenger RNA and Membranes

Studies of the fate of mRNA after termination of protein synthesis with puromycin in a buffer of high ionic strength, or in the absence of Mg2+, have led to the conclusion that in fibroblasts (Lande et al., 1975) and HeLa cells (Milcarek and Penman, 1974) some mRNA species of bound polysomes are directly associated with the ER membranes, independently of ribosomal attachment and the peptidyl tRNA. After extensive release of ribosomes brought about by several procedures, 40-60% of the mRNA always remained attached to the sedimentable ribosome-stripped membrane vesicles. The poly A segments, 150-200 nucleotides in length, which are present at the 3' end of the bound mRNA molecules, maintained their association with the membrane even after the non-poly-A-containing regions of the messengers were selectively digested with T 1 or pancreatic RNase in media of high ionic strength. Similar results were obtained from rat liver (Cardelli et al., 1976) and also from human fibroblasts after the cells had been incubated in a medium containing an inhibitor of initiation, Verrucarin A, which causes extensive polysome runoff (Adesnik et al., 1976). These workers conclude that mRNA contains a site of attachment to the membranes that is protected from attack by pancreatic RNase and is located in or immediately adjacent to the poly A region. This site may constitute a direct recognition sequence for the membrane, or more likely, for proteins that serve to bind the mRNA to the ER membranes (Sabatini and Kreibich, 1976).

Several proteins are found associated with poly A segments in mRNA (Axelsson et al., 1977; Blobel, 1973; Mazur and Schweiger, 1978; Ovchinnikov et al., 1978). These proteins may participate in the formation of ribonucleoprotein particles (mRNP) or may be involved in the binding of mRNA to rough endoplasmic reticulum.

It also seems likely that the strength of the binding by which the mRNA is associated with the membrane may vary according to the condition of iso-

148 Asbjijrn M. Svardal and Ian F. Pry me

lation (Harrison et al., 1974b, Kruppa and Sabatini; 1977; Mechler and Vassalli, 1975).

The actual function of the binding between mRNA and the ER membranes has been the source of much speculation. Some investigators have found that the binding may be important for the transfer of newly synthesized mRNA from the nucleus to the cytoplasm (Faifermann et al., 1971; Shiokawa and Pogo, 1974). Sabatini and Kreibich (1976) proposed that the main function is to ensure the proximity of the mRNA to the ribosomal binding sites and to facilitate reutilization of the messenger when reinitiation occurs after translation has been interrupted.

Adesnik et a/. (1976) suggest the possibility that although mRNAs coding for secretory proteins may not be tightly bound to endoplasmic reticulum membranes, other messengers that code for nonsecretory proteins (e.g., those predominating in Hela cells, fibroblasts) are perhaps directly associated with the endoplasmic reticulum since they are apparently not removed under conditions promoting the disassembly of polysomes in vivo and in vitro.

In conclusion, it can be said that considerable evidence exists indicating that a direct association between mRNA and ER membrane occurs in several cell types, but that the actual biological role remains to be elucidated.

5. HETEROGENEITY IN THE FUNCTION OF ROUGH ENDOPLASMIC RETICULUM WITH RESPECT TO PROTEIN SYNTHESIS

Until recently there was little reason to expect that different cisternae of the same endoplasmic reticulum system are functionally separate with respect to the synthesis of a type or group of proteins. A subdivision of the rough endoplasmic reticulum into smaller compartments engaged in different protein synthesizing activities would create a number of questions that are at present difficult to answer, e.g., How could one envisage that a specific mRNA species could become associated with the "correct" area of endoplasmic reticulum where its translation was to occur?

Recent studies have shown that the endoplasmic reticulum may indeed not consist of identical cisternae but instead be built up of individual microenvironments.

5.1. Compartmentalization of the Synthesis of Proteins Destined for Discharge to the Extracellular Environment

It is now well established that proteins destined for transport out of the cell must first be concentrated within the lumen of the rough endoplasmic


reticulum (Palade, 1975). This is achieved by compartmentalizing the synthesis of secretory proteins on ribosomes attached to the endoplasmic reticulum. Concentration of proteins within the lumen of the rough endoplasmic reticulum presumably serves to channel the protein properly through the secretory apparatus. Not all cells, however, that possess an endoplasmic reticulum and a Golgi complex necessarily secrete proteins, and it is becoming clear that unique groups of proteins that subsequently become incorporated into plasma membrane, mitochondria, lysosomes, peroxisomes, etc., are also probably synthesized on membrane-bound polysomes (Shore and Tata, 1977c). Thus in addition to a compartmentalization of the translation of mRNA for secretory proteins within the endoplasmic reticulum, there must also be a preferential segregation of other groups of mRNA species on the rough endoplasmic reticulum.

5.2. Compartmentalization of Protein Synthesis in the Endoplasmic Reticulum and Specific Posttranslational Modifications

It has been long realized that the polypeptides, in transit from their site of "'synthesis on the ribosomes attached to the rough endoplasmic reticulum, undergo structural changes as the nascent chain appears on the luminal side of the membrane or as completed molecules pass through RER, SER, and the Golgi complex. Examples of such changes are proteolytic cleavage, disulfide bridge formation, and chemical modification (e.g., glycosylation, phosphorylation, hydroxylation, iodination, lipidation) (Shore and Tata, 1977c).

The type of posttranslational processing most studied is glycosylation. In this process glycoproteins are produced by stepwise enzyme-catalyzed additions of sugar residues to certain amino acids; L-asparagine may be substituted by N-acetyl-j3-o-glucosamine; L-serine by j3-o-xylose, N-acetyl a-o-galactosamine, mannose, or galactose; and L-threonine by the latter three sugar residues. In higher animal collagen, hydroxylysine residues undergo j3-o-galactosylation, whereas hydroxyproline present in some plant proteins may be substituted by L-arabinose or L-galactose. Glycoproteins occur most abundantly as secretory products and as components of most cellular membranes (Hughes, 1976) and constituents of certain organelles, e.g., lysosomes (Goldstone and Koenig, 1970) and mitochondria (Martin and Bosmann, 1971). The glycosyl transferases themselves are located in the cell only in association with membranes (Baker et al., 1972; Dallner, 1963; Lawford and Schachter, 1966; Schachter, 1974; Stoolmiller et al., 1972), and the process of glycosylation thus embodies most of the points that stress the necessity for the compartmentalization of the synthesis of specific protein species on membrane-bound polysomes.

Although most proteins synthesized on membrane-bound polysomes are


glycosylated to some degree or another within the endoplasmic reticulumGolgi system, serum albumin is an example of a protein that contains no carbohydrate side chain (Redman and Cherian, 1972). This discovery together with the finding that the secretion of immunoglobulins lacking carbohydrates in MPC-ll mutants is not impaired (Weitzman and Scharff, 1976) has been used to argue that glycosylation is not the underlying requirement demanding that a protein be synthesized by bound ribosomes.

With respect to the sequence with which sugar moieties are added to the protein acceptor, kinetic studies have shown that different sugars may be incorporated into the same protein at different subcellular loci (Melchers, 1971). This, together with the fact that glycosyl transferases are located at different sites (Bouchilloux et al., 1970, 1973), has led to the widely held view that the passage of a single protein occurs sequentially through the various membrane compartments (e.g., rough endoplasmic reticulum -- smooth endoplasmic reticulum -- Golgi -- vesicles -- plasma membrane), each of which contains its own unique set of membrane-bound glycosyl transferases. These, in turn, determine to a great extent the rather complex oligosaccharide patterns of glycoproteins. Studying the biosynthesis of the carbohydrate portion of immunoglobulin, Melchers (1971) found that glucosamine and mannose were added to the protein, while it was still in the rough endoplasmic reticulum. Galactose, on the other hand, and presumably fucose, were incorporated into the oligosaccharide chains within the smooth membranes-Golgi complex.

Another interesting point is that in addition to this classic scheme for the sequential addition of monosaccharide residues to polypeptide acceptors, there is a different mechanism involving a dolichol derivative as an intermediate in a reaction where preas sembled oligosaccharides containing man nose and Nacetylglucosamine are transferred as a unit to a polypeptide acceptor (Lennarz, 1975).

In addition to secretory and cell-surface proteins, the synthesis of membrane glycoproteins and glycoproteins residing inside various nonsecretory organelles (e.g., lysosomes, mitochondria) also presumably occurs on bound ribosomes. This has led to much speculation about the mechanisms whereby newly synthesized proteins are channeled to their proper locations in the cell. An example of the problem is provided by the work of Novikoff et al. (Novikoff et al., 1974; Novikoff, 1976) on thyroid epithelial cells. They showed that two types of secretory granules are apparently formed-A-granules and B-granules. The former contain peroxidase activity, but the latter do not, yet the enzyme can be detected throughout the rough endoplasmic reticulum. This indicates that the mechanism for properly channeling an enzyme from its site of synthesis to the regions of the endoplasmic reticulum forming A-granules must be a posttranslational event. The deposition of newly synthesized enzyme is apparently not restricted to special cisternae, all of whose contents obliga-


torily flow to A-granules. Presumably it is the physicochemical properties encoded within the amino acid sequence of the protein, combined with posttranslational modifications that provide the basis for its proper channeling. However, the forces involved in transport and the means by which macromolecules are moved from the endoplasmic reticulum to their proper destinations-and, furthermore, against an apparent concentration gradient-are still unknown (Palade, 1975).

5.3. Compartmentalization of the Synthesis of Specific Proteins at Discrete Sites within the Rough Endoplasmic Reticulum

It has been proposed (Tata, 1971, 1973) that the rough endoplasmic reticulum may serve to segregate topographically or to compartmentalize the synthesis of certain intracellular proteins in particular regions of the cell. This could be achieved in a number of ways, e.g., RER itself may be compartmentalized.

The physical continuity in vivo between RER and certain organelles, most notably nuclei and mitochondria, is well established (Fawcett 1969); it would be thus natural to assume that the ER associated with these two widely different organelles would perform functions affiliated with the requirements of the respective organelles. In vivo fragmentation of part of the endoplasmic reticulum system during mitosis has been observed; the nuclear-associated ER becomes irregular at the end of prophase and then breaks down into vesicles and bilamellar fragments. As cell division proceeds, these fragments become almost indistinguishable from the rest of the endoplasmic reticulum (Fry, 1977). In plant cells RER is found concentrated near the cell plate only during cell division (Porter and Machado, 1960).

At a more complex level, the ER could also provide a network for the nonrandom segregation of specific mRNAs by creating microenvironments for different populations of polysomes that synthesize different classes of proteins. If true, it might be expected that heterogeneous fractions of rough endoplasmic reticulum would exhibit heterogeneity in their mRNA content. To investigate this problem, Shore and Tata (l977a,b) used two subfractions of RER obtained from liver-one of these was termed "rapidly sedimenting endoplasmic reticulum" (RSER), and the other rough microsomes. Translation of mRNAs isolated from these subfractions, in a heterologous cell-free system followed by specific immunoprecipitation of labeled polypeptides, indicated that whereas mRNA for a secreted protein (albumin) was distributed about equally between the two fractions of RER, mRNAs coding for proteins destined for the mitochondrion are relatively enriched in the rough microsomal fraction. It was concluded that at least one class of cytoplasmic mRNAs-


those coding for polypeptides destined for transport to the mitochondrionmay be segregated or compartmentalized preferentially in one fraction of RER within the cell, whereas mRNA coding for a secreted protein (albumin) is not. To rationalize this finding, Shore and Tata (1977b) and Meier et al. (1978) proposed that compartmentalization of cytoplasmic synthesis of mitochondrial proteins on ER cisternae closely apposed to mitochondria could be an important aspect of the mechanism whereby the endocellular membrane system discriminates between polypeptides destined for transport to different cellular loci.

Using a mouse plasmacytoma cell line (MPC-II cells), which produces only light chain immunoglobulin, Pryme (1974a) studied the synthesis of this protein in two fractions of the rough endoplasmic reticulum. These two fractions presumably have different intracellular localization since one (generally called the microsomal fraction) consisted of membranes released to the postnuclear supernatant (fraction I) following cell disruption by nitrogen cavitation, whereas the other contained membranes that remained associated with the nucleus (fraction 2). By short pulse labeling of the MPC-ll cells and immunoprecipitation of the light chain obtained from the different fractions, Pryme showed that the light chain is primarily synthesized in fraction I. It was concluded that there is a difference in function between the two fractions of membranes and a compartmentalization of the mRNA that codes for light chain. The biological sense remains to be elucidated.

In experiments designed to examine more closely the content of fraction I, Svardal and Pryme (1978), using discontinuous sucrose gradient centrifugation, separated this microsomal fraction into three subfractions termed heavy rough (HR), light rough (LR), and smooth (S) microsomes. MPC-II cells were labeled with [3H] leucine, and nascent polypeptide chains were released from polysomes isolated from HR and LR microsomes, by puromycin/KCl treatment, and analyzed for light-chain immunoglobulin content by an immunoprecipitation technique (Pryme et al., 1973). Polysomes from HR microsomes were found to contain about four times more light-chain polypeptides than those from LR microsomes. These preliminary results suggest that the HR, in comparison to the LR fraction, is enriched in mRNA coding for immunoglobulin light chain, adding support to the microenvironment theory.

In order to establish the site of synthesis of proteins glycosylated at the polysomallevel in MPC-Il cells, Pryme and Svardal (1978) have investigated the incorporation of [3H]glucosamine into membrane-bound polysomes of nuclear-associated endoplasmic reticulum, microsomes, and free polysomes. After short-term incubation, nascent polypeptide chains on membrane-bound polysomes in the former fraction were labeled to a much greater extent than were those in microsomes or free polysomes, suggesting a compartmentalization of the synthesis of proteins glycosylated at an early stage in a specific region of the endoplasmic reticulum system.


MPC-ll cells grown in suspension culture at different densities (6-23 X 105 cells/ml) show considerable differences with respect to both cell-cycle distribution profiles and relative amounts of rough microsomal subfractions (Svardal and Pryme, 1980b). The variability of appearance of HR and LR microsomal subfractions was not merely caused by nutritional differences between the cultures. The results indicate that there is a direct correlation between phase of cell cycle and both amount and relative distribution of rough microsomal subfractions during the cell cycle. From Table III it can be seen that as the S/G2 + M ratio decreases a simultaneous decrease was observed in the amount of HR membranes while the amount of LR material increased. There was little alteration in the amount of S membranes. Using the same cell line, Garatun-Tjeldst0 et al. (1976) demonstrated that immunoglobulin light chain synthesis was maximal in late G,/early S phase. From the results presented in Table III it is obvious that cultures with a high S/G2 + M ratio contained large amounts of HR microsomes and this fraction has been shown to be about four times richer in light chain polypeptides than the LR fraction (see above). There was therefore an apparent correlation between phase of cell cycle, synthesis of light chain, and appearance of HR microsomes. One can thus speculate that ER microenvironments may vary in amount and number according to the specific protein species being synthesized during the various phases of the cell cycle. Abraham et al. (1973) also showed that immunoglobulin light chain synthesis was maximal in late Gdearly S phase, and Svardal and Pryme (1980b) have shown that cultures with a high S/G2 + M cell cycle distribution ratio were rich in HR microsomes, whereas those with lower ratios contained decreased amounts of this fraction, but showed increased amounts of LR microsomes (the S fraction remained extremely constant throughout).

Table III Cell Cycle Distribution and Microsomal Subfractions'

Ratio between Cell cycle distribution subfractions

SIG, + M G1 S G,+ M ratio HR LR S

30 59 11 5.4 0.41 0.18 0.41 38 49 13 3.8 0.15 0.42 0.43 31 48 21 2.3 0.10 0.51 0.39

"Cell cycle distribution percentages were obtained from cytofluorometric analysis of MPC-II cell populations. ['H) Choline labeled microsomal subfractions were isolated by sucrose gradient centrifugation (Svardal and Pryme, 1978) and the ratio of radioactivity in the subfractions was calculated.

154 Asbjiirn M. Svardal and Ian F. Pryme

There was therefore an apparent correlation between phase of cell cycle, synthesis of light chain, and relative appearance of HR microsomes. One can thus speculate that ER microenvironments may vary in amount and number according to the specific protein species being synthesized during the various phases of the cell cycle. Abraham et al. (1973) demonstrated that the maximum amount of membrane-bound polysomes in MPC-l1 cells was found in the late Gt/early S phase and this is consistent with the fact that light chain immunoglobulin synthesis in these cells also occurs primarily in late G 1/ early S phase (Garatun-Tjeldst0 et al., 1976). However, in other phases of the cell cycle the level of membrane-bound polysomes did not decrease to the same extent as did the synthesis of light chain. This supported the idea that not only proteins for export from the cell are synthesized on membrane-bound polysomes, and furthermore suggests that different proteins are synthesized on membrane-bound polysomes at various times during the cell cycle.

Although light chain immunoglobulin synthesis has been shown to be maximal in late GI/early S phase in MPC-ll cells (Garatun-Tjeldst0 et ai., 1976), there is apparently little variation in the template activity of poly (A)rich light chain mRNA during the cell cycle (Abraham et al., 1976). These results, together with the observation that salt-extractable factors from membrane-bound polysomes in GI phase were able to stimulate light chain synthesis by membrane-bound polysomes in the G2 phase of the cell cycle, in contrast to factors from the G2 phase (Pryme, 1974b), suggest a translational control of light chain immunoglobulin synthesis. In conclusion, therefore, there is suggestive evidence indicating a differential translation of mRNA species on different endoplasmic reticulum microenvironments according to stage of cell cycle.

6. CONCLUSIONS

Protein synthesis on membrane-bound polysomes occurs on rough and not on smooth membranes. The proteins synthesized can be divided into at least three groups: (1) those to be secreted from the cell (e.g., albumin, immunoglobulin), (2) those destined to become member constituents of membranecontaining organelles (e.g., endoplasmic reticulum, mitochondria, lysosomes), and (3) those released from the membrane to the cytosol.

Proteins to be secreted from the cell are vectorially discharged into the lumen of the RER, migrate through the SER, and are ultimately accumulated in the Golgi apparatus before being transported to the cell surface.

Polyribosomes may interact with membranes in several ways: (1) by direct binding of 60 S subunits to integral membrane proteins (ribophorins I and II), (2) by an interaction between a hydrophobic (signal) sequence of amino acids emerging from the 60 S subunit and specific proteins in the membrane, and


(3) through a direct association between the mRNA and membrane proteins. Posttranslational modification of proteins occurs in various regions of the endoplasmic reticulum. For example, during hydroxylation and glycosylation in the rough endoplasmic reticulum, there is presumably some form of interaction between nascent polypeptide chains on membrane-bound polysomes and membrane-bound enzymes within the ER. The three-dimensional folding of the completed part of the nascent polypeptide chain which has emerged into the lumen of the ER may also afford attachment or anchorage of membranebound polysomes to the ER.

There is evidence to suggest that the ER may serve to provide microenvironments where the compartmentalization of synthesis of specific proteins occurs. This raises interesting questions concerning how the appropriate mRNA species segregate within predetermined areas of the RER system. An elucidation of the mechanisms involved requires further experimentation.

Light chain immunoglobulin synthesis occurs on membrane-bound polysomes and is apparently compartmentalized within a distinct fraction of the RER. As the rate of synthesis varies according to phase of the cell cycle, one may expect to find that the composition of microenvironments within the ER will vary with respect to type of proteins being synthesized on membranebound polysomes at anyone time during the cell cycle. The relative amounts of RER and SER are known to vary according to a variety of biochemical and physiological parameters, and thus a variation during the cell cycle would not seem unlikely.

ACKNOWLEDGMENTS

We thank the Norwegian Research Council for Science and the Humanities (NA VF) for financial support (Grant C-1l-14-8). Ms. Elin Kalvenes is thanked for help in preparing the manuscript.

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Chapter 4

Biogenesis of Peroxisomes and Glyoxysomes

J. Michael Lord School of Biological Sciences University of Bradford Bradford BD? lOP, W. Yorkshire, U.K.

1. INTRODUCTION

In 1954 Rhodin described a morphologically distinct class of organelles in the cytoplasm of mouse kidney cells, which he termed microbodies. These organelles are characterized by a single limiting membrane and a finely granular matrix. In addition they often contain a denser inner core or nucleoid. Ultrastructural studies have established that organelles with this typical morphology are widely distributed, although not ubiquitously present, in eukaryotic cells.

During the 1960s, extensive cellular fractionation and biochemical studies by de Duve and his colleagues demonstrated that, in rat liver and kidney cells, urate oxidase, D-amino acid oxidase and catalase were present in a novel cellular particle (de Duve et ai, 1960; Baudhuin et ai., 1964). This organelle was named the peroxisome (de Duve, 1969), and it has been confirmed that the morphological counterpart of the peroxisome is the microbody (Allen and Beard, 1965; Leighton et ai., 1960).

Microbodies have now been isolated from a variety of cells, generally by isopycnic sucrose density gradient centrifugation, and their biochemical properties have been analyzed. These properties have been described in several review articles (de Duve and Baudhuin, 1966; Hruban and Recheigl, 1969; Tolbert, 1971; Vigil, 1973; Mazliak, 1975) and published symposia (Hogg; 1969; Novikoff and Allen, 1973). In spite of differences in enzymic constituents and proposed metabolic role, microbodies isolated from animals, plants, and unicellular eukaryotes are characterized by their ability to produce and decompose hydrogen peroxide. Catalase is a ubiquitous enzyme constituent and has

171

172 J. Michael Lord

proved a useful marker both for locating microbodies during cellular fractionation experiments and, following cytochemical staining, for identifying microbodies in situ. On the basis of biochemical criteria, microbodies are now more generally termed peroxisomes or, in the case of specialized plant microbodies, which also contain the glyoxylate cycle enzymes, glyoxysomes.

In this review emphasis will be placed on the properties and functions of peroxisomes in mammalian liver and plant leaves and of glyoxysomes in fatmetabolizing plant seeds. It is tentatively assumed that the mechanism of microbody assembly will have features common to all eukaryotic cell types. Details of the assembly process have not been experimentally elucidated at present and current models, therefore, remain speculative.

2. MORPHOLOGY AND TOPOGRAPHICAL RELATIONSHIP TO OTHER CELLULAR ORGANELLES

Microbodies vary somewhat in size and shape, but generally appear to be spherical with diameters ranging from 0.3 to 1.5 11m. They are delimited by a single membrane, 60-80 A thick, which encloses a finely granular matrix. In addition, many microbodies contain an electron-dense inner core or nucleoid. The nucleoids have a structure characteristic of the cell type and species. In rat liver peroxisomes or castor bean glyoxysomes the nucleoid consists of straight tubules arranged in parallel that give a regular crystalloid structure, whereas in mouse liver peroxisomes the tubules appear as twisted strands. The significance and function of the microbody core is not clear, although it is known to be the site of the enzyme urate oxidase in rat liver peroxisomes (Baudhuin et al., 1965) and has been shown histochemically to be rich in catalase in certain plant cells (Vigil, 1970b; Frederick and Newcomb, 1969).

Following their identification it was originally thought that microbodies were only present in significant numbers in liver and kidney cells and a few other animal and plant cells (de Duve and Baudhuin, 1966). The lack of distinctive morphological features, particularly when nucleoids were absent, probably resulted in microbodies being overlooked during many ultrastructural studies. The advent of a cytochemical staining procedure for the microbody marker enzyme catalase rapidly extended the list of microbody-containing cells; cells can be stained for catalase using 3,3'-diaminobenzidine. The peroxidative activity of the enzyme on the colorless substrate converts it to an insoluble brown reaction product seen under the electron microscope as an osmiophilic electron-dense deposit largely found within microbodies (Novikoff and Goldfischer, 1968; Fahimi, 1969). The staining procedure has now established microbodies as familiar cytoplasmic inclusions in a wide range of animal and plant cells. Certain cells, such as intestinal absorptive cells, contain numer-

Biogenesis of Peroxisomes and Glyoxysomes 173

ous small (150-250 nm in length) catalase-positive organelles termed microperoxisomes (Novikoff et aI., 1973).

Observations of microbody-containing cells have established certain features concerning the topographical relationship between microbodies and other cellular organelles. This relationship may be indicative of the cellular origin of microbodies and, in plant cells, of their metabolic roles. Particularly striking has been the close spatial relationship observed between microbodies on sections of the endoplasmic reticulum (ER). Indeed, direct continuities between the ER and the microbody membrane are frequently encountered. In the case of microperoxisomes, continuities with the ER are so common that Novikoff and Shin (1964) have suggested that some microbodies may maintain a longterm, perhaps permanent, functionally important connection with the ER.

Both rough and smooth ER are seen adjacent to microbodies with direct membrane continuity usually to smooth ER. Ribosomes attached to the cytoplasmic surface of the microbody membrane have not been reported.

Plant peroxisomes and glyoxysomes show a close association with other cellular organelles that can be readily correlated with their functional metabolic roles. Photosynthesizing plant cells catalyze a process known as photorespiration. Glycolic acid, which is produced as a photosynthetic by-product within the chloroplasts, is known to be the major photorespiratory substrate (Tolbert, 1971). Most of the enzymes of the photorespiratory pathway, sometimes known as the glycolate pathway, are exclusively located within leaf peroxisomes. The first requirement of this pathway is that the substrate, glycolic acid, must be transferred from the chloroplasts into the peroxisomes. This metabolic interplay between these organelles is facilitated by the close spatial association between them in photosynthesizing cells. Quite often the peroxisome is seen to be appressed to sections of the chloroplast outer membrane (Frederick and Newcomb, 1969). Another feature of the glycolate pathway is that one of its intermediates, glycine, is apparently further metabolized in mitochondria, presumably after its direct transport into this organelle from the peroxisome. Electron micrographs have often shown a single peroxisome appressed to both a chloroplast and a mitochondrion. Under such circumstances the peroxisomes may lose their characteristic spherical shape by assuming the contours of the organelles with which they are associated.

Glyoxysomes are present in fat-metabolizing plant seeds. During germination such seeds catalyze a massive transformation of the stored triglycerides into carbohydrates. The glyoxylate cycle enzymes playa key role in the gluconeogenic pathway and, together with enzymes of the iJ-oxidation sequence, are exclusively located in the glyoxysomes (Beevers, 1969). The function of the glyoxysomes in gluconeogenesis from fats probably accounts for certain wellestablished morphological features. In higher plant tissues, glyoxysomes are only found in germinating fat-storing seeds-their cellular numbers increase

174 J. Michael Lord

rapidly when gluconeogenesis begins, and they are degraded when fat utilization is completed. Fatty seeds store triglycerides in single-membrane delimited spherosomes. During gluconeogenesis from fat the newly formed glyoxysomes become closely associated with the spherosomes. Often several glyoxysomes have been observed appressed to a single spherosome (Vigil, 1970a).

The general morphology of animal and plant microbodies is illustrated in Figures 1 and 2.

3. BIOCHEMICAL PROPERTIES AND METABOLIC ROLES

3.1. Liver Peroxisomes

Peroxisomes, as the name implies, are cellular sites for hydrogen peroxide metabolism. The classic studies of de Duve established basic biochemical criteria that now function as a general definition of peroxisomes-the association, in a single-cell organelle, of one or more hydrogen peroxide-generating oxidases with an excess of catalase. In peroxisomes isolated from rat liver the characteristic oxidases are urate oxidase, D-amino acid oxidase, and L-a-hydroxy acid oxidase (de Duve, 1969).

Urate oxidase is the most active oxidase in rat liver peroxisomes where it is located in the crystalline cores; it is not a constant hepatic peroxisomal constituent, however, as it is absent from the liver of several species, including man (Afzelius, 1965; Sknitka, 1966). Glycolate and L-lactate are the substrates most readily oxidized by the L-a-hydroxy acid oxidase.

Hydrogen peroxide catalytically generated by the oxidases is immediately converted to water by the associated catalase. Catalase is a major protein component of rat liver peroxisomes, where it has been estimated to account for some 15% of the total organellar protein (de Duve and Bandhuin, 1966). Hydrogen peroxide decomposition can occur either catalatically or peroxidatically. (The catalatic reaction achieves the conversion of 2 molecules of H 20 2

into 2 molecules of H 20 and 1 of O2, which the peroxidatic reaction represents the interaction between HzOz and a reduced cosubstrate which generates 2 molecules of H 20 and an oxidized cosubstrate.) It has been calculated that the peroxidatic reaction accounts for 70% of the hydrogen peroxide degraded by rat liver (Chance and Oshimo, 1971). Ethanol and formate are among the physiologically important substrates for the peroxidatic action of catalase, being oxidized to acetaldehyde and carbon dioxide, respectively.

The earliest concepts regarding the physiological function of hepatic peroxisomes were based on the oxidase/catalase association. Peroxisomal respiration is not coupled to any mechanism analogous to the electron-transport chain of mitochondria. Energy released during peroxisomal oxidations is simply lost as heat rather than conserved, in part, as A TP. Peroxisomal respiration

a

b

FIGURE 1. Ultrastructural features of (a) rat liver peroxisomes (micrograph courtesy of Dr. P. B. Lazarow; prepared by Ms. Helen Shio), and (b) a crystalloid-containing spinach leaf peroxisome (micrograph courtesy of Dr:E. H. Newcomb). Bar represents 1 /lm.

a

b

FIGURE 2. The association of microbodies with the ER illustrated by (a) kidney peroxisomes (micrograph taken from Shnitka, 1966), and (b) castor bean endosperm glyoxysomes (micrograph courtesy of D. E. L. Vigil). Bar represents 1 ~m.


must therefore be regarded as a wasteful process from this point of view. The rate of peroxisomal respiration is almost directly proportional to oxygen tension, reflecting the properties of the main peroxisomal oxidases.

One of the earliest suggested functions of peroxisomes was to segregate the hydrogen peroxide-generating oxidases. In this way the potentially toxic hydrogen peroxide would be produced in a membrane-bound cellular compartment, which also contained catalase to effectively degrade the toxic product. The main objection to this theory is that the peroxisomes do not contain all the hydrogen peroxide-generating oxidases present in the cell (de Duve, 1969). For example, xanthine oxidase is present in the cytosol of liver cells (Schein and Young, 1952), whereas monoamine oxidase is found in the outer mitochondrial membrane (Depierre and Ernster, 1977).

Another possible role for peroxisomes suggested by de Duve and Baudhuin (1966) depends on their ability to oxidize cell sap NADH by means of substrate-mediated electron shuttles. The postulated mechanism involved the peroxisomal oxidase and catalase activities coupled to dehydrogenases present in the cytosol, as shown in Figure 3. As far as the coupled oxidation is concerned, small amounts of lactate dehydrogenase have been reported to be present in peroxisomes (McGroarty et al., 1974). However, the peroxisomal membrane is permeable to both lactate and pyruvate, so the peroxisomal oxidase could theoretically couple readily with cytoplasmic lactate dehydrogenase. Nevertheless, the physiological importance of this mechanism for cell sap NADH oxi-

r-----------------------------------------------------------,

Peroxisome

I I I

I I

oxidase catalase :

--~:- --7--~-- -~~~~:- ---7---~---~-~:~- ____ J etha'nol acetaldehyde

n OH

NAD NAD NADH +H+

Cytoplasm

FIGURE 3. The oxidase/catalase association in peroxisomes and its possible involvement in the reoxidation of cell sap NADH.

178 J. Michael Lord

dation has been questioned, since the Km of the peroxisomal oxidase for lactate is much greater than is the lactate concentrations normally found in the tissue (McGroarty et aI., 1974).

The peroxidatic reaction of catalase may result in the oxidation of ethanol to acetaldehyde. The reduction of acetaldehyde back to ethanol by soluble alcohol dehydrogenase could achieve the oxidation of cell sap NADH (Figure 3). The contribution that peroxisome-mediated oxidation of cell sap NADH would have to the total rate of cell sap oxidation, generally assumed to be largely achieved by mitochondria, cannot be accurately assessed at present.

A recent significant addition to our understanding of the metabolic capabilities of liver peroxisomes has been the demonstration of a peroxisomal fatty acid oxidation pathway (Lazarow and de Duve, 1976). It has long been realized that hypolipidemic drugs such as clofibrate, used clinically in the treatment of hyperlipemas (Oliver, 1963; Hellman et al., 1963), cause a striking increase in the cellular number of liver peroxisomes and a doubling of liver catalase activity when administered to rats (Hess et al., 1965; Svoboda and Azarnoff, 1966; Svoboda et al., 1967, 1969). The first indication that acyl-CoA derivatives may be substrates for or end products of hepatic peroxisomal metabolism was provided by Tolbert and co-workers, who identified carnitine acyl transferase as a peroxisomal enzyme (Markwell et al., 1973; 1976). Liver peroxisomes contain both a short-chain carnitine acyltransferase (using acetylCoA as substrate) and a medium-chain transferase (with octanoyl-CoA as the most effective substrate). It was estimated that in the liver cell approximately 52% of the carnitine acetyl transferase was mitochondrial, 34% was in a microsome-enriched membrane fraction, and 14% was peroxisomal. In contrast, the long-chain transferase, carnitine palmitoyltransferase, was shown to be exclusively mitochondrial. It is of interest that in rat kidney cells, not only carnitine palmitoyltransferase, but also more than 90% of the carnitine acetyl transferase, were found in isolated mitochondria; carnitine acetyItransferase was not found in kidney peroxisomes (Markwell et al., 1973). The presence of carnitine acetyl transferase in rat liver peroxisomes and its absence from rat kidney peroxisomes is an interesting example of differences in enzymic composition and probably functional role between peroxisomes in mammalian cells.

Drug-induced hepatic peroxisome proliferation accompanying enhanced rates of serum lipid breakdown suggested that peroxisomes may contain fatmetabolizing enzymes. Lazarow and de Duve (1976) have confirmed that this is, indeed, the case, and it is now established that rat liver peroxisomes contain a fatty acyl-CoA oxidizing system. Glyoxysomes isolated from germinating castor bean endosperm cells, in which the conversion of stored triglycerides into carbohydrate is the dominant metabolic event (Beevers, 1961), were the first microbodies shown to contain the complete enzyme system necessary for f3-oxidation (Cooper and Beevers, 1969; Hutton and Stumpf, 1969). The initial


studies of Lazarow and de Duve (1976) showed that liver peroxisomes were capable of oxidizing palmitoyl-CoA with the concomitant reduction of O2 to HzO z and NAD to NADH. This finding was consistent with the operation of a peroxisomal i1-oxidation sequence, and Lazarow (1978) confirmed that the site of oxidation was indeed the i1-carbon and that acetyl-CoA was the end product of the sequence. Crotonase, i1-hydroxybutyryl-CoA dehydrogenase, and thiolase have now been added to the list of rat liver peroxisomal enzymes (Lazarow, 1978). The peroxisomal i1-oxidation pathway appears to be identical to that initially discovered in castor bean glyoxysomes (Cooper and Beevers, 1969). This pathway is shown in Figure 4 and resembles that of mitochondria, except that the first dehydrogenase transfers its electrons to O2 producing hydrogen peroxide. Peroxisomal fatty acid oxidation is therefore another metabolic event producing the substrate for catalase. The i1-oxidation pathway enzymes have also been reported to be present in the peroxisomal fraction isolated from the unicellular eukaryotes Tetrahymena pyriformis (Blum, 1973) and Euglena gracilis (Graves and Becker, 1974).

The generation of acetyl-CoA in hepatic peroxisomes provides an expla-

FIGURE 4. The reactions of {3-oxidation sequence identified in fatty seed glyoxysomes and rat liver peroxisomes.

t FADXHP2

FADH2 O2

R-CH = C H - CO-SCoA

rH'O

R-C HOH -~,~~~_S CoA

~NADH2 R-CO - C H2-CO-SCoA

~talase

rCOASH

R-CO-SCoA + CH3-CO-SCoA

180 J. Michael Lord

nation for the presence of peroxisomal carnitine acetyltransferase activity discussed above. This enzyme may function in the intracellular shuttle of acetylCoA generated in peroxisomes (and perhaps other acyl-CoA residues) to mitochondria for further oxidative metabolism coupled to energy production.

Several hypolipidemic drugs have been shown to increase hepatic palmitoyl-CoA oxidation in the rat (Lazarow, 1977). Male rats treated with clofibrate, tibric acid, or Wy-14643 [4-chloro-6-(2,3-xylidino )-2-pyrimidinyl thioacetic acid] showed an 11- to 18-fold increase in the capacity of their livers to oxidize palmitoyl-CoA. Although the livers of rats treated with such drugs were used to facilitate the identification of the peroxisomal fj-oxidation sequence, it should also be stressed that the peroxisomes isolated from normal rats also contain fj-oxidation enzymes, although at a much lower level (Lazarow and de Duve, 1976). Clofibrate treatment has also been shown to increase the activity of hepatic carnitine acetyltransferase (Moody and Reddy, 1974; Goldenberg et aI., 1976). It is noteworthy that although clofibrate treatment increases hepatic fj-oxidation activity by approximately one order of magnitude, Lazarow and de Duve (1976) estimated that this accompanied a 2.5-fold increase in peroxisomal protein over the normal liver value and a 53% increase in the activity of peroxisomal catalase. Clofibrate treatment, therefore, does not simply induce a proliferation of peroxisomes with a fixed enzyme complement, but has a differential effect on the rates of synthesis of various enzymes destined to be segregated in the proliferating peroxisomes.

Recently a cyanide-insensitive fatty acid oxidation system has been identified in human liver peroxisomes (Bronfman et at., 1979).

Following sucrose density gradient fractionation of rat liver homogenates, the bulk of the palmitoyl-CoA oxidation detected across the gradient occurs in the peroxisomes and not in the mitochondria Lazarow and de Duve, 1976). Prior to the discovery of the peroxisomal system, fj-oxidation in mammalian cells was generally considered a function of the mitochondria (Garland et aI., 1969; Bressler, 1970). Lazarow (1978) has questioned the significance of mitochondrial fj-oxidation in rat liver cells, particularly because studies in which fj-oxidation activity was originally attributed to mitochondria may have utilized mitochondrial preparations obtained by differential centrifugation and that could, therefore, have been seriously contaminated by peroxisomes. Tolbert and co-workers (Krahling et at., 1978) have confirmed that the bulk of the hepatic palmitoyl-CoA oxidation occurs in peroxisomes after gradient centrifugation; mitochondria were damaged to such an extent that the rates of respiration were very low with succinate and malate, in addition to palmitoylCoA, as substrates in an isotonic assay medium. ADP-stimulated respiration measured as O2 uptake by gradient-isolated mitochondria was readily detected with the above-named substrates using dilute phosphate buffer as the assay medium. fj-Oxidation was assayed polarographically across a collected gra-


dient to determine its distribution between peroxisomes and mitochondria. It was found that the ratio of palmitoyl-CoA oxidation by the mitochondrial fraction relative to the surviving peroxisomes was 3 to 1. More than 60% of the total catalase activity, however, was recovered in the soluble fraction. Assuming that all the catalase is located in peroxisomes (which may not be a valid assumption), it appears that some two-thirds of the total peroxisomes may have been damaged. The ratio of mitochondrial ,a-oxidation to peroxisomal ,a-oxidation in the normal hepatocyte may therefore be closer to 1. This ratio clearly increases in favor of the peroxisomes in the case of clofibrate-treated rats. Regardless of the contribution of peroxisomal ,a-oxidation to the overall rate of fatty acyl-CoA oxidation in rat hepatocytes, it is clear that it represents a major metabolic capacity of the peroxisome. Thus an important physiological role can be attributed to liver peroxisomes in addition to the classic oxidases/ catalase association, and a plausible biochemical explanation of the mechanism by which hypolipidemic drugs lower plasma lipids is apparent.

3.2. Leaf Peroxisomes

Ultrastructural studies have established that most, if not all, eukaryotic plant cells contain microbodies (Mollenhauer et aI., 1966; Frederick et al., 1968). Detailed accounts of their morphology and biochemical properties can be found in several review articles (Tolbert, 1971; Vigil, 1973; Richardson, 1974; Frederick et al., 1975). Accordingly, only the major recognized cellular functions of specialized plant microbodies are discussed here.

It is well established that during photosynthesis many plant cells take up O2 and evolve CO2 in the light simultaneously with the normal O2 evolution and CO2 fixation of the photosynthetic pathway. This process is known as photorespiration, and its magnitude is such that net photosynthesis is markedly reduced; up to one-half the photosynthetic carbon fixation and reducing power of the chloroplasts may be lost (Jackson and Volk, 1970; Hatch et al., 1971). Photorespiration is enhanced by increased O2 concentration in the photosynthesizing plants environment. A small proportion of the O2 used and CO2

released may come from conventional mitochondrial reactions, but by far the greatest part comes from the metabolism of glycolic acid (Tolbert, 1971). Glycolate is a photosynthetic by-product that is formed within the chloroplast by the oxidative degradation of ribulose diphosphate. Oxygen competes with CO2

for the catalytic site of ribulose diphosphate carboxylase/oxygenase and leads to the formation of one molecule of 3-phosphoglycerate (the normal product of the primary photosynthetic carboxylation) and one of phosphoglycolate (Bowes et al., 1971; Andrews et al., 1973). Phosphoglycolate is dephosphorylated within the chloroplast (Yu et al., 1964). The resulting glycolate is then metab-

182 J. Michael Lord

olized by a well-established pathway in which most of the enzymes are located in leaf peroxisomes (Tolbert et ai., 1968). Within the peroxisomes, glycolate is oxidized to glyoxylate by the enzyme glycolate oxidase, which is the major oxidase in leaf peroxisomes. The hydrogen peroxide generated during this oxidation step is degraded by peroxisomal catalase. Glyoxylate is converted to glycine by aminotransferases, which utilize either serine or glutamate as amino donors (Rehfeld and Tolbert, 1972). Glycine apparently leaves the peroxisomes and enters the mitochondria, where it is converted to serine. Serine reenters the peroxisomes, where it is deaminated to hydroxypyruvate by serine: glyoxylate aminotransferase and subsequently reduced to glycerate by NADH-hydroxypyruvate reductase (Tolbert, 1971). The overall result of the peroxisomal glycolate pathway (Figure 5) is to convert two molecules of glycolate into one of glycerate, which can then reenter the gluconeogenic pathways in the chloroplast. A major contributor to photorespiratory CO2 release is the reaction in which two molecules of glycine are converted into one of serine and one of CO2,

Whereas the photorespiratory glycolate pathway apparently represents the major functional role of leaf peroxisomes, other enzymic activities are present in these organelles including malate dehydrogenase (Yamazaki and Tolbert, 1969) and urate oxidase (Huang and Beevers, 1971). Leaf peroxisomes may thus participate in other metabolic processes not directly related to photorespiration.

The widespread occurrence of metabolically active peroxisomes in photosynthetic cells appears to be somewhat puzzling; these specialized plant organelles function in a process (photorespiration) that is clearly wasteful and markedly reduces net photosynthesis. Current concepts suggest, however, that leaf peroxisomal metabolism may represent a salvage mechanism by which some of the glycolate carbon can be directed back into hexose production. Such concepts regard glycolate production as an unfortunate consequence of the dual carboxylase/oxygenase nature of the enzyme ribulose diphosphate carboxylase (Lorimer and Andrews, 1973). Photorespiration may be regarded as an evolutionary accident. The primordial atmosphere is thought to have contained much CO2 but little O2, The evolution of photosynthesis has increased the atmospheric concentration of O2 until competition between O2 and CO2 for ribulose diphosphate carboxylase is significant, and glycolate production is the result. The original response of the organism may have been to excrete the glycolate, as certain algal species still do this (Merrett and Lord, 1973). Faced with the loss of the whole glycolate molecule, a compound formed in increasing amounts with increasing O2 concentration, peroxisomal metabolism is beneficial to the plant cell in retaining some glycolate carbon. In addition to its gluconeogenic role, the glycolate pathway appears to generate ATP during the conversion of glycine into serine (Bird et ai., 1972).

Biogenesis of Peroxisomes and Glyoxysomes

CHLOROPLAST

R~UF:,di- P--_O~2---··p-gIo,ate

~H20-P Sugar ~H20H CHOH COOH GLYCOLATE

eOOH 3- P- GLYCERATE .. ~

CH 20H CHOH

PEROXISOME

COOH GLYCERATE

JOH I

C=O I

COOH HYDROXYPYRUVATE

i~ "'NSAM'"AS"

CH20H I

CHNH 2 I

COOH .. ~

CHO I

COOH GLYOXYLATE

----"',1 CH2NH 2 I

COOH GLYCINE

MITOCHONDRION

183

FIGURE 5. Localization of the major reactions of the photorespiratory glycolate pathway in plant leaves.

184 J. Michael Lord

3.3. Fatty Seed Glyoxysomes

Many plant seeds, such as those of the castor oil plant, Ricinus communis. store carbon reserves in the form of triglycerides. During the early stages of postgerminative growth, these triglycerides are actively converted into carbohydrate, which nurtures the developing seedling until it becomes photosynthetically competent. Because triglycerides can account for up to 70% of the dry weight of fat-storing seeds, gluconeogenesis from fat is the dominant metabolic event during early development. The glyoxylate bypass of the tricarboxylic acid cycle is known to playa key role in gluconeogenesis. Shortly after the discovery of the glyoxylate cycle in microbial cells growing on acetate (Kornberg and Krebs, 1957), Kornberg and Beevers (1957) identified the key enzymes of this cycle, isocitrate lyase and malate synthase, in the fat-storing endosperm tissue of germinating castor bean seeds. Furthermore, these enzymes have only been found with significant activity in plant tissues actively converting fat to carbohydrate (Beevers, 1961). The intracellular location of the key glyoxylate cycle enzymes was initially thought to be mitochondria (Marcus and Velasco, 1960), until Breidenbach and Beevers (1967) demonstrated that they were housed in a distinctive cytoplasmic organelle, which they therefore termed the glyoxysome. These organelles were originally isolated and characterized from the endosperm of castor bean seedlings. Glyoxysomes contain all the enzymes necessary to convert fatty acids, via acetyl-CoA, into succinate. Ultrastructural studies have shown that glyoxysomes appear to be appressed to the lipid-storing spherosomes (Vigil, 1970a). The lipases necessary for the liberation of free fatty acids from the storage triglycerides are present in the spherosomes themselves (Ory et al.. 1968). An alkaline monoglyceride lipase has also been demonstrated in the membrane of castor bean glyoxysomes (Muto and Beevers, 1974). Once the fatty acids have entered the glyoxysomes, they are activated by long-chain fatty acid thiokinase (Cooper, 1971) and oxidized to acetyl-CoA by the {3-oxidation sequence illustrated in Figure 4 (Cooper and Beevers, 1969). Acetyl-CoA is further metabolized via the glyoxylate cycle, the net result of which is the conversion of two molecules of acetyl-CoA into one of succinate. Succinate then leaves the glyoxysomes and enters the mitochondria, where it converted into oxaloacetate via the tricarboxylic acid cycle reactions and ultimately into carbohydrate via phosphoenolpyruvate and reverse glycolysis (Beevers, 1969).

In fat-metabolizing seeds, fatty acid thiokinase and the {3-oxidation enzymes are exclusively located in glyoxysomes. Thus in these tissues the situation is different from that found in rat liver, where the capacity for {3-oxidation is shared by both peroxisomes and mitochondria.

The significance of this compartmentalization defines the role of glyoxysomes in fat-metabolizing tissues; the association of {3-oxidation and the gly-


oxylate bypass in a distinct cytoplasmic organelle ensures that gluconeogenesis occurs at maximum carbon efficiency. Acetyl-CoA avoids the oxidative decarboxylations of the mitochondrial tricarboxylic acid cycle and 75% of the fatty acid carbon is thus recovered in carbohydrate (Canvin and Beevers, 1961). It should be noted that the mitochondria isolated from castor bean endosperm contain a full complement of active tricarboxylic acid cycle enzymes. Enzymes unique to the tricarboxylic acid cycle, such as fumarase and succinate dehydrogenase, are exclusively located in mitochondria; enzymes unique to the glyoxylate cycle, isocitrate lyase and malate synthase, are exclusively located in glyoxysomes; enzymes common to both cycles, such as citrate synthase, aconitase, and malate dehydrogenase, are present in both mitochondria and glyoxysomes. Glyoxysomes also contain catalase, a-hydroxy acid oxidase, and uricase, thus identifying these organelles as specialized peroxisomes (Breidenbach et a/., 1968; Theimer and Beevers, 1971). Other enzymic activities that include allantoinase (St. Angelo and Ory, 1970), hydroxypyruvate reductase (Lord and Beevers, 1972), and glutamate: oxaloacetate aminotransferase (Cooper and Beevers, 1969) have been identified in glyoxysomes, which may therefore have additional cellular functions other than their major role in gluconeogenesis from fats.

Many other plant tissues, e.g., roots, have been shown to contain microbodies in addition to the specialized leaf peroxisomes or fat-metabolizing cell glyoxysomes. Catalase, a-hydroxyacid oxidase, and urate oxidase have been identified as enzymic components of these organelles, which have been described as nonspecialized plant microbodies (Huang and Beevers, 1971).

Unicellular eukaryotic plants, such as green algae, have been shown to contain microbodies in ultrastructural studies, although problems encountered when isolating these fragile organelles from such cells have prevented a detailed biochemical characterization. Eukaryotic microorganisms growing on acetate or metabolically related substrates synthesize the glyoxylate cycle enzymes. These enzymes are probably located in glyoxysomelike microbodies in green algae (Graves et al., 1972; Collins and Merrett, 1975), fungi (Kobr et a/., 1969; Avers, 1971), and protozoa (Muller, 1975).

4. MICROBODY PROLIFERATION

4.1. Liver Peroxisomes

Developmental studies have shown that a proliferation of peroxisomes in hepatic cells occurs during fetal and early postnatal growth (Peters et al., 1963; Tsukada et al., 1978). A similar effect has been observed in regenerating liver cells following partial hepatectomy (Stenger and Confer, 1966).

186 J. Michael Lord

Most of the studies on proliferating hepatic peroxisomes have utilized the livers of animals, in which proliferation has been experimentally induced by the administration of certain chemical agents. Salicylates, including acetyl salicylate, induce a striking (two- to threefold) increase in the number of microbody profiles in cellular sections of rat liver (Hruban et al.. 1966). The most effective and thoroughly studied proliferating agents have been hypolipidemic drugs, particularly ethyl-a-p-chlorophenoxyisobutyrate (CPIB or clofibrate).

In 1964 Duncan and co-workers reported that clofibrate caused a marked hepatomegaly in rats, which was subsequently shown to accompany a largescale proliferation of hepatic peroxisomes (Hess et al .. 1965; Svoboda et al .. 1967, 1969). Peroxisome proliferation was associated with a marked increase in catalase activity resulting from an enhanced rate of synthesis of this enzyme Reddy et al .. 1971), whereas the peroxisomal oxidase activities remained either unchanged or declined (Hess et al .. 1965; Svoboda et al .. 1967; Leighton et al .. 1975). The effects of clofibrate treatment were shown to be sex-dependent, with male rats responding as described above while female rats did not.

Several structural analogues of clofibrate induce hypolipidemia and peroxisome proliferation (Reddy, 1974; Reddy et al .. 1974; Moody and Reddy, 1976), in common with several other compounds structurally unrelated to clofibrate (Hruban et al .. 1974; Reddy and Krishnakantha, 1975).

Experimentally induced peroxisome proliferation seems to offer a useful system for the study of peroxisome biogenesis but, as yet, such systems have failed to produce any definitive evidence in this respect. For the most part these studies have relied on ultrastructural observations that have made any conclusions regarding biogenetic mechanisms somewhat speculative. Legg and Wood (1970) used the cytochemical staining of catalase in clofibrate-treated rat liver to derive the suggestions that (1) peroxisome proliferation may result by a process of fragmentation or budding from preexisting peroxisomes, (2) catalase is synthesized on rough ER, and (3) newly synthesized catalase is not vectorially discharged into the ER cisternae, but rather is directly transported into the peroxisomes following its release from the membrane bound ribosomes. Moody and Reddy (1976) have stressed that peroxisome proliferation is always accompanied, to a lesser extent, by a proliferation of smooth ER. Other compounds, e.g., phenobarbital, induce a proliferation of smooth ER solely (Conney, 1967), which could therefore occur independently of peroxisome proliferation. However, the close association between the smooth ER and peroxisomes, which has been frequently emphasized, suggests that the simultaneous proliferation of these cellular components may be ontogenetically related.

4.2. Leaf Peroxisomes

The ontogeny of leaf peroxisomes has been followed during the greening of etiolated leaves. The achlorophyllous leaves of germinating seedlings that


have not emerged into the light contain small peroxisomes (0.3 Jlm in diameter) associated with the ER. After the leaves have turned green, the peroxisomes are considerably larger (1.5 Jlm) and are associated with chloroplasts (Gruber et al .. 1973). This peroxisomal enlargement is accompanied by a severalfold increase in the activities of peroxisomal enzymes. It has been suggested on the basis of ultrastructural observations that peroxisome enlargement is achieved by preexisting peroxisomes fusing with vesicles derived from the ER (Gruber et al.. 1973). These vesicles could conceivably transport additional enzymic protein to the peroxisomes. The addition of individual enzymes to preexisting peroxisomes via ER-derived vesicles could explain the observation that individual enzymes of leaf peroxisomes have different developmental patterns and are independently regulated (Feierabend and Beevers, 1972a).

Although the light-dependent increase in leaf peroxisomal enzyme activity normally parallels chlorophyll synthesis and an increase in photosynthetic capacity, these events are not mutually dependent, as peroxisomal enzyme synthesis still occurs when chlorophyll synthesis and chloroplast development are inhibited (Feierabend and Beevers, 1972b).

4.3. Fatty Seed Glyoxysomes

Glyoxysomes and their characteristic enzymes are virtually absent in dry seeds. During the first few days of postgerminative growth there is a rapid assembly of these organelles. In the endosperm tissue of castor bean seedlings germinating at 30 0 C, glyoxysome biogenesis occurs during the first 4 days of growth, and subsequently the cellular content of these organelles rapidly declines (Gerhardt and Beevers, 1970). At the developmental stage, when maximum rates of gluconeogenesis from fats are observed, glyoxysomes can account for up to 20% of the total particulate protein present in homogenates. Glyoxysome formation occurs in the absence of cell division and without net protein synthesis, and the organelles can be isolated without significant breakage during cell fractionation (Kagawa et al.. 1973). The messenger RNAs for castor bean endosperm glyoxysomal proteins are transcribed during germination (Roberts and Lord, 1979a) and are translated on newly assembled ribosomes (Roberts and Lord, 1979b). Developmental studies have shown that castor bean endosperm glyoxysome formation during germination is preceded by proliferation of the ER membrane (Lord, 1978).

5. MODELS FOR THE SYNTHESIS OF MICROBODY COMPONENTS AND THEIR TRANSFER TO THE ORGANELLES

In considering microbody formation, current views hold that all the components of such single-membrane-delimited organelles are synthesized outside

188 J. Michael Lord

the organelles themselves. Microbody biogenesis involves the assembly of these components into the structurally complete and functionally active organelle. In order to simplify the present discussion, these components will be considered to be the microbody membrane consisting of lipids and proteins, and the microbody matrix consisting of the enzymic proteins characterizing the organelle. At the present time detailed analyses of the structural components of microbody membranes and matrices have not been documented, so this view may be an oversimplification.

5.1. Membrane Lipids

The microbody membrane is assumed to be a typically structured cytoplasmic membrane and, as such, its major lipid components are phospholipids. Studies using a variety of eukaryotic tissues have established that the enzymic reactions of phospholipid synthesis, particularly those of the final steps, occur in the ER (Wilgram and Kennedy, 1963; McMurray and Dawson, 1969). (The enzymic synthesis of certain phospholipids by mitochondria is not considered here, because the lipid products of such reactions, e.g., cardiolipin, are probably not present in microbody membranes.) The question to be considered here is: How are these phospholipids transferred from their site of synthesis in the ER to their ultimate destination in the microbody membrane? Two mechanisms can be envisaged-one involving phospholipid exchange and the second involving membrane flow.

5.1.1. Phospholipid Exchange Proteins

Although the enzymes responsible for the final steps in the assembly of membrane phospholipids are confined to the ER, intact tissues supplied with radioactive phospholipid precursors rapidly label these lipids in other cellular membranes, in particular the mitochondria (Wirtz and Zilversmit, 1969; Jungawala and Dawson, 1970). Furthermore, the half-lives of the major phospholipid classes in the microsomal, inner mitochondrial, and outer mitochondrial membranes are similar, suggesting that the various phospholipid pools may be in equilibrium (Wirtz and Zilversmit, 1968). These observations have been explained by the discovery of specific phospholipid exchange proteins in the cytoplasm of a variety of eukaryotic cells (Wirtz and Zilversmit, 1968; McMurray and Dawson, 1969). The biological activity of these proteins is assayed by their ability to catalyze the exchange in vitro of phospholipids between different membrane preparations, particularly microsomes and mitochondria. The purified exchange proteins exhibit specificity with respect to individual phospholipids; in some cases the specificity is absolute (Kamp et aI., 1973), while in others more than one phospholipid class appears to be trans-


ferred (Helmkamp et aI., 1974). The biochemical properties of the exchange proteins and their possible role in the assembly of the mitochondrial membrane has been reviewed by Wirtz (1974). The significance of the exchange proteins with regard to membrane biogenesis depends not on their ability to catalyze a one-for-one exchange between membranes, but on their ability to achieve a net transfer of phospholipid from the ER to another membrane. In this way, newly synthesized phospholipids could be transferred to the membrane of an organelle, such as the microbody, thus expanding this membrane by increasing its lipid content. The significance of such a mechanism cannot be assessed at present. An apparent problem is that phospholipid exchange proteins can only transfer lipids to the cytoplasmic side of the membrane bilayer exposed to the proteins (Johnson et al .. 1975). Transfer of phospholipids from the cytoplasmic monolayer to the opposing monolayer (flip-flop), which is necessary to expand the membrane by adding phospholipids transferred individually, appears to be too slow to permit exchange proteins to synthesize intracellular membranes at physiologically significant rates. Indeed, the low rate of flip-flop permits the opposing monolayers in a membrane to maintain the lipid and protein asymmetry which characterizes all cellular membranes.

5.1.2. Membrane Flow

Although the rate of flip-flop of phospholipids between opposing monolayers is slow, the lipids are free to diffuse laterally in the fluid monolayer, and they apparently exchange positions very rapidly. Once the phospholipid has been synthesized by the enzymic ER membrane protein and has been incorporated into the monolayer adjacent to this protein, it may thus be free to flow through the fluid hydrophobic phase. In this way any intracellular membranes in direct continuity with the ER, either permanently or temporarily, could aquire additional phospholipids while the continuity exists. This consideration may be significant, as the microbody membrane is one of the organelle membranes reported to have direct continuity with the ER. Although it has been suggested that continuity between the ER and peroxisomal membrane may be a relatively long-lasting or even permanent association (Novikoff and Shin, 1964), it seems probable that this continuity is transient. Membrane proliferation may thus be confined to the ER, with the expanding ER membrane giving rise to other cellular membranes, such as the peroxisomal membrane, by a process of vesiculation. The direct continuity occasionally seen between the ER and peroxisomal membrane may be a visualization of the vesiculation step. This model implies that specialized areas of the ER may exist where budding off of membranes generates nascent peroxisomes. These ER regions are presumably specialized, or are actively differentiating prior to vesiculation, because the peroxisomal membrane is not identical to the ER membrane in its

190 J. Michael Lord

biological activity. For example, the peroxisomal membrane does not have the capacity to synthesize phospholipids. The budding off of peroxisomes from the ER is not necessarily a step that defines the ultimate size of the isolated organelles. This step might generate an immature peroxisome, which could increase the surface area of its membrane by fusing with other membrane vesicles subsequently derived from the ER. The budding of vesicles from one membrane and their fusion with another membrane is the basis of the membrane flow hypothesis. The biological significance of membrane flow has been elegantly demonstrated in secretory cells by Palade and his colleagues (Palade, 1975).

One of the strongest arguments supporting the membrane-flow hypothesis is the established lipid and protein asymmetry of opposing monolayers in cellular membranes. If we consider that each intracellular organellar membrane may be individually assembled by a process not involving any preassembled membrane, it is necessary to propose a self-assembly hypothesis for each particular membrane. In this case the various membrane components would be individually transferred from their site of synthesis to the site of membrane assembly, phospholipids by means of specific phospholipid exchange proteins, and integral membrane proteins by some unknown mechanism. If these membrane components could thus be collected together in some fashion, it is reasonable to assume they would form a membrane vesicle under suitable environmental conditions. Indeed, isolated cellular membranes can be dispersed into their individual protein and lipid components by disrupting the bilayer with high concentrations of detergents, these hydrophobic molecules being effectively separated into detergent micelles. If the detergent is subsequently removed, the proteins and lipids spontaneously reform a membrane vesicle. Such reconstituted membranes have one vital difference from the original biological membrane-The reconstituted membranes are almost always symmetrical and the original characteristic asymmetry of the cellular membrane is lost. The symmetry of reconstituted membranes occurs because the formation of the lipid bilayer and the insertion of the membrane proteins take place simultaneously. The asymmetry of cellular membranes occurs because components such as integral membrane proteins are given their orientation at the time they are inserted into the membrane. It is therefore improbable that an asymmetric organellar membrane such as the peroxisomal membrane could be formed by a self-assembly process even if the individual lipid and protein components of that membrane were available on site. Self-assembly of the peroxisomal membrane would require that the organelle could both synthesize its own membrane components and assemble them in a stepwise manner. The peroxisome clearly lacks this capacity. Integral membrane proteins must be synthesized and inserted into a preexisting membrane that has a defined sidedness; the ER is the cellular membrane with this capacity. This consideration does not eliminate the possibility that the peroxisomal membrane could be altered


in some way, either by the addition of further membrane components such as peripheral proteins, or by the removal or modification of existing components.

One further aspect of the synthesis of phospholipids within an expanding membrane such as the ER must be considered. The enzymes responsible for these syntheses are themselves integral components of the membrane. It is not clear whether these enzymes insert the newly synthesized phospholipids into a single monolayer or into both sides of the bilayer. If some of the precursors for phospholipid synthesis cannot cross the hydrophobic bilayer completely, then lipid synthesis might be expected to occur largely in the cytoplasmic monolayer. Because the expanding membrane is clearly a bilayer, it is essential that phospholipids be added to both monolayers. As mentioned earlier, the rate of flip-flop across bilayers is very low. It has therefore been proposed that the growing membrane contains proteins which facilitate the transfer of lipids between mono layers (Bretscher, 1973). This protein-mediated lipid transport must also achieve the established asymmetric distribution of the various phospholipid classes between these monolayers (Nilsson and Dallner, 1975). It seems possible that this asymmetry is not actively maintained, but that it represents a thermodynamic equilibrium between the molecules in the bilayer. Certain phospholipids may have a lower free energy on one side of the membrane than they do on the other.

5.2. Membrane Proteins

There are two classes of protein associated with membranes. Integral proteins are molecules embedded in the lipid bilayer with regions exposed on both sides of the membrane. Peripheral proteins are not integrated into the bilayer but occur at one surface or the other where they are bound to the exposed portions of integral proteins. All integral proteins are asymmetrically inserted into the membrane, thereby giving it sidedness. Peripheral proteins are also asymmetrically distributed being associated with only one side of the membrane.

In considering a mechanism for the synthesis of microbody membrane proteins, the implications of results obtained during studies into the synthesis of proteins in other biological membranes will be applied. One of the most fruitful systems examined to date has been the membrane of the vesicular stomatitis virus (VSV). The viral membrane has the lipid composition of the hostcell membrane, but it incorporates only proteins encoded by the viral genome. There are two such proteins-an integral membrane glycoprotein, designated G, which is largely exposed on the external surface of the host plasma membrane and becomes the viral spike (McSharry et al .. 1971), and a second protein, the matrix protein M, which becomes a peripheral protein on the cytoplasmic side of the host plasma membrane and of the mature viral membrane.

192 J. Michael Lord

Several groups of workers have established that the mRNA for the M protein is largely translated on free cytoplasmic ribosomes and that the product is released as a soluble cytoplasmic protein prior to its incorporation into the virion as a peripheral membrane protein (Morrison and Lodish, 1975; Toneguzzo and Ghosh, 1975). In contrast, the glycoprotein G is exclusively synthesized on membrane bound ribosomes (Grubman et ai., 1975; Morrison and Lodish, 1975.) [Free and bound ribosomes are functionally different in that they synthesize proteins destined to have different ultimate cellular locations; otherwise, the two classes of ribosomes are structurally identical and indeed are interchangeable (Blobel and Dobberstein, 1975).] The closely coupled synthesis, membrane insertion, and glycosylation steps for the G protein have recently been elucidated in a series of elegant experiments. The first 30 codons of the mRNA-encoding G direct the synthesis of an amino-terminal signal sequence (Blobel and Dobberstein, 1975) which identifies the growing polypeptide as one destined to be inserted into the rough ER membrane. Once the signal sequence emerges from the ribosome, it binds to specific receptors on the ER membrane, thereby causing the attachment of the translating polysomes to the membrane. The ribosome-membrane junction creates the correct topographical conditions for the transfer of the elongating polypeptide through the lipid bilayer (Katz et ai., 1977). The signal sequence is removed by a protease, presumably associated with the luminal surface of the ER membrane, as the polypeptide is extruded through. The growing polypeptide passes into the lumen, where it is folded as the initial step toward assuming its final conformation. At this stage two identical carbohydrate chains, which contain N-acetylglucosamine and mannose residues and have been preformed by enzymes located in the ER membrane (Waechter and Lennarz, 1976), are added to the polypeptide (Rothman and Lodish, 1977). This core glycosylation step therefore occurs cotranslationally. At some stage toward the end of the translation of the G protein mRN A, the polypeptide becomes bound to the ER membrane, with some 30 amino acids at the carboxyl terminus remaining in the cytoplasm. Thus when the complete glycoprotein folds into its final conformation it is held in the membrane in an asymmetric orientation with most of the protein and all the carbohydrate exposed on the luminal side of the membrane, and with a short fragment at the carboxyl terminus exposed on the cytoplasmic side (Katz and Lodish, 1979). After this cotranslational insertion into the ER membrane, the protein remains an integral membrane protein with a fixed orientation as it is transported via the Golgi apparatus to the plasma membrane, possibly via vesicles that bud off from one membrane and fuse with another. Certain posttranslational modifications occur during cellular transport, such as the addition of terminal sugar units in the Golgi apparatus (Hunt and Summers, 1976). The vital step of membrane insertion, however, occurs cotranslationally.

Although the synthesis of integral proteins of intracellular membranes,


with certain exceptions, has not been studied in similar detail to the VSV G protein, it is probable that their synthesis and membrane insertion will be mechanistically similar. It is generally assumed that mRNAs for such membrane proteins are translated on bound ribosomes and this has been verified in certain cases, e.g., cytochrome bs (Omura, 1973). This assumption will be maintained in the present discussion: Integral proteins of the microbody membrane are considered to be synthesized on rough ER and inserted into the ER membrane.

The site of synthesis and immediate fate of peripheral membrane proteins may vary depending on which side of the membrane they become associated. Peripheral proteins associated with the cytoplasmic face of the membrane could be synthesized on free polysomes and released into the cytoplasm before their ultimate interaction with the membrane occurs. In contrast, peripheral proteins associated with the luminal face must themselves be passed through the membrane, and this step is considered to occur cotranslationally. The segregation step was initially explained for secretory proteins by Blobel and Dobberstein (1975). Their results led to the formation of the signal hypothesis, a modified account of which was briefly given above for the synthesis of VSV membrane glycoprotein, and details of which can be found elsewhere (Blobel and Dobberstein, 1975). Peripheral membrane proteins that must cross the membrane may be coded for by mRNAs, which contain a signal sequence that directs the ribosome to the membrane. Their synthesis would proceed exactly as described for integral membrane proteins, except that they would not become anchored in the membrane but would pass right through it. Subsequently they could become associated with the luminal face of the membrane.

It should be emphasized that in addition to being a site of protein synthesis, rough ER is also the site of enzymes that catalyze the core glycosylation of proteins (Czichi and Lennarz, 1977). Since core glycosylation is considered to be a cotranslational process (Kiely et al., 1976; Katz et al., 1977), it follows that glycoproteins, particularly membrane-bound or secretory proteins, must be synthesized on rough ER.

5.3. Implications for Microbody-Membrane Biogenesis

The microbody membrane is assumed to be initially fabricated in, and hence directly derived from, the ER. Phospholipids are synthesized by integral enzymic proteins present in the ER membrane. Integral microbody-membrane proteins are synthesized on rough ER and cotranslationally inserted into the membrane. Peripheral proteins associated with the matrix surface of the microbody membrane are segregated by the membrane during synthesis on rough ER. If any of these membrane proteins are glycosylated, core sugars would be added cotranslationally. Transfer of the lipid and protein components of the

194 J. Michael Lord

microbody membrane from their site of synthesis in the ER occurs with these components already inserted into, and asymmetrically orientated within, a largely completed membrane structure. This transfer would be achieved by membrane budding from the ER in the form of a closed vesicle. This closed vesicle would be a nascent microbody.

5.4. Microbody Matrix Proteins

Ribosomes are not seen in animal or plant microbodies examined under the electron microscope. Biochemical analysis has established that microbodies do not possess unique nucleic acids. The RNA and DNA that may be detected in isolated microbody preparations are contaminants, and have been identified as such in castor bean endosperm glyoxysomes (Douglass et al .. 1973). A claim that glyoxysomes isolated from pine seedlings were able to synthesize proteins when incubated with amino acids (Ching, 1970) has not been substantiated. The enzymic proteins of the microbody matrix are the translational products of polysomes situated outside the microbodies themselves. In order to understand the mechanism of matrix protein synthesis and segregation it is necessary to establish the intracellular site of the ribosomes involved in their formation. Once synthesized in the cytoplasm, these proteins must cross a membrane before reaching their final destination. There are various possible modes of synthesis and transport of microbody proteins, and they are discussed below. The importance of the various models has not been experimentally established in the case of microbody proteins and, accordingly, selected evidence supporting their roles in the synthesis of proteins housed in other types of cellular organelles is cited.

5.4.1. Synthesis on Rough ER

The role of membrane-bound ribosomes in the synthesis and segregation of proteins has been elegantly demonstrated in the case of secretory proteins (Palade, 1975). The mechanism of transport across the ER membrane has been formulated into the signal hypothesis (Blobel and Dobberstein, 1975). An important feature of this system is that membrane transport is a cotranslational step. During transport, the amino-terminal signal sequence, which directs the translating ribosome to the membrane, is cleaved off by an ER protease. Cotranslational segregation of microbody matrix proteins into the ER cisternae by a mechanism analogous to that for secretory proteins may occur. Intracisternal sorting may lead to these proteins accumulating in specialized sections of the ER where vesiculation could occur, a step that would release the enzymes into the cytoplasm trapped within the membrane bound microbody. The translation of mRNAs for microbody proteins in a cell-free system devoid of microsomal membranes might generate preproteins some 1500-3000


daltons larger than the corresponding mature microbody proteins. Although the amino-terminal signal sequence seems to be a feature of proteins destined for transmembrane translocation, it is not necessarily essential. Hen ovalbumin mRNA does not encode a cleavable signal sequence (Palmiter et at., 1978), although it has been suggested that the amino-terminal sequence of mature ovalbumin acts as a functional equivalent of a signal sequence and effects the cotranslational segregation of the protein into the cisternal space (Lingappa et at., 1978). Recently evidence has been presented that ovalbumin contains an internal signal sequence (Lingappa et at., 1979).

5.4.2. Synthesis on Organelle-Bound Ribosomes

The synthesis of organellar proteins on ribosomes bound to the cytoplasmic surface of the organelle membrane would ensure direct insertion of the nascent proteins into the organelles. Evidence for the presence of cytoplasmic ribosomes on organelle membranes has only been documented in the case of yeast mitochondria (Butow et at., 1975). In a series of experiments, Butow and his colleagues have presented evidence that the 80 S ribosomes attached to yeast mitochondria are not artifacts arising during cell fractionation and that these polysomes vectorially discharge their translational products into mitochondria (Kellems et at., 1974, 1975). On the other hand, cytoplasmic ribosomes have not been found attached to mitochondria from other organisms or attached to other cellular organelles including microbodies. In castor bean endosperm cells, organelles with similar size and general morphology to glyoxysomes and bearing attached ribosomes have been described. These organelles have been designated ricinosomes (Mollenhauer and Totten, 1970) or dilated cisternae (Vigil, 1970a). It seems unlikely that these organelles represent a form of rough-surfaced glyoxysomes, because they do not stain cytochemically for catalase, and they are only formed in cells in which glyoxysomes are maximally abundant (Vigil, 1970a). Their presence in preparations of isolated castor bean glyoxysomes could, however, account for the high levels of RNA r~ported for such preparations (Gerhardt and Beevers, 1969).

In terms of the segregation of microbody matrix proteins, it seems unlikely at present that they are directly inserted into the organelle during synthesis on ribosomes bound to the cytoplasmic surface of the organelle membrane.

5.4.3. Synthesis on Free Polysomes

The synthesis of microbody matrix proteins on ribosomes bound to a membrane would offer the advantage that the necessary transmembrane translocational step could occur cotranslationally. It is possible, however, that this translocational step might occur posttranslationally. In this case, the protein to be transported across the membrane could be synthesized on free polysomes,

196 J. Michael Lord

released into the cytoplasm and subsequently cross the membrane. In this way enzymic proteins could be added directly to their respective organelles. Recent evidence has shown that such a mechanism operates during the transport of cytoplasmically synthesized proteins into chloroplasts and mitochondria.

In the case of chloroplasts, these studies have concentrated on the transport of the small subunit of the photosynthetic carboxylating enzyme, ribulose diphosphate carboxylase (RuDPCase). Chloroplast RuDPCase is a largemolecular-mass multimeric protein containing eight large 55,000-dalton subunits and eight small 15,000-dalton subunits. The large subunit is encoded by chloroplast DNA and is synthesized within the organelles on 70 S chloroplast ribosomes, whereas the small subunit is encoded by nuclear DNA and synthesized on free 80 S ribosomes in the cytoplasm (Ellis, 1977). The assembly of the functional chloroplast enzyme therefore depends on the transport of the small subunit across both membranes of the chloroplast envelope.

When polyadenylated RNA isolated from pea, spinach, or Chlamydomonas reinhardtii is translated by a heterologous protein-synthesizing system, the small subunit of RuDPCase is synthesized as a precursor of higher molecular mass (20,000 daltons) (Dobberstein et aI., 1977; Cashmore et al., 1978; Chua and Schmidt, 1978; Highfield and Ellis, 1978). When the postribosomal supernatants from the in vitro protein synthesis mixtures are incubated with purified intact chloroplasts, the small subunit precursor is transported into the chloroplasts and processed to the mature size and charge. This in vitro transport process is physiologically significant, since the newly transported small subunits assemble with the endogenous large subunits to form the holoenzyme (Chua and Schmidt, 1978). Because the precursor's amino acid extension appears to be involved in a posttranslational transport mechanism, it has been designated as the transit peptide in order to distinguish it from the signal peptide of the cotranslationally transported precursors of secretory proteins (Chua and Schmidt, 1979). The intraorganellar site of the chloroplast processing enzyme was initially suggested to be the envelope membrane (Dobberstein et al., 1977), but recent evidence suggests that precursor-small subunit processing and holoenzyme assembly are stromal events (Smith and Ellis, 1979). Evidence has been presented that Chlamydomonas small subunit precursor is synthesized on free ribosomes from which it is presumably released into the cytoplasm after chain completion. Interaction with chloroplast membrane receptors may possibly be the first step in the transport process.

The three largest subunits (a, (3, and 'Y) of the F]-ATPase of yeast mitochondria have also been shown to be posttranslationally transported into mitochondria. Yeast F]-ATPase consists of five nonidentical subunits encoded by nuclear DNA, synthesized on 80 S cytoplasmic ribosomes, and then transported across the mitochondrial membranes to the matrix side of the inner membrane. When yeast polyadenylated RNA is translated in a cell-free system each of the three largest F] subunits is made as a larger precursor. When these precursors are incubated with purified intact yeast mitochondria under condi-


tions excluding protein synthesis, they are converted to the mature polypeptides and are simultaneously transported into the mitochondria by an energy-requiring mechanism (Maccecchini et ai., 1979). A similar posttranslational transport into mitochondria has been demonstrated for the cytoplasmically synthesized subunits of the cytochrome bel complex (Schatz, 1979). These data have induced Schatz (1979) to question the role of mitochondrial 80 S ribosomes (see Section 5.4.2) and the vectorial translation model for the insertion of mitochondrial proteins.

The transport of certain toxins, including the diphtheria toxin, and the toxic plant lectins abrin and ricin, is a further example of transport of proteins across a membrane. These toxins consist of two distinct polypeptide chains held together by disulfide bonds. One of these polypeptide chains, the A chain, is the biologically active toxic polypeptide that must cross the plasma membrane in order to exert its toxic effects. The other polypeptide, the B chain, interacts with receptors present in the plasma membrane and facilitates the translocation of the A chain. During this transport process, the B chain, or a large portion of it, remains embedded in the plasma membrane. Details of these transport processes with respect to the diphtheria toxin can be found elsewhere (Boquet et ai., 1976; Pappenheiner, 1977).

5.5. Implications for Microbody Matrix Protein Segregation

In the light of the experimentally established models discussed above, it is conceivable that microbody matrix proteins could be segregated by a membrane, either cotranslationally or posttranslationally. Depending on their mode of segregation, these proteins could be synthesized on bound ribosomes, free ribosomes, or on both classes of ribosomes. Synthesis on bound ribosomes would imply that the proteins were initially segregated by the ER membrane and temporarily housed in the cisternal space before reaching the microbody matrix after a vesiculation step. Synthesis on free ribosomes may result in the release of completed polypeptides, perhaps in the form of higher-molecularmass precursors, into the cytoplasm from which they might be transported into preexisting or nascent microbodies. The determination of the precise mode of microbody matrix protein segregation must await further experimental evidence identifying the site of synthesis and membrane translocation.

6. THE SYNTHESIS OF MICROBODY COMPONENTS

6.1. Membrane Lipids

The phospholipid composition of the membrane fraction recovered following the osmotic disruption of rat liver peroxisomes, spinach leaf peroxisomes, and castor bean glyoxysomes has been compared to that of the microsomal

198 J. Michael Lord

membranes isolated from the respective tissues. In each case, the microbody and microsomal membranes were similar in their phospholipid compositionPhosphatidylcholine, phosphatidylethanolamine, and phosphatidyl-inositol were the major components and accounted for approximately 50%, 30%, and 10% of the total lipid phosphorus, respectively (Donaldson et al .. 1972). The similarity of the glyoxysomal and ER membranes was confirmed in a more detailed analysis of these membranes isolated from castor bean endosperm. The phospholipid, free fatty acid and sterol composition was the same in both types of membrane, as was the fatty acid composition of the individual phospholipid classes (Donaldson and Beevers, 1977). This indicates that there is a single site of origin for these lipids.

The intracellular site of synthesis of the major peroxisomal membrane phospholipids in rat liver cells is the ER (Wirtz, 1974). Likewise the major glyoxysomal phospholipids in castor bean endosperm are synthesized by enzymes exclusively located in the ER membrane (Lord et al .. 1973; Moore et al .. 1973; Bowden and Lord, 1975).

Both rat liver and castor bean endosperm tissue contain phospholipid exchange proteins that could account for the transfer of phospholipid molecules from the ER to the microbody membrane. The exchange activity is low in castor bean endosperm cells, and the labeled phosphatidylcholine formed when the total particulate fraction from such cells was incubated with CDP[14C]choline was subsequently largely confined to the ER membranes (Lord, 1976). It seems likely that glyoxysomal membrane phospholipids are derived by membrane flow from the proliferating ER. In keeping with this, the labeling kinetics of various cellular membranes when intact castor bean endosperm is incubated with radioactive choline (Kagawa et al .. 1973), acetate (Donaldson, 1976) or CDP-choline (Lord, 1978) have established that the ER membrane becomes labeled before the glyoxysomal or mitochondrial membranes. Developmental studies with castor bean endosperm have indicated that a proliferation of the ER precedes the formation of glyoxysomes during the early stages of germination (Lord, 1978).

6.2. Membrane Proteins

Microbody membrane proteins have not been well characterized, particularly with respect to any enzymic activities they may possess. Certain enzymic proteins have been located in the microbody membrane, however, and they include NADH cytochrome c reductase, which has been found in rat liver and leaf peroxisomes and castor bean glyoxysomes (Donaldson, et al .. 1972). Castor bean glyoxysomal membranes also contain an alkaline lipase (Muto and Beevers, 1974), cytochrome hs (Donaldson and Beevers, 1977), and cinnamic acid 4-hydroxylase and p-chloro-N-methylaniline-N-demethylase (Young and Beevers, 1976). All these enzymic activities are also present in the ER mem-


brane. In addition, certain of the gluconeogenic pathway enzymes present in castor bean glyoxysomes appear to be peripheral proteins associated with the matrix surface of the glyoxysomal membrane. When the glyoxysomal fraction isolated by sucrose density gradient centrifugation is diluted, the organelles are osmotically disrupted, and certain matrix enzymes, e.g., catalase and isocitrate lyase, are almost completely solubilized. In contrast, enzymes such as malate dehydrogenase and citrate synthase remain associated with the glyoxysomal membrane to a significant extent, whereas malate synthase is almost completely bound to the membrane (Beiglmayer et al., 1973; Huang and Beevers, 1973). These enzymes associated with the glyoxysomal membrane are not adsorption artifacts generated during organelle subfractionation, nor are they integral membrane proteins, as they can be readily removed from the membrane by washing with 0.15 M KC (Huang and Beevers, 1973).

The polypeptide components present in washed glyoxysomal membranes have been separated by SDS-polyacrylamide gel electrophoresis (Brown et al., 1974).

The polypeptide profile of washed glyoxysomal membrane is similar to that of washed microsomal membranes (Bowden and Lord, 1976a). Isoelectric focusing showed that glyoxysomal and microsomal membrane proteins also share similar profiles on the basis of charge (Brown et al., 1976). Antibodies raised against glyoxysomal membrane proteins from both watermelon cotyledons (Hock, 1974) and castor bean endosperm (Bowden and Lord, 1977) recognize antigenic determinants in the ER membrane. These criteria establish that polypeptide components of the glyoxysomal membrane are also present in the ER membrane. Evidence that these polypeptides are initially inserted into the ER membrane was obtained when intact castor bean endosperm tissue was incubated with [35S]methionine. The kinetics of protein labeling clearly indicated that microsomal membrane proteins became radioactively labeled before their glyoxysomal membrane counterparts (Bowden and Lord, 197 6b).

A further indication that the ER is responsible for the synthesis of the glyoxysomal membrane proteins has been provided by the demonstration that many of the constituent proteins of these membranes are glycosylated. The sugars present in KCP-washed organelle membranes isolated from castor bean endosperm have been identified by gas liquid chromatography of their trimethylsilyl derivatives and are listed in Table I.

Studies with mammalian tissues have established that the core sugars, mannose and N-acetyl-glucosamine, are initially transferred to a carrier lipid, usually dolichol monophosphate (Waechter and Lennarz, 1976). These monosaccharide lipids are assembled into an oligosaccharide lipid. The core glycosylation step involves the en bloc transfer of the oligosaccharide moiety from the oligosaccharide lipid to the nascent polypeptides. This cotranslational step is catalysed by enzymes present in the ER membranes (Katz, et al., 1977). Likewise, in castor bean endosperm the ER membrane catalyzes the synthesis

200 J. Michael Lord

Table I Carbohydrate Content of Washed Castor Bean Membranes'

Sugar ER Glyoxysome Mitochondria

Mannose 0.35 0.56 1.08 N-Acetylglucosamine 0.31 0.48 0.70 Galactose 0.60 1.34 1.32 Fucose 0.15 0 0 Arabinose Trace 0.65 1.64 Glucose 0.19 Trace 0.32 Xylulose 0 0 1.09 Mannosamine 1.00 0 0 Sialic acid 0 0 0

'Expressed in micromoles sugar per milligram lipid-free protein.

of dolichol monophosphate mannose and the incorporation of this monosaccharide lipid into a man nose and N-acetyl-glucosamine-containing oligosaccharide lipid (Mellor and Lord, 1979a,b). Castor bean ER also catalyzes transfer of the man nose-containing oligosaccharide moiety from its lipid carrier to a chemically denatured acceptor protein, sulfitolysed ribonuclease A (Mellor et ai., 1979). This reaction was first demonstrated by Pless and Lennarz (1977) using hen oviduct membranes and is regarded as the in vitro equivalent of the cotranslational core glycosylation step (Kiely et ai., 1976).

Studies with secretory cells have emphasized the role of the Golgi apparatus during protein glycosylation. This cellular fraction contains glycosyltransferases that add terminal sugars such as galactose (Schacter, 1974). In the nondividing, nonsecreting castor bean endosperm cell the Golgi apparatus is not well developed, and galactose is incorporated into glycoproteins by an ER galactosyltransferase (Mellor and Lord, 1979c). The functional role of the castor bean ER in the glycosylation of glyoxysomal membrane proteins was confirmed by incubating intact tissue with either labeled galactose or GDPmannose-Iabeled glycoprotein appears in the ER membrane before the glyoxysomal membrane (Mellor and Lord, 1978, 1979a).

The ER has also been identified as the site of synthesis and segregation of the peripheral glyoxysomal membrane protein, malatc synthasc. Gonzalez and Beevers (1976) first demonstrated that the microsomal fraction isolated from germinating castor bean endosperm cells at a developmental stage when the rate of glyoxysome formation is actively increasing was found to contain a large proportion of the total cellular malate synthase. Microsomal and glyoxysomal malate synthases are physically and serologically identical and the microsomal enzyme is ultimately sequestered in glyoxysomes (Bowden and Lord, 1978;


Lord and Bowden, 1978). At no time can soluble malate synthase be detected during cellular fractionation. Glyoxysomal malate synthase is a glycoprotein and, predictably, the microsomal enzyme has already been glycosylated prior to its transfer to the glyoxysomes (Mellor et af., 1978). It seems likely that malate synthase is synthesized on bound ribosomes, glycosylated and fixed as a peripheral membrane protein on the luminal surface of the ER membrane. In support of this concept nascent malate synthase chains are preferentially associated with bound, rather than free, polysomes (Lord and Bowden, 1978).

Collectively, these studies indicate that proteins which span the microbody membrane are synthesized, glycosylated if appropriate, and given their asymmetric membrane orientation in the ER. The insertion of newly synthesized phospholipids into the ER membrane in the vicinity of these integral proteins permits this proliferating membrane to provide the limiting membrane of other cellular organelles, including peroxisomes and glyoxysomes. The microbody membrane is derived from the proliferating ER by a process of membrane flow. The direct continuity occasionally seen between the ER and microbody membranes may be a visualization of the budding process. After cotranslational insertion into the rough ER membrane, transmembrane proteins destined for the microbody membrane may be sorted from other integral membrane proteins on the basis of some structural information possessed only by microbody membrane proteins. If such a sorting step occurs, it would precede the vesiculation step and could account for the different catalytic properties of the ER and microbody membranes.

6.3. Matrix Proteins

The microbody matrix protein most extensively studied in terms of its site of synthesis and pathway of intracellular transport is peroxisomal catalase of rat liver. One of the earliest studies in this respect was that of Higashi and Peters (1963), who examined the intracellular distribution of labeled catalase in rat liver at various times after the animals had been injected with [14C]leucine. Apart from an unexplained early peak in the supernatant fraction, their results indicated that catalase antigen initially appeared in the rough microsomal fraction and was subsequently transferred to the peroxisomes. Higashi and co-workers have also reported the immunoprecipitation of nascent catalase chains from both free and bound polysomes, although they suggested that the membrane bound ribosomes were primarily responsible for peroxisomal catalase synthesis (Kashiwagi et al., 1971). The conclusion to be drawn from these studies, that peroxisomal catalase is synthesized and segregated by the rough ER, has not been substantiated in several subsequent investigations.

Redman et al. (1972) repeated the experiment of Higashi and Peters (1963) but used improved cellular fractionation procedures, and compared the

202 J. Michael Lord

intracellular pathway of newly formed liver catalase with that of a known secretory protein, albumin. Radioactive catalase and albumin both appeared in the liver fraction some 5-10 min after the rats were injected with [14C]leucine, but they appeared in different intracellular locations. Catalase first appeared in the soluble cytoplasmic fraction while albumin was found in the rough ER. Albumin was transported from the rough ER to the smooth ER in accordance with the predicted secretory protein pathway. Catalase, on the other hand, did not enter the rough or smooth ER and appeared to move directly from the cytoplasm into the peroxisomes. These data suggest that catalase either enters preformed peroxisomes or that it crosses the ER membrane in specialized regions from which peroxisomes are being generated by vesiculation.

Lazarow and de Duve (1973a,b) have confirmed that newly synthesized catalase is released into the cytoplasm rather than discharged across the ER membrane. In a detailed kinetic study these workers established that catalase antigen first appears in the form of a hemeless precursor of approximately the same molecular weight as the catalase monomer, but that does not accompany catalase through its chemical purification procedure. This catalase precursor was first recovered in the high speed supernatant fraction from where it was transferred into the peroxisomes where it acquired heme and then aggregated to form the active tetrameric catalase molecule.

Robbi and Lazarow (1978) have immunochemically isolated catalase synthesized when rat liver polyadenylated RNA was translated in either the rabbit reticulocyte lysate or the wheat germ cell-free protein-synthesizing systems. Catalase antigen immunoprecipitated from the translational systems and completed catalase immunoprecipitated from peroxisomes had identical monomer molecular masses (66,000) on the basis of their relative mobilities in SDS gels. This molecular mass was some 4000 daltons larger than than determined for purified peroxisomal catalase. Robbi and Lazarow (1978) concluded that catalase is not synthesized as a higher-molecular-mass precursor in the cell-free systems, because its apparent molecular mass did not change when it entered the peroxisomes. The smaller size of the purified catalase was attributed to a decrease that occurred during the chemical purification procedure.

Goldman and Blobel (1978) have likewise shown that the molecular weights of both catalase and uricase isolated from a cell-free system translating rat liver mRNA were identical to those of the mature peroxisomal enzymes. These workers further established that both catalase and uricase are exclusively synthesized on free, rather than on bound, polysomes. The fidelity of the polysome separation technique was confirmed by examining the synthesis of albumin. This product was exclusively synthesized by mRNA located in the bound polysome fraction. Moreover, newly synthesized albumin, in contrast to both catalase and uricase, was cotranslationally segregated when dog pancrease microsomal vesicles were added to the translational system.


At the present time, therefore, it appears likely that rat liver catalase is synthesized on free ribosomes and is released into the cytoplasm before its posttranslational transfer into preexisting or forming peroxisomes. It is also possible that catalase and uricase contain uncleaved signal sequences and thus are not synthesized as larger precursors. These conclusions cannot be applied to glyoxysomal malate dehydrogenase at present, because Walk and Hock (1978) have demonstrated that the translation of watermelon cotyledon mRNA in the wheat germ system results in the synthesis of a malate dehydrogenase precursor 5000 daltons larger than the mature enzymic polypeptide. The proteolytic processing and membrane segregation of glyoxysomal malate dehydrogenase have not yet been demonstrated. The malate dehydrogenase results do suggest, however, that it is still too early to generalize about a mechanism for the synthesis and segregation of microbody matrix proteins. The elucidation of the detailed mechanism for microbody biogenesis must await further experimental evidence which, in the light of techniques currently available, should not be long forthcoming.

ACKNOWLEDG MENTS

I am particularly grateful to Dr. P. B. Lazarow, Dr. E. H. Newcomb, and Dr. E. L. Vigil for kindly providing me with electron micrographs.

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Blobel, G., and Dobberstein, 8., 1975. Transfer of proteins across membranes. 1. Presence of proteolytically processed and unprocessed nascent immunoglobulin light chains on membrane-bound ribosomes of murine myeloma, J. Cell Bioi. 67:835-851.

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Bowden, L., and Lord, J. M., 1976a, Similarities in the polypeptide composition of glyoxysomal and endoplasmic reticulum membranes from castor bean endosperm, Biochem. J. 154:491-499.

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Bowden, L., and Lord, J. M., 1977, Serological and developmental relationships between endoplasmic reticulum and glyoxysomal proteins of castor bean endosperm, Planta 134:267-272.

Bowden, L., and Lord, J. M., 1978, Purification and comparative properties of microsomal and glyoxysomal malate synthase from castor bean endosperm, Plant Physiol. 61:259-265.

Bowes, G., Ogren, W. L., and Hageman, R. H., 1971, Phosphoglycolate production catalysed by ribulose diphosphate carboxylase, Biochem. Biophys. Res. Commun. 45:716-722.

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Vigil, E. L., 1970a, Cytochemical and developmental changes in microbodies (glyoxysomes) and related organelles of castor bean endosperm, J. Cell Bioi. 46:435-454.

Vigil, E. L., 1970b, Intracellular localization of catalase (peroxidatic) activity in plant microbodies, J. Histochem. Cytochem. 17:425-428.

Vigil, E. L., 1973, Structure and function of plant microbodies, Subcell. Biochem. 2:237-285. Waechter, C. J., and Lennarz, W. J., 1976, The role of polyphenol-linked sugars in glycoprotein

synthesis, Annu. Rev. Biochem. 45:95-112. Walk, R. A., and Hock, 8., 1978, Cell-free synthesis of glyoxysomal malate dehydrogenase,

Biochem. Biophys. Res. Commun. 81:636-643. Wilgram, G. F., and Kennedy, E. P., 1963, Intracellular distribution of some enzymes catalysing

reactions in the biosynthesis of complex lipids, J. Bioi. Chem. 238:2615-2619. Wirtz, K. W. A., 1974, Transfer of phospholipids between membranes, Biochim. Biophys. Acta

344:95-117. Wirtz, K. W. A., and Zilversmit, D. B., 1968, Exchange of phospholipids between liver mito

chondria and microsomes 'in vitro,' J. Bioi. Chem. 243:3596-3602. Wirtz, K. W. A., and Zilversmit, D. 8., 1969, The use of phenobarbital and carbon tetrachloride

to examine liver phospholipid exchange in intact rats, Biochim. Biophys. Acta 187:468-476. Yamazaki, R. K., and Tolbert, N. E., 1969, Malate dehydrogenase in leaf peroxisomes, Biochim.

Biophys. Acta 178:11-20. Young, 0., and Beevers, H., 1976, Mixed function oxidases from germinating castor bean endo

sperm, Phytochemistry 15:378-385. Yu, Y. L., Tolbert, N. E., and Orth, G. M., 1964, Isolation and distribution of phosphoglycolate

phosphatase, Plant Physiol. 39:643-647.

Chapter 5

The Subcellular Biochemistry of Thyroid

H. J. Hilderson, G. Van Dessel, A. Lagrou, and W. Dierick RUCA-Laboratory for Human Biochemistry and UIA-Laboratory for Pathological

Biochemistry University of Antwerp Groenenborgerlaan 17 I, B2020 Antwerp, Belgium

1. INTRODUCTION

The thyroid gland elaborates, stores, and releases into the bloodstream hormones (iodinated derivatives of tyrosine), that, in the adult animal, are chiefly concerned with the regulation of energy production. It differs from most endocrine glands by using extracellular storage of its hormonal products (in most endocrine glands there are only limited provisions for intracellular storage and there is no extracellular storage system). A high percentage (up to 95%) of the iodide captured is stored as iodothyronyl and iodotyrosyl groups incorporated in the polypeptide chains of thyroglobulin (the normal content of adult thyroid is 10 mg iodine, i.e., equivalent to the need for 6 months). Microscopically the gland consists of a series of follicles (spherical cystlike vesicles), that are lined by a single layer of epithelial cells (follicular epithelium). The lumen (inner space) of the follicle is filled with colloid, composed mainly of thyroglobulin and some mucinous material. The plasma membrane of these cells have very specialized regions. The stretch of plasma membrane facing the stroma, which is rich in blood vessels, lymphatics, and connective tissue, is designated as the basal or antiluminal membrane. That part of the plasma membrane that is oriented toward the lumen is called the apical or luminal border of the follicular cell. It shows numerous microvilli that are small in resting cells, but often long and branched in stimulated ones. Shortly after stimulation, the apical portion of the follicular cell develops cytoplasmic processes that fuse around portions of the follicular colloid and thereby lead to their endocytotic uptake into the cell, where they form colloid droplets that can fuse with lysosomes. The

213

214 H. J. Hilderson et al.

cytoplasma contains microtubules and microfilaments subjacent to the apical plasmalemma, which seem important to endocytosis in a manner that is not completely understood. In addition to the normal subcellular components (nucleus, mitochondria, lysosomes, well-developed Golgi apparatus, prominent endoplasmic reticulum), thyroid cells also contain two types of secretory granules, called A and B granules. The A granules are peroxidase-positive and appear to arise mainly from the endoplasmic reticulum in the apical portions of the cells. The B granules are peroxidase-negative, and they apparently form from Golgi-endoplasmic reticulum-lysosome (GERL) (Novikoff et ai., 1974).

The most crucial biochemical events in the thyroid are the uptake of extracellular iodide (the normal daily diet has no more than 200 /lg; its concentration in blood is less than 10-7 M) and its transport to the interior of the cell (Taurog, 1978) where it is oxidized and coupled to the thyroglobulin (Figure 1). Thyroglobulin is stored in the follicle lumen. After resorption from the colloid by pinocytosis, the thyroglobulin is hydrolysed intracellularly (by lysosomes) with formation of free iodothyronines (T3 = 3',3,5-triiodothyronine, T4 = T x = 3',5',3,5-tetraiodothyronine). Nonhormonally active iodotyrosines are also formed in this process (MIT = 3-monoiodotyrosine, DIT = 3,5-diiodotyrosine). The iodothyronines are secreted, while the iodotyrosines are deiodinated and the iodide reutilized. Every step in thyroid metabolism, leading to synthesis and secretion of thyroid hormones, seems to be stimulated by thyroidstimulating hormone or thyrotropin (TSH). As TSH has many effects on the structure as well as on the function of the thyroid gland, the study of the subcellular biochemistry of the thyroid gland is of major importance.

Subcellular biochemistry intends to provide information not only about the chemical nature and physical properties of membranes, but also on the chemical constituents and enzyme activities localized in the various cellular compartments. Therefore it is essential that isolated organelles and/or membrane fragments retain their structural organization and their original metabolic activities. The characterization of isolated fractions may include microscopy and the measurement of the concentration relative to the homogenate of a variety of markers. For most tissues, markers and fractionation schemes are derived from studies on rat liver. However, enzymes that are appropriate markers for liver may not occur in some other tissues. Also, marker enzymes for liver may, in other organs, have a different physiological function and localization. In thyroid glands the interpretation of experimental data is complicated further by the drastic procedures required for homogenization, resulting in damage to some subcellular organelles and disruption of membranes into small vesicles.

This review on thyroid tissue deals with homogenizing procedures, cell fractionation, and isolation of enriched subcellular fractions. Furthermore,

Subcellular Biochemistry of Thyroid 215

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attention is focused on the methods used and the validity of markers. This review does not deal with the overall metabolism of the thyroid cells. No attempt has been made to review the possible biochemical and clinical applications of studies on the subcellular biochemistry of thyroid.

2. CELL FRACTIONATION

2.1. Disruption and Homogenization of Thyroid Tissue

Careful anatomical preparation is essential in order to facilitate the homogenization procedure. Adhering adipose and connective tissue must be removed. For maximal preservation of intact subcellular organelles, thyroid tissue must be homogenized as gently as possible. The homogenization technique to be used is also dependent on the subcellular fractions one wishes to isolate. All manipulations must be performed at 4°C in order to prevent autolysis.

In order to obtain the distribution patterns for a series of markers, the minced tissue is generally homogenized in 0.25 M sucrose with an all glass loose fitting Ten Broeck homogenizer and filtered through a double layer of cheesecloth. The filtrate is taken as 100%. It still contains whole cells, blood cells, and connective tissue. Further homogenization in a Potter-Elvehjem homogenizer is possible without supplementary damage to cell components (Dierick and Hilderson, 1967).


To isolate nuclei, the tissue is minced, washed repeatedly, and homogenized in a Ten Broeck homogenizer (0.32 M sucrose, 3 mM with respect to MgCI 2, adjusted to pH 7.4 with NaHC03) (Hilderson and Dierick, 1974). To isolate nuclei on a larger scale, thyroid tissue is homogenized in a Virtis homogenizer at low speed (Voets et al., 1978). When high recoveries of RNA-polymerase activities in the nuclear fraction are required hypertonic sucrose solution is preferred as homogenization medium (Voets et al., 1979a).

To analyze the distribution of thyroid membranes, a two-step procedure is applied: (1) minced tissue is treated in a Virtis homogenizer (100 g in 300 ml isotonic sucrose medium, pH 7.4 at 11,000 rpm for 2 X 1 min) or in a Waring Blendor (medium speed), after which (2) the resulting suspension is homogenized further in a Potter-Elvehjem homogenizer (Teflon pestle; 3000 rpm; five strokes) (Hilderson et al., 1975).

For the preparation of plasma membrane-enriched fractions, homogenization is the critical step (for details see Section 4.5). The final concentration of the tissue in the homogenization medium (2-3 gin 20 ml)also seems to be critical (Wolff and Jones, 1971).

2.2. Localization of Marker Enzymes in Thyroid (Preliminary Studies)

The study of markers is based on the premise that each morphologically distinguishable cell component is characterized by some constituent or combination of features that render that particular cell component unique. Markers are chemical, enzymic, or antigenic constituents that have previously been shown to be localized exclusively, or at least concentrated at known cellular sites. They can therefore be used to define the distribution of material from these sites in the isolated fractions.

For thyroid tissue the subcellular localization of {1-glucuronidase was investigated first (Hilderson et aI., 1965; Dierick and Hilderson, 1967). The enzyme is sedimentable from a homogenate and displays latent enzyme activity (45-55% for a whole homogenate). It can be released by repeated freezing and thawing, preincubation at pH 5, osmotic shock, and exposure to detergents. Moreover, a lag period (time elapsing before free enzyme activity increases) is apparent in the velocity curve at 37"C ranging from 45 to 50 min. Similar Km values for both free enzyme activity and total enzyme activity are found, pointing to the presence of an impermeable membrane between {1-glucuronidase and the substrate.

Indeed, if an enzyme is located inside a particle with a permeable membrane the substrate itself will reach the selected enzyme molecules by pene-


trating the particles during the incubation period. As a result, the latent enzyme activity will decrease with increasing substrate concentration and different values for the substrate concentration will be obtained at Vj2 (V= maximal velocity) for total and free enzyme activity.

For cytochrome oxidase, latent enzyme activity (80-90%) is found with whole homogenates. No soluble enzyme activity is detectable. The curve of release of cytochrome oxidase, occurring with prolonged preincubation, differs from that of ,3-glucuronidase. These observations are best explained by assuming that in thyroid homogenate these two enzymes are released from different particles. These results were confirmed by Herveg et at. (1966). In liver, ,3-glucuronidase is known to be a lysosomal enzyme, whereas cytochrome oxidase displays a mitochondrial localization. Summarizing, one can say that the same localization occurs in thyroid tissue.

Similar studies were performed on other lysosomal hydrolases (,3-glycerophosphatase, di-Na-phenylphosphatase, phosphodiesterase, ribonuclease, deoxyribonuclease) and on the soluble enzyme adenosine deaminase (Hilderson et at., 1970). The release of the different acid hydrolases in the whole homogenate during the incubation period does not proceed in the same way for all enzymes. This could be a consequence of the presence of different substrates in the incubation mixtures. The release of latent enzyme activity as a function of ageing of a whole homogenate at 4°C, repeated freezing and thawing and preincubation at 3rC is different for all hydrolases investigated. This can be explained by (1) the existence of different types of lysosomes in the same cell, (2) the existence of differing thyroid cell types (or cells in different metabolic states), (3) leaks in the membranes through which the enzymes can leave the particles or be reached by the substrates in different ways, and (4) an internal structural organization of the lysosomes resulting in differential release of hydrolases in the different experimental conditions (de Duve, 1963). Furthermore, part of the released enzyme activity is not recovered in the soluble fraction but remains sedimentable ("sedimentable free enzyme activity") (Hilderson et at., 1971a). The significance of this phenomenon was discussed by Shibko and Tappel (1965)-first the substrates penetrate into the particle through gaps in the membrane; the enzymes, however, cannot yet leave the damaged particle (free enzyme activity increases, sedimentable free enzyme activity increases, and soluble enzyme activity remains unchanged). When the destruction of the membrane becomes more extensive, the enzymes are able to leave the particles and soluble enzyme activity increases. During the preincubation period, free enzyme activity never equals total enzyme activity; part of the enzyme activity remains latent. This is in agreement with the results of Allen and Gockerman (1964) for liver cells.


Two substrates (,B-glycerophosphate, di-Na-phenylphosphate) used for studying phosphatase activities yield differing results in a number of experiments: (1) differences in latent enzyme activity values for whole homogenate, (2) differences in the conversion of latent enzyme activity to free enzyme activity, and (3) differences in distribution patterns during differential pelleting. Furthermore, gel filtration of aIM NaCi thyroid tissue homogenate shows different elution profiles for both enzymic activities (Hilderson et aI., 1971 b). Therefore, at least two acid phosphatases (phenylphosphatase and ,B-glycerophosphatase) are present in thyroid tissue.

From these introductory studies, the following enzymes can be choosen as markers in order to initiate studies on differential pelleting and gradient centrifugation-cytochrome oxidase (mitochondrial marker), ,B-glycerophosphatase, di-Na-phenylphosphatase, phosphodiesterase, and ribonuclease (lysosomal markers).

2.3. Differential Pelleting

A group of authors (De Groot and Carvalho, 1960; Kogi and Van Deenen, 1961; Satyaswaroop, 1971) prepared four fractions starting with a 0.25 M sucrose homogenate using essentially the original scheme devised for rat liver homogenization (Schneider and Hogeboom, 1950). Balasubramaniam et al. (1965) also presented a four-fraction scheme for dog thyroid, using different centrifugation conditions (800g X 15 min; 15,000g X 20 min; 100,000g X 90 min, supernatant). Chabaud et al. (1971) presented a six-fraction scheme for sheep thyroid (1000g, 3300g, 1O,000g, and 25,000g X 10 min and 145,000g X 60 min) designated as PIO, P33, PlOD' P25Q, P1450, and supernatant. The distribution patterns of different glycosidases were interpreted as being related to those of acid phosphatases and used as an indication of a lysosomal localization. However, a substantial part of the enzyme activities were recovered in the final supernatant and the distribution patterns themselves did not completely coincide with each other and with acid phosphatase.

In our approach for devising a different pelleting scheme, ,B-glucuronidase and cytochrome oxidase were used as markers to look for the best resolution between Iysosomes and mitochondria (Dierick and Hilderson, 1967). Whole homogenate was centrifuged for 10 min at 700g. The supernatant was collected and the sediment of the second washing was called the "nuclear fraction" (N). The three supernatants were then pooled and spun down for 15 min at 25,000g followed by washing the sediment twice under identical conditions. The sediment of the last washing was designated as the "mitochondrial fraction" (M). At this stage the three supernatants were again pooled and centrifuged for 15 min at 73,000g. The resulting sediment was suspended in 0.25 M sucrose (L,


light mitochondrial fraction). Finally, the supernatant of this run was centrifuged again (60 min at 104,000g), yielding a microsomal fraction (P) and a supernatant (S fraction). Adsorption did not interfere with the distribution patterns for both iJ-glucuronidase and cytochrome oxidase. The maximum specific activities for both enzymes were found in the M fraction (Figure 2). No cytochrome oxidase was found in the S fraction, while iJ-glucuronidase was frequently present in large amounts. Other possible markers for lysosomes and mitochondria (except for succinate dehydrogenase that is also present in appreciable amounts in the S fraction) resulted in essentially the same distribution patterns. From these results, one can conclude that this differential pelleting scheme fails to separate lysosomes from mitochondria. This is a consequence of the higher g values (25,000) required to pellet mitochondria. This phenomenon was also observed by other authors (Jablonski and McQuillan, 1967; Fisher et aI., 1968). As thyroid tissue is tough, homogenization always results in appreciable damage to subcellular organelles. The membranes are disrupted, and a large population of vesicles (diameter smaller than 0.2 p,m; Hilderson, unpublished results) appears. The bulk of these vesicular structures, probably derived from endoplasmic reticulum and plasma membranes, are recovered in the M and L fractions. Finally, more than 80% of the proteins is found in the S fraction. This is because of the typical architecture of thyroid follicles containing large amounts of thyroglobulin, stored extracellularly in the follicle lumen.

For alkaline phosphatase as well as for ATPase and 5' -nucleotidase, differential pelleting results in distribution patterns comparable with each other and showing the highest relative specific activity in the L fraction (Lagrou et al., 1974a; De Wolf et aI., 1978b).* This could imply a joint localization in plasma membranes. This is in accordance with the results of Yamashita and Field (1970) and Wolff and Jones (1971) who, using histochemical techniques, locate 51-nucleotidase and ATPase in thyroid plasma membranes. The distribution patterns of lipid bound sialic acid (Lagrou et al., 1974a), cholesterol (Hilderson et al., 1975), and adenyl ate cyclase (Hilderson et al., unpublished results) are similar to the patterns recorded for plasma membrane markers. Therefore, the bulk of these constituents and enzymes is probably localized in plasma membranes. However, a large fraction of the plasma membrane markers (also endoplasmic reticular membrane markers; see Section 2.5) are recovered in the S fraction. This fraction can be subfractionated by centrifuging at 104,000g for 16 hr yielding a sediment SO' a viscous red fraction Sb, a yellow fraction Sc layered over the sediment Sa, and a colorless supernatant Sd

*Mutsazaki et al. (1973) omitted the isolation of the L fraction and found the highest specific activity in their P fraction.

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(Van Dessel et al., 1977). When the Sb fraction is applied onto the Sepharose 2B column and eluted with 0.14 M NaCI pH 7.4 several markers are eluted in two peaks. The first peak is eluted with the void volume, the second one is retarded. The bulk of the lipids (85% of the cholesterol, 70% of the total phospholipids, and 70% of the sphingomyelin), some proteins (3% of the total), and most of the plasma membrane markers as well as of the endoplasmic reticular membrane markers are recovered in the first peak. Therefore, this peak represents membranous material (De Wolf et al., 1978b).

The unique pattern (almost equally distributed throughout the different fractions) of ~-glucosidase activity results from the presence of two different enzymes-a soluble ~-glucosidase with alkaline pH optimum and an acid ~glucosidase localized in lysosomal membranes (Hilderson et al., 1975; see also Section 2.4).

Lactate dehydrogenase is predominantly recovered in the S fraction. Therefore in thyroid as in other cells it is probably localized in the cytosol.

From the distribution patterns, no clear-cut conclusion can be drawn regarding the preferential localization of individual phospholipids in the different types of subcellular membranes (G. Van Dessel, A. Lagro!!, H. J. Hilderson, and M. De Wolf, unpublished results).

Chabaud et af. (1974) attempted to isolate a Golgi-rich microsomal preparation. Alkaline phosphatase, 5'-nucleotidase, and acid phenyl phosphatase were all enriched mostly in their C fraction in which there was no clear differentiation between lysosomes and plasma membranes. This is not surprising, as this fraction corresponds to a part of our M fraction. Therefore, substantial overlap in distribution of different cell components must occur. Moreover, they report that 20-30% of total glycosyltransferase activities are recovered in the post microsomal supernatants. Spiro and Spiro (1973) performed similar experiments. Mannosyltransferase was found entirely bound to the particulate fraction while substantial amounts of sialyltransferase and galactosyltransferase were recovered in the high speed supernatant. In light of our experiments, these results do not exclude membrane localization of those transferases. Using our approach, we were indeed able to recover galactosyltransferase activity in the Sb fraction (Hilderson et al., unpublished results).

2.4. Gradient Centrifugation Studies

Gradient centrifugation studies have been performed in order to determine sedimentation constants for cell organelles and to get more precise information about subcellular localization of some enzymes and constituents. Gradient centrifugation can also be applied for preparative purposes.


From zonal differential gradient centrifugations in 0.35-0.50 M sucrose gradients s values for bovine thyroid mitochondria ranging from 0.23 X 10-9

to 0.56 X 10-9 sec are obtained (Hilderson et al., 1971c). These values are in the same range as those for rat liver lysosomes. Therefore, higher centrifugation forces are necessary for sedimenting thyroid mitochondria than for rat liver mitochondria. It is impossible to calculate a sedimentation constant for bovine thyroid lysosomes, as no peak can be detected for acid hydrolases.

After 2 or 3 hr equilibration at 2"C of an M + L fraction in a 1.0-2.0 M sucrose density gradient using an SW-rotor at 73,300g cytochrome oxidase bands as a symmetrical peak (1.18 ± 0.01 gjcm3). Acid hydrolases are distributed throughout the whole gradient displaying a smooth maximum at densities higher than the mitochondrial peak. /3-Glycerophosphate, a stabilizing agent for lysosomes (Hilderson et al., 1971 b), does not affect the position of the mitochondrial peak. However, a symmetrical lysosomal peak forms, completely overlapping the mitochondrial peak.

After buoyant-density equilibration of an M + L fraction in a B-XIV or HS-zonal rotor, two distinct zones can be observed in the sedimentation profiles (Figure 3)-zone I at lower densities, and zone II at higher densities (Hilderson et al., 1975). Cytochrome bs is only present in zone I. In rat liver cytochrome bs is largely confined to smooth endoplasmic reticulum (Hinton et al., 1970). This indicates that in our experiments smooth endoplasmic reticular material is only present in zone I. The rest of the gradient (zone II) is probably free of smooth endoplasmic reticulum. 5'-N ucleotidase, ATPase, and alkaline phosphatase profiles are confined to zone I, running parallel to each other. Therefore, zone I also contains plasma membranes. Furthermore, the relative specific concentration of sphingomyelin is higher in zone I than in zone II. This also points to a localization of plasma membranes in zone I. This result is also compatible with the statement made by Morn~ et al. (1974) that increasing amounts of sphingomyelin are present in the sequence-smooth endoplasmic reticulum < Golgi apparatus < secretion vesicles < plasma membranes (membrane flow differentiation).*

Zone II is the only part of the gradient where RNA is found. Therefore, rough endoplasmic reticulum must be localized in zone II. In this zone the

*During the life of a cell, membrane material moves from one subcellular component to another in the following sequence: rough endoplasmic reticulum, smooth endoplasmic reticulum, Golgi apparatus, secretion granules or vesicles (exocytosis), and plasma membranes. Moreover, on endocytosis, membrane material moves from plasma membranes to vacuoles and secondary lysosomes. During these processes, membrane differentiation occurs and the composition and organization of membranes is changed by different means, such as addition of new enzymes and constituents, removal or destruction or inactivation of enzymes previously present. These combined processes are collectively designated as membrane flow differentiation.

Subcellular Biochemistry of Thyroid

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cytochrome oxidase profile shows a peak with density 1.19 g/cm3• This peak is very probably due to the presence of intact mitochondria. Indeed, mitochondria obtained by a mild homogenization procedure (collagenase treatment of thyroid tissue) also equilibrate at a similar density (1.17-1.19 g/ cm3) (Hilderson and Dierick, 1973). The cytochrome oxidase profile also extends into zone I, indicating to what extent mitochondria are damaged during the experiment. After a more traumatizing homogenization procedure a double peak can be observed (De Wolf et al., 1978a). Part of the succinate dehydrogenase activity (5-10%) is recovered at the top of the gradient (Hilderson et al., unpublished results) (not shown in Figure 3). In the gradient the distribution profile coincides completely with cytochrome c oxidase and monoamine oxidase.

NADPH-cytochrome c reductase and glucose-6-phosphatase are found in both zones. As NADPH-cytochrome c reductase is located in rat liver endoplasmic reticulum (see Lee et al., 1969) and rat liver glucose-6-phosphatase is also located in the endoplasmic reticulum (see de Duve, 1971), it can be concluded from the NADPH-cytochrome c reductase and the glucose-6-phosphatase profiles (see Section 2.5) that these enzymes qualify as endoplasmic reticulum markers in thyroid. Furthermore, phospholipid profiles, reflecting the distribution of membranous material, generally follow the glucose-6-phosphatase profiles rather closely.

Acid phosphatase is found throughout the whole gradient. This is less pronounced for the other hydrolases that show a peak around density 1.19 g/cm3•

However, their profiles do not coincide. This may be because of heterogeneity of the lysosomal population or differences in release of enzymes from disrupted lysosomes, or both. f3-Glucosidase is recovered from several bands. A peak at the top of the gradient (soluble enzyme) has a pH optimum of 8.5-9.0 and a Km value of 2.3 mM. A band at density 1.19 g/cm3 shows a maximum activity at pH 4.0 and has a Km value of 1.15 mM. The activity in the gradient has a lysosomal localization. This can be concluded from the acid pH optimum and from the existence of latent enzyme activity with a lag period for f3-glucosidase in the M + L fraction. Also, in rat kidney two f3-glucosidases were identifiedone associated with lysosomal membranes and the other one, differing in pH optimum, in the supernatant. In the cytoplasma of rat liver only a Iysosomelinked f3-glucosidase has been found (Patel and Tappel, 1969a,b). Similar sedimentation profiles as for the M + L fraction are found for the P fraction. However, the f3-glucosidase band recovered at density 1.13 g/cm3 had increased appreciably, and the band at 1.19 g/ cm] diminished accordingly. This could be because of the presence at density 1.13 g/ cm] of damaged lysosomes and would imply that the lysosomal f3-glucosidase is membrane bound.

About 80% of the lactate dehydrogenase activity is recovered at the top of the gradient (Hilderson et al., unpublished results; not shown in Figure 3). The remaining 20% is distributed throughout the gradient. This could be


because of nonspecific adsorption or to inclusion of cytosol within small vesicles formed during the homogenization procedure (Schengrund and Rosenberg, 1970).

Cholesterol profiles run parallel to the 5'-nucleotidase and alkaline phosphatase profiles (plasma membrane markers) in zone I. In zone II, however, there is a tendency to follow the glucose-6-phosphatase profile (endoplasmic reticulum marker, see Section 2.5.). The molar ratio of cholesterol/phospholipids decreases from 0.37 to 0.19 as a function of increasing density. Although there is controversy about the presence of cholesterol in rat liver endoplasmic reticulum membranes (Thines-Sempoux, 1974; Glauman et aI., 1974), its presence is confirmed for bovine thyroid by (1) the cholesterol profiles in the experiments reported here, (2) the relatively high cholesterol/phospholipid molar ratio in zone II (0.19) compared with the low ratio (0.088) in purified bovine thyroid nuclei (Hilderson et al., 1974), and (3) the shift of the position of plasma and endoplasmic reticulum membranes after digitonin treatment. Indeed, when an M + L fraction is exposed to low concentration of digitonin and is then subjected to density equilibration, the distributions of most markers have changed. 5'-Nucleotidase shifts from 1.13 to 1.15 g/cm3 and alkaline phosphatase from 1.13 to 1.14 g/cm3• The profile for glucose-6-phosphatase has changed from a double-peak distribution (densities 1.13 and 1.20 g/cm3)

to a continuous distribution with broad maximum at density 1.16 g/cm3• The cholesterol profile shows a shift of a maximum from 1.13 g/cm3 to 1.15-1.16 g/cm3, whereas the maximum at 1.20 g/cm3 has vanished.

Catalase is chiefly recovered after differential pelleting in the Sc and Sd fractions (De Wolf et al., 1978b; Figure 2). This may reflect the extent to which the peroxisomes have been damaged during the homogenizing process. After isopycnic gradient centrifugation in a zonal rotor, the sedimentable fraction equilibrates as a sharp peak around density 1.19 g/cm3, suggesting that the localization is in one single organelle. Tailing of the band suggests that during the movement through the gradient enzyme is continuously released from the cell organelle. Urate oxidase, detectable in whole homogenates and M + L fractions, is present in concentrations too low for quantitative measurements.

Ribosomes are released from the rough endoplasmic reticulum membranes when heparin is incorporated in both gradient and homogenization medium. However, heparin seems to have a damaging effect on bovine thyroid membranes. Similar results are obtained with the chaotropic agent pyrophosphate. This not only releases ribosomes but also removes the outer membrane from the mitochondria (Figure 4; De Wolf et al., 1978b).

After lO-hr centrifugation of M + L fractions in an HS-zonal rotor, most markers approach their isopycnic zones. Satisfactory resolution is obtained between subcellular components. After 24-hr centrifugation good resolution is

12 ~ a d

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4 0

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10

8

6

4

2

0 I I I I I I I I I 0 10 20 30 40 15 10 5 0

FRACTION NUMBER

FIGURE 4. Biphasic centrifugation of a pyrophosphate-treated M + L fraction in a HS zonal rotor. Experimental conditions: An M + L fraction was treated overnight at 4 0 C with 50 mM sodium pyrophosphate (pH 7.4 in 5 mM Tris/HCl buffer). A 12-ml sample was injected. After an initial phase (9000 rpm for 5 hr), the denser part of the gradient was replaced (d-f), and the centrifugation was continued for an additional 19 hr at 9000 rpm (a-c). (a,d) (- - - -) Protein; (_._.) A2Ml; (-) A2BO; (-0-0) slope of gradient. (b,e) ( .... ) 5'-Nucleotidase; (-) glucose-6-phosphatase; (- - - -) acid phosphatase; (--0--0) cytochrome c oxidase; (- 0-0) monoamine oxidase. (c,f) (_. - .) Guaiacolperoxidase; (--) p-phenylenediamine peroxidase; (- 0 - 0 ) 3,3' -diaminobenzidine tetrachloride peroxidase.


lost. Because the mitochondria migrate faster through the gradient than do the other subcellular components, the denser section of the gradient, containing the mitochondria, can be replaced after centrifuging for 5 hr.

2.5. Localization of Biochemical Markers (Supplementary Studies)

Fisher et al. (1968) and M ushahwar et al. (1972) report the presence of monoamine oxidase not only in mitochondria, but also in thyroidal endoplasmic reticular membranes. The enzymic reactions of both enzymes would proceed by similar pathways, the microsomal enzyme being susceptible to inhibition by anions. After discontinuous gradient centrifugation of a pellet previously sedimented at 105,000g for 1 hr, monoamine oxidase was recovered in all four fractions, whereas the mitochondrial marker succinic oxidase was found almost exclusively located in one fraction. However, it must be stressed that the method used to determine succinic oxidase (measured by oxygen uptake) depends on the presence of functionally intact respirating mitochondria, and therefore does not reflect the real distribution of mitochondrial material. Furthermore, we were able to demonstrate that especially in damaged mitochondria, even in the presence of succinate, respiration is hardly detectable (Hilderson and Dierick, 1973). In addition, when an M + L fraction was subjected to buoyant-density equilibration, monoamine oxidase (always coinciding with cytochrome c oxidase) could not be demonstrated in regions in which only endoplasmic reticular membranes are present. On the basis of these experimental data and the distribution profile of the enzyme shown in Figure 4, and because in other tissues this enzyme is located in the outer mitochondrial membrane, it is safe to use monoamine oxidase as marker for outer mitochondrial membranes in thyroid.

Cardiolipin (Section 4.2) can be considered an absolute marker for mitochondria because its relative specific concentration always coincides with the relative specific activities of both cytochrome oxidase and monoamine oxidase. Bovine thyroid mitochondria display a very low cholesterol-to-phospholipid molar ratio ( < 0.1). Therefore, the determination of this ratio is important in detecting contamination with other membranes (G. Van Dessel, A. Lagrou, H. J. Hilderson, and M. De Wolf, unpublished results).

When released from the lysosomes, acid phenyl phosphatase is probably nonspecifically adsorbed on other subcellular components, masking the real distribution of lysosomes. By means of isoelectric focusing, it was found that the enzyme displays three different isoelectric point values-pH 4.8, a continuous zone around 8.0, and pH 9.0 (Boddin et al., 1974). In addition, the enzyme activity is adsorbed at pH 5.5 on hydroxyl apatite-, CM-cellulose-, DEAE-cellulose columns and is only eluted at relatively high ionic strengths (Boddin et al., 1975). The enzyme also adsorbs on APUMP-agarose (agaroseaminophenyluridylic acid) columns and thereby interferes with the affinity


chromatography of thyroidal ribonuclease on these columns. (Boddin et al .. 1976). Addition of heparin or pyrophosphate to both suspension medium and gradient results in an increase of phenyl phosphatase at the top of the gradient. (This was shown to be caused by soluble enzyme by buoyant-density gradient centrifugation in a zonal rotor.) Moreover, the bulk of phenylphosphatase activity is eluted with the soluble protein fraction during Sepharose 2B column chromatography of an Sb fraction. (The suspension and elution was with 0.14 M NaCI/10 mM Tris/HCl buffer pH 7.4.) Summarizing, one can say that acid phenyl phosphatase is not an ideal marker for lysosomes.

Acid ribonuclease (isoelectric point values: 4.3, 6.3, and 9.7) also exhibits adsorption on the same columns. However, the enzyme activity is eluted at lower ionic strength. Phosphodiesterase (isoelectric point values 4.7-5.9) displays only minor adsorption. Therefore, acid ribonuclease and phosphodiesterase are better markers for lysosomes than is phenylphosphatase.

N-Acetylglucosaminidase exhibiting latent enzyme activity and displaying a lag period in isoosmotic incubation mixtures, nearly coincides with the other lysosomal hydro lases during isopycnic gradient experiments. It is probably a good marker for thyroid lysosomes.

It is generally believed that glucose-6-phosphatase is only present in the endoplasmic reticulum of tissues in which gluconeogenesis occurs (liver, kidney, intestinal mucosa). Tn those tissues glucose-6-phosphate can also be hydrolyzed by nonspecific phosphatases (Beaufay and de Duve, 1954a,b). As thyroid tissue contains only minor amounts of glycogen (Merlevede et al .. 1963), the hydrolysis of 80% or more of added glucose-6-phosphate by specific enzymes differing from lysosomal acid phosphatase is rather unexpected (Hilderson et al., 1976).

In bovine thyroid homogenates glucose-6-phosphatase activity occurs at a similar rate (2.5 /-Lmoles/min per mg protein at pH 6.5) to that in rat liver homogenates (Beau fay and de Duve, 1954a,b), whereas acid phenylphosphatase activity amounts to 625 /-Lmo1es/min per mg protein at pH 5 (ratio 1 : 250). In the subcellular fractions the following ratios of glucose-6-phosphatase : acid phenyl phosphatase activity were found: N fraction, 1 : 290; M fraction, 1 : 320; L fraction, 1 : 38; P fraction, 1 : 47; and supernatant, 1 : 166. Glucose-6-phosphatase activity is recovered chiefly in the M fraction, and there is appreciable enrichment in the L fraction. The distribution pattern is different from that of NADPH-cytochrome c reductase, 5'-nucleotidase, alkaline phosphatase, and acid phenylphosphatase.

No divergence between acid phenylphosphatase and glucose-6-phosphatase activities can be observed when measuring pH optima under different conditions. Tartrate ions stimulate phenylphosphatase activity in both M + Land P fractions, but slightly inhibit glucose-6-phosphatase activity. Both enzymic activities are strongly inhibited by fluoride ions, glucose-6-phosphatase activity being less affected.


Preincubation of a P fraction at pH 5 reduces both enzymic activities (phenyl phosphatase by 20%, glucose-6-phosphatase by 50%). In rat liver preincubation at pH 5 does not affect i3-glycerophosphatase, but inhibits completely glucose-6-phosphatase (Beaufay and de Duve, 1954a).

In sonicated thyroid microsomes deoxycholate inhibits both glucose-6-phosphatase and acid phosphatase activities. In nonsonicated microsomes increasing amounts of deoxycholate inhibit phenyl phosphatase activity, but stimulate glucose-6-phosphatase activity at low concentration and inactivate it at higher concentrations. This can be explained by the occurrence of a 30% latency for glucose-6-phosphatase that is unmasked by sonication and deoxycholate treatment. Therefore, glucose-6-phosphatase activity in thyroid can be associated with membranes present in the P fraction. This is in agreement with the results obtained for rat liver microsomes (Beaufay and de Duve, 1954b).

After treatment of an M + L fraction with increasing amounts of digitonin for 10 min at 50 C, a double S curve is obtained when glucose-6-phosphatase activity in the supernatant is plotted against digitonin concentration (Figure 5). The first part of the double-S curve does follow the acid phenylphosphatase activity, the second part follows NADPH-cytochrome reductase. These results are compatible with a dual localization of glucose-6-phosphatase activity. The part of the double-S curve coinciding with phenylphosphatase could reflect glucose-6-phosphatase activity owing to the presence of acid phosphatase (lysosomes), whereas the section of the curve that coincides with NADPH-cytochrome reductase could be attributable to the presence of a specific enzyme associated with endoplasmic reticulum membranes.

In highly purified nuclear fractions glucose-6-phosphatase activity is only present with the same relative specific activity as other endoplasmic reticulum enzymes (NADPH-cytochrome c reductase, peroxidases) (Voets et at., 1979b). As no separate endoplasmic reticulum membranes could be detected in those purified nuclear fractions by electron microscopy, glucose-6-phosphatase is probably present as a constituent of the outer membrane of the nuclear envelope.

After subjecting a sodium chloride extract of thyroid tissue to gel chromatography on Sephadex G-200, i3-glycerophosphatase does not display any glucose-6-phosphatase activity. The profile of glucose-6-phosphatase coincides with the profile of acid phenylphosphatase. Glucose-6-phosphatase to phenylphosphatase activities display a molar rate ratio of 1 : 1000. Kinetic studies on the phenyl phosphatase peak indicate that the hydrolysis of phenylphosphate and glucose-6-phosphate in this peak must be caused by the action of one single enzyme. Kinetic studies, however, on washed microsomal pellets suggest that the two substrates are hydrolyzed there by different enzymes.

The distribution pattern for glucose-6-phosphatase can be corrected by subtracting the glucose-6-phosphate hydrolysis owing to phenyl phosphatase from the total glucose-6-phosphate hydrolysis. Thus in the homogenate, 21 %

230

~ 100 « ~ z a: w c.. ~ 80 w :r: I-z >-!:: 60 > I-U « ...J « :2: 40 x « :2: w :r: l-LL. 20 o I-Z w U a: ~ 0

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i

I

I I

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H. J. Hilderson et al.

. ""~-----./ ................ "'" ",,"" ----

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o 5 10 15 20

FIGURE 5. The hydrolysis of glucose-6-phosphate (pH 6.5), phenylphosphate (pH 5), and NADPH-cytochrome c reductase activity in the 104,000g X 60 min supernatants after treatment of microsomal fractions with increasing amounts of digitonin (10 min,S 0 C). Abscissa scale: Amount of digitonin (mg/ml M + L 1 : 1). (_._.) Acid phenylphosphatase activity, (-) glucose-6-phosphatase activity, (- - -) NADPH-cytochrome c reductase activity.

of the glucose-6-phosphatase activity is caused by the presence of phenylphosphatase, in the N fraction 27%, in the M fraction 30%, in the L fraction 4%, in the P fraction 5%, and in the S fraction 17%. After correction, the distribution pattern becomes similar to the pattern for NADPH-cytochrome c reductase.

In gradient studies the soluble glucose-6-phosphatase activity at the top is almost entirely attributable to phenylphosphatase. Within the gradients glucose-6-phosphatase activity caused by acid phenyl phosphatase is only of the order of a few percent. The glucose-6-phosphatase distribution profiles corrected for phenylphosphatase activity nearly coincide with the profiles of total phospholipids and of NADPH-cytochrome c reductase (M + L fraction).


After Sepharose 2B column chromatography of an Sb supernatant fraction, the slowest migrating glucose-6-phosphatase activity is entirely attributable to acid phenylphosphatase. The activity eluted with the void volume is membrane bound and is not the result of phenyl phosphatase (De Wolf et ai., 1978b).

Summarizing, one can say that in bovine thyroid tissue the glucose-6-phosphatase activity is neither entirely caused by acid phenyl phosphatase nor by iJ-glycerophosphatase. The enzyme is not a good marker for differential pelleting, but can be used as a marker for endoplasmic reticulum membranes in fractionations involving centrifugation in a zonal rotor.

The presence of glucose-6-phosphatase in endoplasmic reticulum membranes has also been reported in other non liver tissues, such as small intestine (Lygre and Nordlie, 1969), brain, testes, pancreas, and adrenal (Collila et ai., 1975), lung, spleen, and erythrocytes (Nordlie et ai., 1965).

3. LOCALIZATION OF ENZYMES AND CONSTITUENTS IN BOVINE THYROID TISSUE

3.1. Subcellular Localization of Lipolytic Enzymes

The characterization and subcellular localization of phospholipases in bovine thyroid is of special interest because of their possible role in the biosynthesis of prostaglandins (Haye et ai., 1974) and their eventual involvement in some pathological conditions (e.g., cold nodules, Hashimoto's disease) (De Wolf et ai., 1976).

Bovine thyroid contains at least four phospholipase activities differing in their pH-optima and requirement for Ca2+ ions. These are two phospholipases with acid pH optima, one neutral phospholipase, and one with an alkaline pH optimum. Sodium taurocholate increases the acid A2-type activity (pH 4) and decreases the acid AI-type activity (pH 4) suggesting the presence of different enzymes. The lack of influence of Ca 2+ ions and EDT A and the acid pH optima point to a lysosomal localization (De Wolf et ai., 1976; De Wolf et ai., 1978a). The rather heat-stable neutral phospholipase activity (pH 6.5) is essentially of type A2 and is inhibited by Ca2+ ions. The specificity of the alkaline Ca2+stimulated phospholipase A activity (pH 8.5) cannot be determined because of interference by lysophospholipase (De Wolf et ai., 1979). The heat lability of alkaline phospholipase and its easy solubilization from the membranes are compatible with Al specificity (De Wolf et ai., 1978d). An additional acid lipase has been detected displaying a rather broad pH optimum (pH 4-6.5) (De Wolf et ai., 1978c).

Lipolytic enzymes in sucrose homogenates are 40-50% sedimentable (De Wolf et al.,1976). The neutral and alkaline phospholipases are more readily


solubilized than are the other lipolytic activities. They show their highest relative specific activity in the M fraction (Figure 2). Their distribution patterns differ from those of plasma membrane and mitochondrial markers.

More precise information about the subcellular localization of lipolytic enzyme activities is obtained by subjecting M + L or P fractions to isopycnic gradient centrifugation in a B-XIV zonal rotor (Section 2.4; De Wolf et al., 1977). A considerable amount of acid phospholipase AI, acid phospholipase A2, and lipase is found at the top of the gradient. In the gradient, acid phospholipase Al displays a bimodal distribution (densities 1.19 and 1.22 g/cm3).

Acid phospholipase A2 is mainly localized at the edge of the rotor, while activity decreases gradually toward lower densities. Lipase activity shows one peak between the densities 1.18 and 1.19 g/cm3, having a shoulder in the region of higher densities. The distribution profiles for acid phospholipase AI, acid phospholipase A2, and acid lipase closely parallel those of lysosomal marker enzymes. These enzymes can be considered lysosomal.

Neutral phospholipase A2 is mainly found at the top of the gradient. In the gradient, the distribution looks very similar to the distribution of lysosomal markers, especially ribonuclease. Centrifugation of a pellet obtained from a NaHC03 homogenate by 15 min at 73,300g in a HS-zonal rotor (De Wolf et al., 1977) results in a distribution profile that runs parallel with acid phosphatase, {1-glucosidase, and acid phospholipase A2. This enzyme is probably lysosomal.

For alkaline phospholipase A a completely different distribution profile is found, not paralleling that of marker enzymes for mitochondria, lysosomes, endoplasmic reticulum, or plasma membranes. As this alkaline phospholipase can only be detected in the M and L fractions, this enzyme could probably be particle bound and easily solubilized during the fractionation procedure. Its subcellular localization is therefore hard to establish.

Approaching the problem of the subcellular localization of lysophospholipase by isopycnic gradient centrifugation in a B-XIV zonal rotor does not immediately provide an unambiguous answer. Indeed, different distribution profiles for the enzyme were recorded depending on whether or not albumin was added in the enzyme assay mixture. The results obtained in the absence of albumin suggest that lysophospholipase is chiefly localized in more dense microsomal elements (perhaps rough endoplasmic reticulum). However, from the distribution profiles found in the presence of albumin in the assay mixture a bimodal localization could not be excluded (De Wolf et al., 1979). A double localization of lysophospholipase has already been reported in bovine liver (Vandenbosch and De long, 1975).

3.2. Subcellular Localization of Peroxidase Activities

It is generally believed that the iodination of thyroglobulin in thyroid is mediated by peroxidase (De Groot, 1977). Peroxidase activity can be detected


using guaiacol, p-phenylene diamine, benzidine, or 3,3'-diaminobenzidine as cosubstrates. It can also be assayed by measuring iodination or incorporation of radioactive iodide. However, one must be cautious when using 3,3'-diaminobenzidine, as proteins other than peroxidases could also be able to interact (Morrison, 1973). One must also keep in mind that it is also possible that different cosubstrates are oxidized by different enzymes.

When reviewing the literature relating to the subcellular localization of peroxidase activities and the iodination process, conflicting results and interpretations become apparent. Strum and Karnovsky (1970), using a cytochemical method with 3,3'-diaminobenzidinetetrachloride as cosubstrate, tend to localize the iodination process at the microvilli. However, Hosoya et al. (1973), using similar experiments, localize iodination in the endoplasmic reticulum. Novikoff et al. (1974) claim that iodination takes place in the colloid near the microvilli. Conflicting results are also obtained when applying autoradiography. Indeed, Strum and Karnovsky (1970) found labeling in the follicle lumen after 10 sec of incorporation. In contrast, Croft and Pitt-Rivers (1970) found the label inside the cells, provided that the incorporation time did not exceed 55 sec. When fixation is delayed for 2 min, the label is predominantly found over the peripheral region of the follicle lumen. Therefore, iodide seems to be captured initially within the cells. In agreement with these results, Edwards and Morrison (1976) demonstrated that after prefixation of the tissue, the label is localized within the cells and not in the follicle lumen. The bulk of the label was found at the level of the endoplasmic reticulum. Using biochemical methods, Hosoya et al. (1971) suggested that the localization of guaiacol peroxidase is in the rough endoplasmic reticulum. Also, Suzuki et al. (1977) found in their subfractions of bovine thyroid plasma membranes that the highest specific activity of guaiacol peroxidase was in that fraction that also had the highest specific activity for NADPH-cytochrome c reductase (which is an endoplasmic reticulum marker). Finally, Matsukawa and Hosoya (1979a,b) found that during incubation of thyroid slices with Na_131I, iodine atoms are preferentially incorporated into newly synthesized, less iodinated thyroglobulin, rather than into preformed thyroglobulin, and therefore suggest that the iodination occurs, at least to a certain degree, in apical vesicles before the thyroglobulin is secreted into the colloid lumen. Iodination seems to be a very rapid process, and the conflicting results found in the literature could be attributable to the speed of this phenomenon.

De Wolf et al. (1978b) systematically investigated the subcellular localization of thyroid peroxidase activity. After differential pelleting, peroxidase activities are present at their highest relative specific activity in the M fraction. After centrifugation of an M + L fraction for 24 hr in a HS-zonal rotor (buoyant-density equilibration) p-phenylenediamine, guaiacol, and 3,3'-diaminobenzidine tetrachloride peroxidase profiles do not coincide. In the presence of heparin (with or without digitonin) or pyrophosphate (Figure 4) the peak of guaiacol peroxidase always shifts along with glucose-6-phosphatase to lower

234 H. J. Hilderson et aI.

densities, whereas p-phenylenediamine peroxidase is less affected. In these experiments, catalase does not equilibrate with the peroxidase activities. Using electrophoresis on polyacrylamide gels and sucrose gradients, different profiles are found for guaiacol, 3,3'-diaminobenzidine tetrachloride, and p-phenylenediamine peroxidase activities. DEAE-Sephadex A-50 column chromatography of a Na2C03 extract of thyroid resulted in double peak distributions for all peroxidase activities (Figure 6). However, the percentage distribution of the individual peroxidase activities was different in both peak fractions; this can hardly be explained by the presence of one single-enzyme protein. Different Sepharose 2B elution profiles (Sb fraction, see Section 2.5) are also obtained for guaiacol, p-phenylenediamine, and 3,3'-diaminobenzidine peroxidase activities. From these results, one can conclude that a localization of peroxidase activity in plasma membranes or peroxisomes is unlikely, that different peroxidases are likely to exist in thyroid, and that guaiacol peroxidase is localized in rough endoplasmic reticular membranes. As guaiacol peroxidase always coincides with a part of the second peak of glucose-6-phosphatase, one can also say that this enzyme activity belongs to a specialized domain of the rough endoplasmic reticulum or to membranes very closely related to them, e.g., apical vesicles (Strum and Karnovsky, 1970) or A granules (Novikoff et al., 1974).

3.3. Subcellular Localization of RNA-Polymerase Activity

Different classes of RNA-polymerase activities have been described in eukaryotic cells. In calf thyroid, enzyme forms I and III both consist of at least two components (lA, IB, IlIA, IIIB) (Spaulding, 1977). A large fraction of these enzyme activities are recovered in the cytoplasm, (lJthough the more logical localization for these enzymes would be in the nucleus. These results could be attributable to a procedural artifact, or they may reflect the real situation. One could argue that RNA-polymerase molecules, not engaged in a transcription complex with chromatin (free RNA-polymerase), could either (in vivo) diffuse freely to the cytoplasm or be squeezed out of the nucleus by the cell fractionation technique (Austoker et al., 1974). A quantitative comparison of RNApolymerase activities in different subcellular fractions is difficult to achieve. In the presence of 0.25 M ammonium sulfate, an inhibitor of the initiation of transcription, RNA-polymerases found in a transcription complex with chromatin (bound RNA-polymerase) are chiefly measured. In the absence of ammonium sulfate, both free and bound RNA polymerase activities are measured. However, the amount of endogenous DNA differs in each subcellular fraction. The optimal conditions for RNA-polymerase determination differ for each enzyme type and differ further in each subcellular fraction. During the fractionation procedure, both free and bound RNA-polymerase activities can be released in different ways. Finally, regulatory factors may be distributed unequally


60 0.6 a

z 40 0.4 ::ii :::) ...J 0 U UJ :r: 20 0.2 I-::ii 0 a:: LL.

0 UJ a:: 0 0.0 UJ > 0 u UJ a:: b >- l t: 60 i\ > j:: i.\ u « n, UJ :r: in \ I- 40 LL. it \i 0 ·f \ i l- I, :. z .: !\ UJ U f !' a:: - : \ UJ 20 f \ \ c..

,\ : \ \ " .

0 .. ' : ...................

I I I I I 0 10 20 30 40

FRACTION NUMBER

FIGURE 6. Chromatography on DEAE-Sephadex A-50 of an Na2CO, extract of a P fraction. A sample of the Na2CO, extract (3 ml, 6 mg of protein/ml) was loaded on the column (2.5 cm X 12 cm) and eluted first with 50 ml 0.01 M Na2CO" pH 10.5, and then with 200 ml of a linear gradient of 0-0.6 M NaCI in 0.01 M Na2CO" pH 10.5. KI (0.1 ffiM) was added to the eluent to stabilize the enzyme preparation. (a) (- - -) Catalase; (. -' -) iodination of monoiodotyrosine; (--) iodination of tyrosine; (.-. -) slope of NaCI gradient. (b) (. -' -) Guaiacol peroxidase; (. . .) p-phenylenediamine peroxidase; (--), 3,3'-diaminobenzidine tetrachloride peroxidase.


throughout the subcellular fractions. The distributions recorded for RNApolymerase activities can therefore only be qualitative.

Lin et al. (1976) extracted RNA-polymerase activity from rat liver nuclei isolated in isotonic or hypertonic sucrose. Activities of RNA-polymerase I, II, and III from nuclei isolated in hypertonic media were much higher (3.8-, 1.5-, and 27-fold, respectively) than those from nuclei isolated in isotonic media. RNA-polymerase III was virtually absent in the latter nuclei. Concurrent with the reduced recovery of polymerases from these nuclei, the corresponding cytosol fraction contained higher level of all three enzymes. Thus a large proportion of the "cytoplasmic" RNA-polymerase III in "isotonic cytosol" fraction corresponds to nuclear RNA-polymerase III. Furthermore, Kellas et al. (1977) were able to show that RNA-polymerase I activity exists as two discrete pools-a pool of "free" activity (IA) and a pool of enzyme engaged in a transcription complex (Is). The free form of the enzyme is lost to the cytoplasmic fraction on nuclear isolation.

In Chinese hamster kidney cells Austoker et al. (1974) found RNA-polymerase I activity in both nuclear and cytoplasmic fractions. In both fractions the addition of DNA more than doubled the apparent activity of this enzyme, which supports the view that unbound polymerase I exists in those cells. No free RNA-polymerase II seems to be present in the nuclear or cytoplasmic fraction. Bound RNA-polymerase II is found to be concentrated in the nucleus. The proportion in the cytoplasm (15.6%), corresponding closely to the amount of total cellular DNA in this fraction, is probably due to nuclear damage. RNA-polymerase III (free and bound) is found in both nuclei and cytoplasm.

In bovine thyroid homogenate RNA-polymerase I, II and III, measured in 0.25 M ammonium sulfate (high ionic strength, conditions more optimal for bound RNA-polymerase II determination), account, respectively, for 10.9%, 84.7%, and 4.3% of total RNA-polymerase activity. In the absence of ammonium sulfate (low ionic strength, conditions more optimal for RNA-polymerase I determination) the percentage values are, respectively, 58.3%, 38.3%, and 3.4%. Table I shows the composition of the RNA-polymerase activity in the fractions obtained during the preparation of a nuclear fraction according to Widnell and Tata (1966) (see also Section 4.1). Table II summarizes the percentage activities of the different RNA-polymerase activities in each isolated fraction. After differential pelleting, the distribution pattern for total RNApolymerase activity in the presence of 0.25 M ammonium sulfate coincides with the distribution pattern for endogenous DNA, indicating that under these experimental conditions bound RNA-polymerase II is chiefly measured (Voets et al., unpublished results).

On the basis of these experiments, one can say that bound RNA-polymerase II is almost completely recovered in the nuclei, a portion (4.5%) of the enzyme being extracted during the centrifugation step through 2.2 M sucrose.

Subcellular Biocbemistry of Thyroid

Table I Composition (% ) of the RNA-Polymerase Activity in Fractions

Obtained during the Isolation of Bovine Thyroid Nuclei"

Low ionic strength High ionic strength

Fractionb II III II III

H 58.3 38.3 3.4 10.9 84.7 4.3 S\ 79.2 17.1 3.7 77 NO' 23 N\ 24.1 67.0 8.8 5.8 90.2 4 S, 52.3 47.7 NO 3.2 91.3 5.5 N, 19.9 76.8 3.16 5 91.7 3.3

'Voets et al. (1978), according to Widnell and Tata (1966). bA homogenate (H) prepared in 0.25 M sucrose-3 mM MgCl, (pH 7.4) is layered on 0.32 M sucrose-I mM MgCl, (pH 7.4), and centrifuged for 10 min at 700g. The supernatant (S\) is aspirated, and the sediment (N I ) is purified by centrifuging through 2.2 M sucrose-I mM MgCl, (pH 7.4). The subsequent pellet is designated as N, and the supernatant as S,.

'ND, Not detectable.

237

The major part of the RNA-polymerase I is recovered in S" some activity being released from the nuclei during the final purification step. RNA-polymerase III behaves in an intermediate way. In an S, fraction the bulk of the activity is accounted for by polymerase I. In the nuclear fraction RNA-polymerase II represents the major part of the enzyme activity.

Table II Percent" and Relative Specific Activities (RSA) of RNA-Polymerase Activitiesb

1+11+111 II III

% RSA % RSA % RSA % RSA

Low ionic strength S·' , 32 0.32 65 0.7 15 0.17 43.7 0.5 N\ 63 14 30 7 84 19 71 16 S, 4 1.5 6.5 2 4.5 1.5 NOd NO N, 62 60 27 25 82 78 56.3 54

High ionic strength S\ 7 0.07 58 0.6 NO NO 40 0.4 N\ 92 21 40 9 92 20.9 66.4 15.1 S, 5 1.7 1.5 0.5 4.6 1.5 5.7 1.9 N, 87 83 38 36 95 90.4 54.4 52

'Homogenate taken as 100%, mean values of at least two experiments. bFractions obtained during the isolation of bovine thyroid nuclei according to Widnell and Tata. 'Sj, Nj, S" N,; see Table I. "NO, Not detectable.


Summarizing, one can say with certainty that in bovine thyroid RNApolymerase II is localized exclusively in the nuclei. The possibility exists that this is also true for RNA-polymerase I and III although one cannot yet exclude a double localization of these enzymes in nuclei and cytoplasm.

4. ISOLATION AND CHARACTERIZATION OF THYROID ORGANELLES, SUBCELLULAR COMPONENTS, AND MEMBRANES

4.1. Nuclei

For the isolation of bovine thyroid and bovine liver nuclei a method adapted from that of Widnell and Tata (1966) was used (Hilderson and Dierick, 1974). After homogenization in 0.32 M sucrose solution (3 mM with respect to MgCI2, adjusted to pH 7.4 with NaHC03), the homogenate is filtered through two layers of cheesecloth. Per 25 ml filtered homogenate are added 15 ml chilled 0.32 M sucrose solution (3 mM MgCI2, pH 7.4) and 12 ml chilled glass distilled water. Portions of this diluted homogenate are layered over 0.32 M sucrose and centrifuged at 700g for 10 min. The pellets (N] in Tables I and II) consist of red blood cells, cellular debris, nuclei, and some contamination with other elements. Most of the postnuclear fraction (fraction S]), however, is retained in the top layer. The crude pellet is further purified by centrifugation through 2.2 M sucrose (l mM MgCI2, pH 7.4) yielding nuclear fraction A (N2 in Tables I and II) and a supernatant fraction S2. To avoid damage during resuspension, the A pellet is left standing overnight (4 0 C, 0.32 M sucrose, 3 mM MgCI2, pH 7.4), after which it is resuspended easily and centrifuged at 1000g for 20 min, yielding the purified nuclear fraction B.

During the run in the A-XII zonal rotor, no DNA and RNA is lost from the nuclei. It cannot be precluded that other material escapes from the nuclei. No respiratory activity can be detected in this nuclear fraction C (Hilderson and Dierick, 1973). In nuclear fraction C the relative specific concentration* for DNA is 70. For RNA-polymerase, the relative specific activities range from 31 to 150, depending on the assay conditions. Therefore, one can conclude that a highly purified nuclear fraction has been obtained.

From the rate of sedimentation in the A-XII zonal rotor, the density of the nuclei at any given point in the sucrose gradient can be computed (Hilderson and Dierick, 1974). For normal thyroid nuclei, an isopycnic density of 1.41 g/ cm3 is found, for hypertrophic thyroid nuclei 1.43 g/ cm3, and for bovine

*The relative specific concentrations were measured with respect to homogenate protein including follicular colloid.

Subcellular Biochemistry of Thyroid

Table III Protein, RNA, and DNA Content in Bovine Thyroid and Liver

Nuclei"

Normal Hypertrophic Content thyroid nuclei thyroid nuclei Liver nuclei

Proteins 35.2 ± 5.4 60.4 ± 8.4 53.9 ± 11.9 DNA 14.4 ± 1.7 13.7 ± 2.0 11.9 ± 1.2 RNA 2.9 ± 0.5 3.4 ± 0.8 3.9 ± 0.7 DNA/protein 0.422 ± 0.076 0.260 ± 0.056 0.196 ± 0.021 RNA/protein 0.082 ± 0.017 0.065 ± 0.016 0.069 ± 0.012 RNA/DNA 0.214 ± 0.051 0.255 ± 0.059 0.330 ± 0.062

'In picograms per nucleus.

239

liver nuclei 1.30 g/cm3• It is also possible to compute the sucrose impermeable space in thyroid nuclei-3.88%, whereas the value for bovine liver nuclei is 7.76%. Johnston et al. (1968) report values of 10.2% for parenchymal rat liver nuclei and 23.7% for mouse liver nuclei.

The protein RNA and DNA contents in bovine thyroid and liver nuclei are represented in Table III. The percentage composition of both normal and hypertrophic thyroid nuclei is given in Table IV. For hypertrophic thyroid nuclei, it is similar to that of rat liver nuclei. The high percentage value found for protein in hypertrophic thyroid nuclei causes a drop in percentage values for the other components. RNA and DNA contents are similar in both kinds of nuclei. Comparing those DNA values with values for rat liver nuclei (Johnston et aI., 1968) one can conclude that adult bovine thyroid and adult bovine liver are chiefly tetraploid.

The mean nuclear diameter for normal thyroid tissue ranges from 6 to 10 ~m (with a peak at 8.4 ~m). For hypertrophic thyroid tissue the diameter ranges from 6 to 12 ~m (with a peak at 9.0 ~m; 1.8% of the nuclei have a

Table IV Percentage Composition of Normal and Hypertrophic Thyroid Nuclei

Tissue Protein DNA RNA Phospholipids

Bovine hypertrophic 76.0 17.2 4.3 2.4 thyroid nuclei

Bovine normal 64.6 26.4 5.32 3.66 thyroid nuclei

Rat liver nuclei 72.4 20.0 3.4 4.1 (Kay et al .. 1972)


diameter> 10.5 Jim). The range is 6-11.1 Jim (with a peak at 8.3 Jim) for bovine liver. Histograms for fetal thyroids show the normal pattern in fetuses from mother animals having a normal thyroid. Fetuses from mothers having a hypertrophic thyroid display the hypertrophic pattern. Therefore, one must conclude that anti thyroidal agents do cross the placental barrier and reach the developing foetus in utero.

The nuclei are relatively large and contain only small amounts of lipid. The determination of their true lipid composition is difficult, since mitochondrial contamination especially, and/or microsomal contamination could give rise to error (Van Dessel et al .. 1979a). Mitochondrial contamination could include cardiolipin, for example, and microsomal contamination could include dolichol. Therefore, highly purified nuclear fractions (nuclear fraction C) are required. Moreover, precautions must be taken in order to avoid postmortem changes of the lipid composition.

The results for both normal and hypertrophic thyroid are shown in Table V. The lipid composition of hypertrophic thyroid nuclei does not differ essentially from that of normal thyroid nuclei. In both cases cholesterol is the more important neutral lipid ( ~65% of total neutral lipids). The molar ratio cholesterol/phospholipid is 0.09. Cholesterol esters account for less than 1 % of total cholesterol content. The percentage phosphorus distribution shows phos-

Table V Lipid Composition of Bovine Thyroid Nuclei"

Composition

Total lipids Free cholesterol Cholesterolesters Total phospholipids

Normal bovine thyroid nuclei

2.2 ± 0.1 0.15 ± om

less than I % of cholesterol 2.0 ± 0.1

Hypertrophic bovine thyroid nuclei

2.1 ± 0.5 0.16 ± 0.01

1.9 ± 0.4

% Distribution of total phospholipids (PC + PE + PI + PS + SM = 100%)

Phosphatidylcholine 58.5% 60.7% Phosphatidylethanolamine 25.7% 23.6% Phosphatidylinositol + 15.4% 13.0%

phosphatidylserine (PI + PS) Sphingomyelin 0.65% 2.3% Other (% total phospholipids) Not determined 6.3 ± l.l

"All values. except for the percent distribution of the phospholipids, are expressed in picograms per nucleus and represent the mean values for at least 10 experiments.


phatidylcholine to be the main phospholipid. A very low value is found for sphingomyelin. Phosphatidylethanolamine, phosphatidylserine, and phosphatidylinositol are present in intermediate concentrations. Extremely low values « 1 %) are found for lysoderivatives, indicating the absence of "postmortem" changes. The nature of the remaining phospholipids is still uncertain. The presence of glycolipids cannot be demonstrated. Our data are in good agreement with the results reported elsewhere (Rouser et aI., 1968), except for the low values for lysoderivatives. Our results support the statement by Rouser et at. (1968) that there seems to be no striking organ specificity for nuclear membranes.

From the phospholipid content one can calculate the number of monolayers. In thyroid nuclei mean values of 5-6 were found. These figures represent more than the amount of lipid required to build a double envelope. However, it is possible that (1) the distance between two phospholipid molecules is overestimated in the calculations (Vandenheuvel, 1965); (2) the molecules are stacked more closely in a membrane than in a Langmuir trough; (3) the nuclear surface is not necessarily smooth, but displays a "pleated sheet" aspect; (4) nucleoplasmic lipids interfere; or (5) some fragments of endoplasmic reticulum remain attached to the nuclear envelope during the entire procedure.

4.2. Mitochondria and Lysosomes

In order to investigate the percentage of total proteins contributed by mitochondria and lysosomes to a thyroid homogenate, enriched mitochondrial and lysosomal fractions were prepared. As in sucrose density gradients, mitochondria and lysosomes move faster than do the other cell components, the denser part of the gradient containing the mitochondrial and lysosomal peaks were removed after 5 hr of centrifugation of an M + L fraction in a HS zonal rotor (De Wolf et at., 1978b). In the mitochondrial peak fraction the relative specific activities of both cytochrome oxidase and monoamine oxidase ranged from 40 to 60, the relative specific activities of the other components being lower than in the original M + L fraction. In the lysosomal peak fraction, the relative specific activities for the hydrolases ranged from 10 to 15. Using the method of Leighton et at. (1968) to calculate the % contribution of each organelle to the total protein in the homogenate, a value of 2.0 ± 0.8% was found for thyroid mitochondria and 3.6 ± 2.2% for thyroid lysosomes (G. Van Dessel, A. Lagrou, H. J. Hilderson, and M. De Wolf, unpublished results).

Total lipid content of the purified mitochondrial fraction amounted to 0.39 mg/mg protein. Phospholipid accounted for 85-90% of the lipid. Phosphatidylcholine (39%), phosphatidylethanolamine (26%), cardiolipin (10%), and


sphingomyelin (12%) were the major mitochondrial phospholipids. In contrast to other tissues, thyroidal mitochondria contain relative high amounts of sphingomyelin and acid phospholipid. This was also reported by Satyaswaroop (1971). However, whether this finding is an artifact or not remains to be resolved. Cholesterol and cholesterolesters represent 6.8 and 2.8%, respectively, of total lipid (G. Van Dessel, A. Lagrou, H. J. Hilderson, and M. De Wolf, unpublished results).

Van den Hove-Vandenbroucke and De Nayer (1976) have studied the separation of pig thyroid subcellular organelles in metrizamide gradients. Using continuous gradients, it was found that intact mitochondria banded at 1.18 gjcm3, and mitochondria damaged by hydrostatic pressure banded at 1.14 and 1.16 gj cm3• Two different lysosome species « 1.14 and 1.15 gj cm3) were found. After centrifuging an M + L fraction (prepared in 0.25 M sucrose) through a dense pad of 20% (w jv) metrizamide more than 70% of the cytochrome c oxidase was recovered in the pellet. The authors report that this pellet is not contaminated with lysosomal material (mitochondrial pellet). About 40% of the lysosomal material remained at the interface. After washing this material with 0.25 M sucrose, the resulting lysosomal fraction was essentially free of mitochondria and showed high latent enzyme activity for {3-glycerophosphatase. The final lysosomal fraction represented 8-10% of the initial hydrolytic activity of the homogenate, and it retained its high buoyant density in sucrose gradients. However, applying this procedure in our laboratory, a substantial contamination of the so-called mitochondrial pellet with lysosomes was noted as 19% of N-acetylhexosaminidase and 15% of acid phenylphosphatase activity present in the M + L fraction were recovered in this pellet. Thirty-five percent of the glucose-6-phosphatase activity was also found in that fraction, indicating contamination with endoplasmic reticulum membranes. Furthermore, the socalled final lysosomal fraction still contained some cytochrome c oxidase activity. We did obtain better results using a dense pad of 30% (w jv) metrizamide, resuspending the pellet in 0.25 M sucrose and centrifuging again through 30% (w Iv) metrizamide. In this way a mitochondrial pellet was obtained with a 12% yield of protein and with only traces of acid hydrolase activity. No 5/-nucleotidase could be detected. At the interface, about 40-50% of the acid hydrolases, 50% glucose-6-phosphatase, and 30% 5'-nucleotidase, alkaline phosphatase, and NADPH-cytochrome c reductase were recovered. Ten percent of the cytochrome c oxidase activity was found in that interface. After washing most of the cytochrome c oxidase activity was removed. However endoplasmic reticular and plasma membrane markers could still be detected.

Finally Simon et al. (1979) were able to separate three sUbpopulations of lysosomes from rat thyroid by using Sepharose 2B filtration. Each subpopulation consists of secondary lysosomes, digesting thyroglobulin corresponding to

Subcellular Biochemistry of Tbyroid 243

different compartments (a rapid, slow, and very slow compartment) in thyroid cells.

4.3. Golgi-Rich Fractions

For thyroglobulin synthesis it is generally believed that the internal sugars are added as soon as the growing polypeptide chain moves into the channels of the endoplasmic reticulum. Spiro and Spiro (1973) suggest that the peripheral sugars (sialic acid, galactose) are added later on (by the smooth endoplasmic reticulum, the Golgi apparatus, or even plasma membranes). In order to test this possibility it is important to separate endoplasmic reticular membranes from Golgi apparatus and plasma membranes. The resolution obtained by differential pelleting is not good enough. Better results are obtained with gradient centrifugation (Spiro and Sprio, 1973; Chabaud et al., 1974; Ronin and Bouchilloux, 1978). Spiro and Spiro (1973) centrifuged light particles in a discontinuous density gradient and showed a distribution of sialyl- and galactosyltransferases different from that of the mannosyltransferase. The former enzymes are found in the low-density fractions together with 5' -nucleotidase. Mannosyltransferase is more concentrated in the high-density fractions together with RNA and NADPH-cytochrome c reductase.

Chabaud et af. (1974) used a discontinuous sucrose gradient in order to subfractionate their microsomal fractions isolated in stabilizing medium for Golgi membranes. Their fraction 1 (0.4-1.05 M sucrose interface) is highly enriched in morphologically recognizable Golgi fragments (Bouchilloux et al., 1970), and its RNA protein ratio is very low. The glycosyltransferases (sialyltransferase, galactosyltransferase, N-acetylglucosaminyltransferase) exhibit a distribution profile quite distinct from that of plasma membrane markers. These authors suggest that their results are consistent with a common preferentiallocalization of glycosyltransferases in the thyroid Golgi apparatus. They also suggest that N-acetylglucosamine, and perhaps even galactose, would be incorporated at an earlier stage, possibly at the periphery of rough endoplasmic reticulum. However, neither Spiro and Spiro (1973) nor Chabaud et al. (1974) could localize the different glycosyltransferases with certainty. Absolute markers for both apical plasma membranes and Golgi apparatus would be necessary to solve this problem.

In order to localize the glycosyltransferases involved in the ganglioside biosynthesis in bovine thyroid Pacuszka et al. (1978) attempted to isolate Golgi membranes by centrifuging a 300g X 10 min supernatant, layered on 1.25 M

sucrose. The yellow layer, formed at the interphase, is collected and washed. This fraction represents 3 mg protein/l0 g thyroid gland (recovery: 0.3%). It contains less than 1 % of 5'-nucleotidase and glucose-6-phosphatase. No total


recoveries (balance sheet) are reported, so that it is not possible to exclude inhibition or activation phenomena. Finally, it is not shown that this subcellular fraction corresponds to the morphological entity called Golgi apparatus. However, from this study one can indeed conclude that bovine thyroid gland contains the glycosyltransferase activities that constitute a biosynthetic route for the stepwise formation of GD 1• * from lactosylceramide. The glycosyltransferases are highly enriched in the interface fraction (ranging from 77- to 97-fold). Thyroid tissue also contains the glycosyltransferase activities required for another sequential synthesis of GD3, GD2, GD1b, and GT1b, starting from GM3•

Three of the enzyme activities (catalyzing the formation of GD2, GD1b, and GT1 b) are highly enriched in the interface fraction. In contrast CMP-N-acetylneuraminic acid: GM3-sialyltransferase activity does not appear to be predominantly localized in that fraction. This observation could imply that differential subcellular localization of the glycosyltransferase may be an important factor in the regulation of ganglioside biosynthesis.

4.4. Protein-Synthesizing Polyribosomes

Several authors have used polyribosomal cell-free systems from thyroid in an attempt to study the steps leading to the formation of the thyroglobulin molecule. Nunez et al. (1965) and Morais and Goldberg (1967) prepared microsomal fractions from sheep and calf thyroid and showed that these incorporated labeled amino acid into 19 S thyroglobulin. They showed the presence of several successive peaks corresponding to different classes of polyribosomes. Kondo et al. (1968) found that the distribution throughout a sucrose gradient of ribosomes and polysomes were similar to those from other mammalian tissues. The heavier polyribosomes (containing lO-35 ribosomes) showed a higher proportion of incorporation into thyroglobulin-related protein than did the smaller ones. Furthermore, De Nayer and De Visscher (1969) were able to demonstrate that in order to obtain labeled thyroglobulin, nonradioactive thyroglobulin must be present in the incubation medium. They suggested that an exchange of subunits takes place between newly synthesized and mature thyroglobulin molecules. Lecocq et al. (1971) evaluated quantitatively polysomes and ribosomes and found the following distributions: 15-20% monomers, 4% of dimers, 5% trimers, 7% tetramers, 7-8% pentamers, 9% hexamers, and 50-60% heavy polysomes.

Chebath et al. (1977a,b) prepared thyroglobulin-specific polysomes using

*The nomenclature of Svennerholm (1970) is used: GM3• monosialosyl-Iactosylceramide; GD3•

disialosyl-Iactosylceramide; GD2• disialosyl-N-triglycosylceramide; GD.a• disialosyl-N-tetraglycosylceramide; GD. b• disialosyl-N-tetraglycosylceramide; GT.b• trisialosyl-N-tetraglycosylceramide.


an indirect immunoprecipitation technique. Thyroglobulin-specific polysomes represented 30-45% of the total polysomes. These workers also demonstrated that synthesis of thyroglobulin peptides in a reticulocyte cell-free protein synthesizing system was three times greater when using mRNA from immunoprecipitated polysomes than with total mRNA. Finally, they showed that the specific mRNA displays a molecular mass of 2.8 X 106 daltons (enough to code for the thyroglobulin peptide chain). This report was confirmed by Vassart et al. (1977).

Van Voorthuizen et al. (1978) using frozen tissue as starting material for the preparation of polysomes and thyroglobulin mRNA were able to show that normal goat thyroid contains a population of large membrane-bound polysomes engaged in thyroglobulin synthesis. In contrast, such polysomes are absent from the thyroid of goats with hereditary congenital goiter and thyroglobulin deficiency. The authors suggest that there could be a defect in the processing of thyroglobulin (pre)mRNA.

4.5. Plasma Membranes

Plasma membranes from thyroid are characterized by the presence of specific receptors for thyroid stimulating hormone (TSH) and by the occurrence of active transport for iodide. Gangliosides and phospholipids would be involved in processing the TSH signal from the extracellular into the intracellular compartment of the thyroid cell (Kohn, 1978; Van Dessel et al., 1979b). Phospholipids could also playa role in the active transport of iodide (Lagrou et al.. 1974b).

Plasma membranes display similar densities as smooth endoplasmic reticular membranes. Therefore cell rupture techniques must be used that avoid formation of small or mixed vesicles, or both, and that produce large sheets of plasma membrane that can be sedimented at low centrifugal forces. A tissue as "solid" as thyroid contains more collagenous tissue and fibrous material than liver tissue. This results in serious problems in homogenizing (Evans, 1978).

The different methods described in the literature do not control sufficiently contamination through other cell components. Too few marker enzymes are assayed and only few data are reported concerning recovery (i.e., by using a balance sheet). Therefore some authors refer to their isolated fractions only as to "membranes" (Amir et al., 1973; Moore and Wolff, 1973, 1974; Macchia and Meldolesi, 1974; Smith and Hall, 1974; Bhattacharyya and Wolff, 1975; Mehdi and Nussey, 1975; Ichikawa et al .. 1976; Moore and Feldman, 1976; Davies et al .• 1977; Ashbury et al.,1978).

Two methods are generally applied: (1) a homogenization procedure (Neville, 1960) in buffered water (NaHC03, phosphate buffer, Tris HCl


buffer) (Yamashita and Field, 1970; Amir et aI., 1973; Moore and Wolff, 1973, 1974; Roques et aI., 1975; Suzuki et aI., 1977); and (2) a homogenization procedure in isoosmotic sucrose or in 0.4 M sucrose (Stanbury et al., 1969; Wolff and Jones, 1971; Macchia and Meldolesi, 1974; Mehdi and Nussey, 1975). All methods use gentle homogenization techniques.

Yamashita and Field (1970) isolated bovine thyroid plasma membranes by the method of Neville (1960). A 2500-rpm pellet from a NaHC03 homogenate is mixed with 63% sucrose. After flotation through a discontinuous sucrose gradient, the material at the interface between 1.16 and 1.18 g/cm3

layers of density is collected as the plasma membrane fraction. Electron microscopy shows that structures in this fraction consist of vesicular membranous material and a layer consisting of plasma membrane sheets with a minimal contamination with other cellular components. The relative specific activity of the adenyl ate cyclase activity in this fraction is 10-20 times that of the whole homogenate, and the ATPase relative specific activity 10 times. Cytochrome c oxidase cannot be detected but NADPH-cytochrome c reductase has a sixfold increase in its specific activity. Amir et al. (1973) and Azukizawa et al. (1977) also isolated plasma membrane-enriched fractions according to the Neville method. The yield of their purified membranes is 0.54 mg/g wet thyroid tissue. Electron micrographs show that the membranes form large vesicular structures. Amir et al. (1973) report minimal contamination of other subcellular components. The relative specific activities of membrane bound enzymes are increased 10- to 40-fold (5'-nucleotidase, 11; ATPase, 19; adenylate cyclase, 42), whereas the relative specific activities of lysosomal and mitochondrial enzymes are reduced approximately 10-fold (acid phosphatase, 0.11; succinic dehydrogenase, 0.25), as well as the endoplasmic reticular membrane markers (glucose-6-phosphatase, 0.08; NADPH-cytochrome c reductase, 0.12). However, no overall recoveries of the enzymic activities are reported (by using a balance sheet) so that the detection of activation or inhibition phenomena was not possible. Indeed, the higher rise in relative specific activity of adenyl ate cyclase (42) in comparison to 5'-nucleotidase (11) might be the result of a more efficient activation of adenylate cyclase in purified fractions (Yamashita and Field, 1970). This could also be the case for Mg2+_ ATPase. In our laboratory a pellet isolated from a NaHC03 homogenate by 15 min at 3000g was subjected to a similar discontinuous gradient centrifugation. The bulk of the plasma membranes was recovered from the 30-40% sucrose interface. However, appreciable contamination was noted. The final relative specific activity values were: cytochrome c reductase, 8; glucose-6-phosphatase, 6.4; cytochrome oxidase, 7.2; hexosaminidase, 2.8; acid phosphatase, 2.4; alkaline phosphatase, 4.9; and 5'-nucleotidase, 3.8. A similar distribution of membranes was also found with a pellet isolated after 15 min at


39,000g (Jansegers et af., unpublished results). Roques et af. (1975) isolated plasma membranes from porcine thyroid glands by the same method. The yield of purified membranes was about 1 mg/IO g thyroid. The activities of 5'nucleotidase and alkaline phosphatase were concentrated in the plasma membrane fraction, the respective relative specific activities being 7.9 and 12.0. As shown by the content of NADPH-cytochrome c reductase (relative specific activity 4), and cytochrome c oxidase (relative specific activity 1.4), the authors were unable to avoid the presence of other subcellular material in the plasma membrane fraction. A protamine kinase activity was found to be present in this fraction, displaying a relative specific activity of 10-12.

Verrier et af. (1976) isolated membranes from thyroid cells obtained by trypsinization of porcine glands and maintained in culture conditions in the presence or absence of thyrotropin or dibutyryl cyclic AMP. As shown by electron microscopy the plasma membrane fraction displays a high degree of purity. In the presence of thyrotropin adenyl ate cyclase was stimulated (Verrier et af., 1974). The protein, phospholipid, cholesterol, and sialic acid content of the three types of plasma membrane preparations used by Verrier et af. (1974) are very similar. The molar ratio reported for phospholipid/cholesterol (0.8) would mean that more cholesterol than phospholipid molecules are present in the plasma membranes (a somewhat surprising result). The amino acid and carbohydrate composition of the plasma membranes is similar to that of other eukaryotic plasma membranes. SDS-polyacrylamide gel electrophoresis disclosed the presence of more than 20 protein bands, of which six correspond to glycoproteins. (The apparent molecular weight of the bands ranged from 24,000 to 240,000.) Aspartic and glutamic acid, leucine, alanine, and glycine are the more abundant amino acids, whereas the sulfur-containing amino acids and histidine are less represented.

Suzuki et af. (1977) isolated two subfractions of plasma membranes (H and L: heavy and light membranes) from bovine thyroid glands using a slight modification of the method of Yamashita and Field (1970). The specific binding of [125I]_TSH is similar in both fractions. The H-membranes have greater adenyl ate cyclase activity. The L-membranes have higher relative specific activities for 5'-nucleotidase and MgH -ATPase. The relative specific activities of (Na +, K+)ATPase and alkaline phosphatase are similar in the two fractions. Contamination with other subcellular components cannot be excluded. The extent to which these membrane fractions are representative is subject to question, as the recovery of proteins (0.08%) and marker enzymes in the plasma membranes is very low (Evans, 1978). The H- and L-membranes, differing in some of their enzyme activities, may represent different parts of the plasma membrane. Electron microscopy demonstrates that plasma membranes and Hmembranes are very similar as both fractions contain junctional complexes,


long membrane sheets and vesicles. The L-membranes consist mainly of short membrane sheets and vesicles and of only a few junctional complexes.

Investigating the subcellular localization of the Long-Acting Thyroid Stimulator Inhibitor (LATS inhibitor)* in bovine thyroid gland, Nitiyanant and Dunlap (1978) prepared plasma membranes by the method of Yamashita and Field (1970), with the exception that a continuous sucrose gradient was employed. Only 2% of the original adenyl ate cyclase activity was found in their SG 1 fraction (largest increase of specific activity). SG 1 is the fraction corresponding to the membranous material equilibrating at a density of 1.10 g/cm3

during continuous gradient centrifugation. No microsomal NADPH-cytochrome c reductase could be detected in SG 1• No yield for protein or other enzymes in this fraction nor data allowing computation of relative specific activities were reported. On lower-power electron micrographs, occasional microsomes are seen, constituting only a very small fraction of the recognizable structures. The authors do not indicate how smooth endoplasmic reticular membranes are differentiated from plasma membranes. High-power electron microscopy shows only minor contamination.

The second method of purifying plasma membranes from thyroid tissue gland begins with a homogenization procedure in isotonic or slightly hypertonic sucrose media. Divalent ions are generally added to reduce vesicularization. Stanbury et al. (1969) disrupt calf and human thyroids by nitrogen microcavitation. Centrifuging of a pellet previously prepared by 45 min at 104,000 g and layered over ficoll yielded two bands, one just below the interface and another at the bottom of the tube. The lighter fraction, consisting of somewhat larger vesicles, had been enriched in fragments derived from plasma membranes(Na+, K+)ATPase, 5'-nucleotidase, and sialic acid were concentrated in that fraction. The authors were unable to show adenyl ate cyclase activity. Substantial contamination by other membranous material did occur.

Wolff and Jones (1971) explored a number of methods for initial tissue disruption and found that the Polytron homogenizer proved the most practical of the systems tested (see Section 2.1). The fractionation procedure is complex. The authors found the highest relative specific activity for TSH-sensitive adenylate cyclase in the "11K-top fraction" (purification factor ll-fold).t After further purification by discontinuous sucrose gradient centrifugation, the high-

*In Graves' disease the thyroid gland seems to be stimulated by an immunoglobulin called "Long-Acting Thyroid Stimulator" (LATS) or "Thyroid-stimulating autoantibody" (Tsaab). Homogenates of normal thyroid cancel the biological activity of LA TS presumably by binding the circulating LATS molecules to the injected plasma membrane preparations. The antigen responsible for that binding is designated as "LATS inhibitor."

tThis fraction corresponds to the pale upper layer of a pellet obtained after sedimentation at 11,000 rpm (14,000g) for 10 min.


est relative specific activity is found in the 37% sucrose boundary fraction (purification factor: 80- to ISO-fold). Other plasma membrane markers are also purified during this procedure. Wolff and Jones (1971) report that electron microscopy still shows contamination of the 37% sucrose boundary fraction with mitochondrial, endoplasmic reticular, and lysosomal fragments. The high purification factors reported for this 37% sucrose boundary fraction are rather unusual compared to the consistently lower figures reported by others (Yamashita and Field, 1970; Amir et al .. 1973). Moreover, one wonders if figures of this order of magnitude are compatible with the degree of contamination shown by electron microscopy or if, using this method, one is not simply purifying enzymes instead of membranes.

In our laboratory (De Wolf et al., 1978e), and using the same method, less plasma membrane marker activity was recovered in the 11K-top fraction. We also noted considerable contamination. The highest relative specific activity for the plasma membrane markers (S'-nucleotidase and alkaline phosphatase) was found at the 37%-40% sucrose interface. However, an increase in relative specific activity for the endoplasmic reticulum marker glucose-6-phosphatase was also found at that interface, but no NADPH-cytochrome c reductase. However, the relative specific activity of this enzyme in smooth endoplasmic reticulum is substantially lower than in rough endoplasmic reticulum (Hilderson et al .. 1975). Therefore, one cannot exclude the possibility that the 37% boundary fraction, as well as the 11K-top fraction of Wolff and Jones, do still contain endoplasmic reticulum membranes. The high rise of TSH-sensitive adenylate cyclase and of the other "stimulated" plasma membrane enzyme markers could be caused by a more efficient activation in purified fractions (Yamashita and Field, 1970).

Macchia and Meldolesi (1974), starting with a pellet sedimented after I,SOOg for 10 min also used a discontinuous gradient-centrifugation procedure yielding three bands and a sediment. The fraction accumulating at the 37% sucrose interphase contains TSH-sensitive adenyl ate cyclase with the highest relative specific activity (22) and the highest capacity for [125I]_TSH binding. However, no recoveries were reported that would allow conclusions concerning activation or inhibition phenomena.

Mehdi and Nussey (197S) made a 1000g supernatant up to 1.31 M with respect to sucrose, and layered it over the homogenization buffer containing 2.0 M sucrose. The material unable to enter the 2 M sucrose amounted to 90% of the adenyl ate cyclase activity of the 1 OOOg supernatant. However, no further indications were given for the purity of these membrane fractions.

Friedman et al. (1977) prepared a crude plasma membrane fraction from bovine thyroid starting from a pellet sedimented by 2S00g for 10 min. This fraction gave an enrichment by a factor 4 over the whole homogenate when


plasma membrane marker enzymes (5'-nucleotidase, ATPase, alkaline phosphatase) were assayed. No data on the purity of this fraction were reported. The authors were able to demonstrate that circulating thyroid hormones may regulate thyroid function by a "short loop" feedback mechanism (inhibition) effected prior to, as well as following, generation of thyroidal cAMP.

Ong et al. (1976) isolated plasma membranes according to the method of Wolff and Jones (1971) with some modification. The plasma membrane fractions were rich in adenylate cyclase as well as in (Na+, K+)ATPase and 5'nucleotidase activities. Basal adenylate cyclase was found in the four plasma membrane fractions, the highest activities being noted in the 45% and 30% sucrose interfaces. The other two enzymes showed enhanced activities in the 30% and 35% sucrose interfaces. Some contamination by mitochondria and endoplasmic reticular membranes was present in these fractions. No values on whole homogenate were reported, so that relative specific activities could not be computed. These results demonstrate that basal adenylate cyclase does not follow 5'-nucleotidase or ATPase, but that is is concentrated in a denser membrane fraction.

The distribution of plasma membranes on discontinuous or continuous sucrose gradients in a zonal rotor was analyzed in our laboratory (De Wolf et al., 1978e; Jansegers et al., 1979). Optimal isolation conditions were investigated including studies on the homogenizer, the homogenization medium, dilution of the homogenate, and centrifugation and g values. The best results are obtained after homogenization in water (adjusted to pH 7.4 by the addition of NaHC03) by means of a Waring Blendor (using 250 ml of a 1 : 5 homogenate blended for 15 sec at high speed). After discarding the pellet sedimented by 17 min at 3000g, a crude plasma membrane fraction is obtained by centrifuging at 39,000g for 15 min (the purification factor is 5, and the recovery approx. 20%). Isopycnic gradient centrifugation of a crude plasma membrane fraction results in 20-fold purification (De Wolf et aI., 1978e). However, contamination with endoplasmic reticular membranes (as measured by glucose-6-phosphatase) persists. The profiles of 5'-nucleotidase and alkaline phosphatase do not coincide, indicating that bovine thyroid plasma membranes are heterogenous. The phospholipid and cholesterol profiles after buoyant-density equilibration suggest the presence of four major membrane-containing regions. In all methods investigated the distributions of fluoride stimulated adenylate cyclase do not parallel the distribution of 5' -nucleotidase and alkaline phosphatase, but display maxima at higher densities (Jansegers et al., 1979).

Pochet et al. (1974) prepared horse thyroid plasma membranes by partition of a 600g pellet in an aqueous two phases system, yielding a plasma membrane enriched fraction at the interface. This fraction was only slightly con-


taminated with lysosomes and mitochondria as judged by the low activity of acid phosphatase and cytochrome c oxidase. It contained (N a +, K +)-A TPase activity and was enriched in adenylate cyclase. (The enzyme was stimulated by TSH and PGE1 and the purification was ninefold). In our laboratory, pellets obtained after 600g for 10 min, and 38,000g for 10 min, respectively, were subjected to similar distribution experiments. Plasma membrane markers were distributed over the two phases in a heterogenous way. The recovery of proteins at the interface was 0.32% and 4.32%, respectively, for the 600g X 10 min and 38,000g X 10 min pellet. Comparable relative specific activity values for 5'-nucleotidase were obtained at the interface for both kinds of pellets (approx. 5). Minor amounts of alkaline phosphatase were found at the interface for the 600g X 10 min pellet experiment. In contrast, a relative specific activity value of 5 was obtained for the 38,OOOg pellet. However, in those plasma membrane enriched fractions contamination still exists as acid phosphatase was purified fivefold, and glucose-6-phosphatase sixfold.

As a conclusion one can say that up to now it is not possible to prepare pure fractions that reflect the real plasma membrane situation in vivo. Vesicularization into small fragments due to the homogenization procedure is the major problem. Better results are obtained starting from dispersed thyroid cells of cell cultures (Tong et al .. 1962).

5. SUMMARY

In this review the subcellular localization of enzymes and constituents in thyroid is discussed. Conditions and results of differential pelleting and gradient centrifugation studies are described with special attention to the validity of the markets used (Table VI). Special approaches to the isolation and characterization of thyroid organelles and membranes are extensively reviewed (Table VII).

Subcellular fractionation of thyroid tissue has been shown to be an arduous task. Classic approaches for differential pelleting and gradient centrifugation, which have been proved successful for rat liver, are not always equally satisfactory for thyroid. The major problem is the toughness of the tissue requiring rather traumatizing homogenizing procedures. Nevertheless, the fractionation procedures did allow the subcellular localization of some enzymes and constituents to be established with a high degree of certainty. Furthermore, enriched subcellular fractions have been isolated which have been useful for biochemical studies concerning the specific function of this tissue.

Enz

yme

or c

onst

itue

nt

DN

A

RN

A-p

olym

eras

e I

RN

A-p

olym

eras

e II

RN

A-p

olym

eras

e II

I C

ytoc

hrom

e c

oxid

ase

Mon

oam

ine

oxid

ase

Suc

cina

te d

ehyd

roge

nase

C

ardi

olip

in

f3-G

lyce

roph

osph

atas

e

N-a

cety

l gl

ucos

amin

idas

e A

cid

f3-g

luco

sida

se

Aci

d ph

enyl

pho

spha

tase

f3-G

lucu

roni

dase

A

cid

lipas

e, a

cid,

and

ne

utra

l ph

osph

olip

ases

A

cid

RN

ase

Pho

spho

dies

tera

se

Tab

le V

I Su

bcel

lula

r L

ocal

izat

ion

of S

ome

Enz

ymes

and

Con

stit

uent

s in

Thy

roid

Tis

sue

Loc

aliz

atio

n V

alid

ity a

s m

arke

r

Alm

ost

com

plet

ely

in n

ucle

i G

ood

mar

ker

Pre

dom

inan

tly

in n

ucle

i N

ot v

alid

Nuc

lei

(bou

nd t

o ch

rom

atin

) M

oder

atel

y go

od

mar

ker

Pre

dom

inan

tly

in n

ucle

i N

ot v

alid

M

itoc

hond

ria

(inn

er m

embr

ane)

G

ood

mar

ker

Mit

ocho

ndri

a (o

uter

mem

bran

e)

Goo

d m

arke

r M

itoc

hond

ria

Goo

d m

arke

r M

itoc

hond

rial

mem

bran

es

Abs

olut

e m

arke

r

Lys

osom

es

Goo

d m

arke

r

Lys

osom

es

Goo

d m

arke

r L

ysos

omal

mem

bran

es

Goo

d m

arke

r L

ysos

omes

U

nrel

iabl

e

Lys

osom

es

Goo

d m

arke

r L

ysos

omes

G

ood

mar

kers

Lys

osom

es

Goo

d m

arke

r

Lys

osom

es

Unr

elia

ble

Com

men

t

Inta

ct n

ucle

i re

quir

ed.

Can

be

extr

acte

d du

ring

the

iso

latio

n of

nuc

lei.

Dou

ble

loca

liza

tion

not

exc

lude

d.

Mea

sure

men

ts o

nly

qual

itat

ive.

Int

act

nucl

ei r

equi

red.

A

ssay

s ex

pens

ive

for

rout

ine

anal

yses

. S

ee R

NA

-pol

ymer

ase

l.

Bal

ance

she

et le

ss t

han

100%

.

Mor

e ea

sily

sol

ubili

zed

than

cyt

ochr

ome

c ox

idas

e.

Can

be

used

for

det

ecti

on o

f co

ntam

inat

ion

with

m

itoc

hond

rial

mem

bran

es.

Dis

tinc

t fr

om a

cid

phen

ylph

osph

atas

e. D

oes

not

disp

lay

any

giuc

ose-

6-ph

osph

atas

e ac

tivi

ty.

Onl

y in

the

abs

ence

of

solu

ble

alka

line

f3-

gluc

osid

ase.

N

onsp

ecif

ic a

dsor

ptio

n on

to

othe

r m

embr

anes

. D

ispl

ays

gluc

ose-

6-ph

osph

atas

e ac

tivi

ty.

Ass

ay t

oo c

ompl

icat

ed f

or r

outi

ne a

naly

ses.

Cal

ibra

tion

cur

ves

requ

ired

(no

lin

eari

ty in

plo

ts o

f ac

tivi

ty v

ersu

s in

cuba

tion

tim

e or

enz

yme

conc

entr

atio

n (H

ilde

rson

et

ai.,

1970

).

Bal

ance

she

et n

ot 1

00%

ow

ing

to i

nhib

itio

n an

d ac

tiva

tion

ph

enom

ena

in t

he d

iffe

rent

sub

cell

ular

fra

ctio

ns

(Hil

ders

on e

t al

., 19

70).

N

Ut

N ?= :- ;, 5: .. ~ =

~ ft

Glu

cose

-6-p

hosp

hata

se

End

opla

smic

ret

icul

um m

embr

anes

G

ood

mar

ker

Onl

y su

itab

le in

the

abs

ence

of

the

bulk

of

acid

C

Il =

phen

ylph

osph

atas

e.

r::r

f")

NA

DP

H-c

ytoc

hrom

e c

End

opla

smic

ret

icul

um m

embr

anes

M

oder

atel

y go

od

Mor

e co

ncen

trat

ed in

rou

gh e

ndop

lasm

ic r

etic

ulum

!!.

e:: re

duct

ase

mar

ker

mem

bran

es.

Q> ..

Cyt

ochr

ome

b,

End

opla

smic

ret

icul

um m

embr

anes

R

elia

ble

mar

ker

Too

low

con

cent

rati

on f

or q

uant

itat

ive

anal

yses

and

1:1

:1

(pro

babl

y sm

ooth

) th

eref

ore

not

prac

tica

l fo

r ro

utin

e us

e.

S'

f") :r'

Lys

opho

spho

lipas

e ?

Not

val

id

Fur

ther

mor

e to

o di

ffic

ult

for

rout

ine

anal

yses

. .. e

Gua

iaco

lper

oxid

ase

Spec

iali

zed

dom

ain

of e

ndop

lasm

ic

Mod

erat

ely

good

P

resu

mab

ly o

nly

pres

ent

in s

ome

part

s of

rou

gh

~ ..

reti

culu

m m

embr

anes

m

arke

r en

dopl

asm

ic r

etic

ulu

m-i

t do

es n

ot r

efle

ct t

he w

hole

.. <0

<

dist

ribu

tion

of

thos

e m

embr

anes

. ~ ....

PPD

-per

oxid

ase

Spec

iali

zed

dom

ain

of e

ndop

lasm

ic

Mod

erat

ely

good

S

ee g

uaia

col p

erox

idas

e. P

roba

bly

dist

inct

fro

m

>-3

:r'

reti

culu

m m

embr

anes

m

arke

r gu

aiac

ol pe

roxi

dase

. <0

< .. ::, D

AB

-per

oxid

ase

? U

nrel

iabl

e N

onsp

ecif

ic c

osub

stra

te.

a.

Ade

nyla

te c

ycla

se

Pla

sma

mem

bran

es

Goo

d m

arke

r E

quil

ibra

tes

at h

ighe

r de

nsiti

es t

han

5'-n

ucle

otid

ase.

N

a+,K

+-s

tim

ulat

ed

Pla

sma

mem

bran

es

Goo

d m

arke

r D

iffi

cult

to d

iffe

rent

iate

tec

hnic

ally

fro

m m

itoch

ondr

ial

ouab

ain-

sens

itiv

e A

TPa

se.

AT

Pas

e 5'

-Nuc

leot

idas

e P

lasm

a m

embr

anes

G

ood

mar

ker

Cho

lest

erol

P

redo

min

antl

y in

pla

sma

Mod

erat

ely

good

N

o si

ngle

loc

atio

n, a

s it

is a

lso

pres

ent

to s

ome

degr

ee in

m

embr

anes

m

arke

r en

dopl

asm

ic r

etic

ulum

mem

bran

es.

Lip

id-b

ound

sia

lic a

cid

Pre

dom

inan

tly

in p

lasm

a M

oder

atel

y go

od

No

sing

le l

ocat

ion

(e.g

., Iy

soso

mes

). T

oo d

iffi

cult

for

mem

bran

es

mar

ker

rout

ine

anal

yses

. C

atal

ase

Pero

xiso

mes

G

ood

mar

ker

But

can

be

solu

biliz

ed d

urin

g ce

ntri

fuga

tion

. U

rate

oxi

dase

Pe

roxi

som

es

Rel

iabl

e B

ut a

ctiv

ity

too

low

for

qua

ntit

ativ

e an

alys

es.

Ade

nosi

ne d

eam

inas

e ?

? P

rese

nt in

the

S f

ract

ion

(cyt

osol

, fo

llicl

e lu

men

?).

Alk

alin

e iJ

-glu

cosi

dase

?

? S

ee a

deno

sine

dea

min

ase.

L

acta

te d

ehyd

roge

nase

C

ytos

ol

Goo

d m

arke

r iJ

-Gal

acto

sida

se

Lys

osom

es

Goo

d m

arke

r C

haba

ud e

t al

. (1

971)

. a

-Man

nosi

dase

L

ysos

omes

G

ood

mar

ker

Cha

baud

et a

f. (1

971)

. A

cid

prot

ease

and

L

ysos

omes

R

elia

ble

Peak

e et

al.

(196

6)

pept

idas

e H

erve

g et

al.

(196

6).

N

(Con

tinue

d)

UI

(H

Enz

yme

or c

onst

itue

nt

Gal

acto

sylt

rans

fera

se

N-A

cety

l gl

ucos

amin

yltr

ansf

eras

e M

anno

sylt

rans

fera

se

Sia

lylt

rans

fera

se

Tab

le V

I (C

ontin

ued)

Su

bcel

lula

r L

ocal

izat

ion

of S

ome

Enz

ymes

and

Con

stitu

ents

in T

hyro

id T

issu

e

Loc

aliz

atio

n

Mem

bran

es (

eith

er e

ndop

lasm

ic

reti

culu

m, G

olgi

app

arat

us, o

r pl

asm

a m

embr

anes

) M

embr

anes

Mem

bran

es

Mem

bran

es

Val

idity

as

mar

ker

Com

men

t

? Sp

iro

and

Spir

o (1

973)

C

haba

ud e

t af

. (1

974)

N

ot u

nam

bigu

ousl

y lo

caliz

ed.

? C

haba

ud e

t af

. (1

971)

S

ee g

alac

tosy

ltra

nsfe

rase

. ?

Spir

o an

d Sp

iro

(197

3)

Cha

baud

et

af.

(197

4)

See

gala

ctos

yltr

ansf

eras

e.

? S

piro

and

Spi

ro (

1973

) C

haba

ud e

t af

. (1

974)

S

ee g

alac

tosy

ltra

nsfe

rase

.

• Abb

revi

atio

ns:

PP

D.

p-ph

enyl

ened

iam

ine;

DA

B.

3.3'

-dia

min

oben

zidi

ne te

trac

hlor

ide.

~ .... ;: : ::c f =

~ It

Cel

l co

mpo

nent

Col

loid

dro

plet

s

Nuc

lei

Mit

ocho

ndri

a

Lys

osom

es

Gol

gi a

ppar

atus

Tab

le V

II

Isol

atio

n of

Thy

roid

Org

anel

les,

Sub

cellu

lar

Com

pone

nts,

and

Mem

bran

es'

Ref

eren

ces

Bal

asub

ram

ania

m e

t 01

. (1

965)

Hild

erso

n an

d D

ieri

ck

(197

4)

(I)

De

Wol

f et

01.

(I 9

78b)

; (2

) V

an D

esse

l et

al.

(197

8);

(3)

Van

den

H

ove-

Van

den

Bro

ucke

an

d D

e N

ayer

(I 9

76)

See

mit

ocho

ndri

a

Spi

ro a

nd S

piro

(19

73)

Met

hods

use

d

Dif

fere

ntia

l pe

lletin

g

Dif

fere

ntia

l pe

llet

ing,

dis

cont

inuo

us

sucr

ose

grad

ient

cen

trif

ugat

ion,

ra

te z

onal

cen

trif

ugat

ion

in a

n A

XII

zon

al r

otor

(I)

Dif

fere

ntia

l pe

lletin

g; (

2)

buoy

ant-

dens

ity

equi

libr

atio

n in

a

HS

-zon

al r

otor

; (3

) ce

ntri

fuga

tion

of

an

M +

L f

ract

ion

thro

ugh

a de

nse

pad

of 2

0% (

w Iv

) m

etri

zam

ide

See

mit

ocho

ndri

a

Dis

cont

inuo

us g

radi

ent

cent

rifu

gati

on

Com

men

t

Ele

ctro

n m

icro

grap

hs s

how

tha

t in

the

JS,

OO

Og

pelle

t co

lloid

dro

plet

s ar

e pr

esen

t. T

his

frac

tion

doe

s al

so c

onta

in o

ther

sub

cell

ular

co

mpo

nent

s.

Hig

hly

puri

fied

nuc

lear

fra

ctio

n. S

ingl

e po

pula

tion

(tet

rapl

oid)

. L

ipid

com

posi

tion

in

agre

emen

t w

ith l

iter

atur

e. N

o gl

ycol

ipid

s de

tect

able

. W

ater

-sol

uble

sub

stan

ces

and

enzy

mes

may

be

extr

acte

d du

ring

the

is

olat

ion

proc

edur

e.

Puri

fied

mit

ocho

ndri

al f

ract

ion

still

co

ntam

inat

ed b

y ot

her

subc

ellu

lar

com

pone

nts

(Iys

osom

es,

endo

plas

mic

re

ticu

lum

, pl

asm

a m

embr

anes

).

Mit

ocho

ndri

a w

ould

acc

ount

for

2.0

±

0.8%

of

tota

l th

yroi

d pr

otei

n.

Lys

osom

al f

ract

ion

still

con

tam

inat

ed b

y ot

her

subc

ellu

lar

com

pone

nts.

Lys

osom

es w

ould

ac

coun

t fo

r 3.

6 ±

2.2

% o

f to

tal

thyr

oid

prot

ein.

D

istr

ibut

ion

of s

ialy

l-an

d of

ga

lact

osyl

tran

sfer

ase

diff

eren

t fr

om t

hat

of

man

nosy

ltra

nsfe

rase

. B

alan

ce s

heet

not

es

ta b

lishe

d.

(Con

tinue

d)

rF.I = if i!.

;:- DO .. 1:1

:1 o· " if 3 5.. ~

Q ... -l =...., a 6:

N

(JJ

(JJ

Tab

le V

II (

Con

tinu

ed)

~

="

Isol

atio

n of

Thy

roid

Org

anel

les,

Sub

cellu

lar

Com

pone

nts,

and

Mem

bran

es·

Cel

l co

mpo

nent

R

efer

ence

s M

etho

ds u

sed

Com

men

t

Bou

chill

oux

et a

l. (1

970)

; D

isco

ntin

uous

gra

dien

t ce

ntri

fuga

tion

T

he 0

.4-1

.0S

M s

ucro

se i

nter

face

con

tain

s C

haba

ud e

t al.

(197

4)

mor

phol

ogic

ally

rec

ogni

zabl

e G

olgi

fr

agm

ents

. T

he d

istr

ibut

ion

of

glyc

osyi

tran

sfer

ases

dif

fers

fro

m t

hat

of

plas

ma

mem

bran

es.

Bal

ance

she

et n

ot

esta

blis

hed.

The

gly

cosy

ltran

sfer

ases

are

not

lo

caliz

ed w

ith c

erta

inty

. P

acus

zka

et a

l. (1

978)

D

isco

ntin

uous

gra

dien

t ce

ntri

fuga

tion

N

ot u

nam

bigu

ousl

y pr

oven

tha

t th

e is

olat

ed

frac

tion

corr

espo

nds

to G

olgi

app

arat

us.

Poly

som

es

De

Nay

er a

nd D

e V

issc

her

Dif

fere

ntia

l pe

lletin

g, g

radi

ent

The

dis

trib

utio

n th

roug

hout

a s

ucro

se g

radi

ent

(196

9);

Kon

do e

t al.,

ce

ntri

fuga

tion

is

sim

ilar

to t

hose

fro

m o

ther

mam

mal

ian

1968

tis

sues

. Po

lyri

boso

mes

are

abl

e to

syn

thes

ize

prot

eins

. L

ecoc

q et

al.

(I 9

71)

Dif

fere

ntia

l pe

lletin

g, g

radi

ent

I S-2

0% a

re m

onom

ers,

4%

dim

ers,

S%

ce

ntri

fuga

tion

tr

imer

s, 7

% t

etra

mer

s, 7

-8%

pen

tam

ers,

9%

he

xam

ers,

SO

-60%

hea

vy p

olys

omes

. C

heba

th e

t al

. (I

977a

,b);

In

dire

ct i

mm

unop

reci

pita

tion

T

hyro

glob

ulin

-spe

cifi

c po

lyso

mes

rep

rese

nt

Vas

sart

et a

l. (1

977)

; 30

-4S

% o

f to

tal

poly

som

es.

The

spe

cifi

c V

an V

oort

huiz

en e

t al

. th

yrog

lobu

lin m

RN

A is

lar

ge e

noug

h to

(I

978)

co

de f

or t

he w

hole

thy

rogl

obul

in p

eptid

e ;t

ch

ain.

...

Pla

sma

mem

bran

es

Yam

ashi

ta a

nd F

ield

Fl

otat

ion

of a

2S0

0-rp

m p

elle

t fr

om a

S

tudy

of

TS

H e

ffec

t on

ade

nyla

te c

ycla

se.

; (1

970)

N

aHC

O,

hom

ogen

ate

thro

ugh

a E

lect

ron

mic

rosc

opy

show

s th

at t

he i

sola

ted

5:

disc

ontin

uous

suc

rose

gra

dien

t pl

asm

a m

embr

ane

frac

tion

s do

con

sist

of

~ ... '"

(Nev

ille

met

hod)

ve

sicu

lar

mat

eria

l an

d of

pla

sma

mem

bran

e 0 =

sh

eets

with

(m

inim

al)

cont

amin

atio

n.

~

Enr

ichm

ent:

10-

to 2

0-fo

ld.

;:,

:--

Am

ir e

t al

. (1

973)

; A

zuki

zaw

aet

al.

(197

7)

Roq

ues

et a

l. (1

975)

Ver

rier

et

al.

(197

6)

Suzu

ki e

t al

. (1

977)

Nit

riya

nant

and

Dun

lap

(197

8)

Sta

nbur

y et

al.

(196

9)

Wol

ff a

nd J

ones

(19

71)

Nev

ille

met

hod

Nev

ille

met

hod

Try

psin

izat

ion

of p

orci

ne t

hyro

id

glan

ds

Slig

ht m

odif

icat

ion

of th

e m

etho

d of

Y

amas

hita

and

Fie

ld (

I 970

)

Met

hod

of Y

amas

hita

and

Fie

ld

( 197

0)

Nit

roge

n m

icro

cavi

tati

on o

f th

yroi

d tis

sue,

cen

trif

ugat

ion

in l

ow i

onic

st

reng

th b

uffe

rs, c

entr

ifug

atio

n of

a

104,

000g

pel

let

laye

red

over

fic

oll

Dis

rupt

ion

in a

Pol

ytro

n ho

mog

eniz

er, c

ompl

ex

frac

tion

atio

n pr

oced

ure,

di

scon

tinuo

us g

radi

ent

cent

rifu

gati

on

Bin

ding

of

TS

H to

iso

late

d bo

vine

thy

roid

pl

asm

a m

embr

anes

. B

alan

ce s

heet

not

re

port

ed.

Ele

ctro

n m

icro

grap

hs s

how

lar

ge

vesi

cula

r st

ruct

ures

. V

ery

prob

ably

co

ntam

inat

ed.

Isol

ated

fra

ctio

n st

ill c

onta

ins

othe

r su

bcel

lula

r co

mpo

nent

s, a

pro

tein

kin

ase

bein

g en

rich

ed in

thi

s is

olat

ed f

ract

ion.

A

naly

sis

of p

hosp

holip

id a

nd p

rote

in c

onte

nt

of t

he e

nric

hed

plas

ma

mem

bran

e fr

actio

n.

Isol

atio

n of

two

subf

ract

ions

(H

and

L).

R

ecov

ery

of p

rote

in a

nd o

f m

arke

r en

zym

es

in t

he p

lasm

a m

embr

anes

is v

ery

low

. E

lect

ron

mic

rosc

opy

show

s th

at to

tal

plas

ma

mem

bran

es a

nd H

-mem

bran

es c

onta

in

junc

tion

al c

ompl

exes

, lo

ng m

embr

ane

shee

ts,

and

vesi

cles

. T

he p

rese

nce

of L

AT

S i

nhib

itor

in p

lasm

a m

embr

ane

frac

tions

. B

alan

ce s

heet

not

re

port

ed.

No

data

allo

win

g th

e co

mpu

tati

on

of th

e re

lativ

e sp

ecif

ic a

ctiv

ities

men

tione

d.

Sub

stan

tial

con

tam

inat

ion

by o

ther

m

embr

anou

s m

ater

ial.

Ver

y hi

gh e

nric

hmen

t of

plas

ma

mem

bran

e m

arke

rs (

80-

to I

SO-f

old)

. A

s fa

r as

we

are

awar

e th

is r

esul

t ha

s ne

ver

been

con

firm

ed

by o

ther

s. E

lect

ron

mic

rosc

opy

reve

als

cont

amin

atio

n. G

luco

se-6

-pho

spha

tase

is

also

enr

iche

d, b

ut n

ot t

rue

of N

AD

PH

cy

toch

rom

e c

redu

ctas

e. C

onta

min

atio

n w

ith e

ndop

lasm

ic r

etic

ulum

ver

y lik

ely.

(Con

tinue

d)

\fJ = et' & =

&i .. til o· " =- a 1;;' ::t

'< sa. ..., = '< a Q:

N

<.II

-.I

Tab

le V

II (

Con

tinu

ed)

Isol

atio

n of

Thy

roid

Org

anel

les,

Sub

cellu

lar

Com

pone

nts,

and

Mem

bran

es'

Cel

l co

mpo

nent

R

efer

ence

s

Mac

chia

and

Mel

dole

si

(197

4)

Meh

di a

nd N

usse

y (1

975)

Fri

edm

an e

t al

. (1

977)

Ong

et

al.

(197

6)

De

Wol

f et

al.

(197

8e);

Ja

nseg

ers

et a

l. (1

979)

Poch

et e

t al

. (1

974)

'Abb

revi

atio

n: L

A T

S:

Lon

g-A

ctin

g T

hyro

id S

tim

ulat

or I

nhib

itor

.

Met

hods

use

d

Dis

cont

inuo

us g

radi

ent

cent

rifu

gati

on

Dis

cont

inuo

us g

radi

ent

cent

rifu

gati

on

Dis

cont

inuo

us g

radi

ent

cent

rifu

gati

on

Met

hod

of W

olff

and

Jon

es (

1971

)

Met

hod

of Y

amas

hita

and

Fie

ld a

nd

of W

olff

and

Jon

es,

disc

ontin

uous

gr

adie

nt c

entr

ifug

atio

n in

zon

al

roto

rs

Par

titi

on o

f a

600g

pel

let

in a

n aq

ueou

s tw

o ph

ases

sys

tem

Com

men

t

Stu

dy o

f T

SH

eff

ect.

Bal

ance

she

et n

ot

esta

blis

hed.

S

tudi

es o

n L

A T

S.

No

indi

cati

ons

abou

t th

e pu

rity

of

thei

r m

embr

ane

frac

tions

. In

hibi

tion

of th

yroi

d ad

enyl

ate

cycl

ase

by

thyr

oid

horm

one.

No

data

on

the

puri

ty o

f th

eir

isol

ated

fra

ctio

ns r

epor

ted.

N

o da

ta a

llow

ing

the

calc

ulat

ion

of re

lati

ve

spec

ific

act

iviti

es.

The

res

ults

sho

w t

hat

basa

l ad

enyl

ate

cycl

ase

does

not

fol

low

5'

nucl

eoti

dase

or

AT

Pas

e, b

ut t

hat

it is

co

ncen

trat

ed in

a d

ense

r m

embr

ane

frac

tion.

In

all

frac

tion

s ob

tain

ed c

onta

min

atio

n w

ith

endo

plas

mic

ret

icul

um p

ersi

sts.

Bov

ine

thyr

oid

plas

ma

mem

bran

es s

eem

to

be

hete

roge

nous

. A

deny

late

cyc

lase

doe

s no

t pa

rall

el t

he d

istr

ibut

ion

of 5

' -nuc

leot

idas

e or

al

kali

ne p

hosp

hata

se,

but

disp

lays

max

ima

at h

ighe

r de

nsiti

es.

Eff

ect

of T

SH

and

PO

E!

on t

hyro

id a

deny

late

cy

clas

e. P

lasm

a m

embr

ane

frac

tion

stil

l co

ntam

inat

ed w

ith o

ther

sub

cell

ular

co

mpo

nent

s.

N

'JI

oc ;: ~ :=; i5: ~ ~ =

~

I:>

:--


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Boddin, M., Lagrou, A., Hilderson, H. J., and Dierick, W., 1976, Studies on the specificity of bovine thyroid ribonuclease type IVb, Abstr. Cornrnun. 10th IUB Congress, Hamburg, 176, Abstr. No. 04-3-323.

Bouchilloux, S., Chabaud, 0., Michel-Bechet, M., Ferrand, M., and Athouel-Haon, A. M., 1970, Differential localization in thyroid microsomal subfractions of a mannosyltransferase, two N-acetylglucosaminyltransferases and a galactosyltransferase, Biochern. Biophys. Res. Cornrnun.40:314-320.

Chabaud, 0., Bouchilloux, S., and Ferrand, M., 1971, Caracterisation, isolement et proprietes de glycosidases thyroldiennes: i3-Galactosidase, 13-N-acetylglucosaminidase et i3-mannosidase, Biochirn. Biophys. Acta 227: 154-170.

Chabaud, 0., Bouchilloux, S., Ronin, c., and Ferrand, M., 1974, Localization in a Golgi-rich thyroid fraction of sialyl-, galactosyl- and N-acetylglucosaminyltransferases, Biochirnie 56:119-130.

Chebath, J., Chabaud, 0., Berge-Lefranc, J. L., Cartouzou, G., and Lissitzky, S., 1977a, Molecular weight of the thyroglobulin messenger RNA of sheep thyroid gland, Biochern. Biophys. Res. Cornrnun. 79:267-273.

Chebath, J., Chabaud, 0., Becarevic, A., Cartouzou, G., and Lissitzky, S., 1977b, Thyroglobulin messenger ribonucleic acid translation in vitro, Eur. J. Biochern. 77:243-252.

Colilla, W., Jorgenson, R. A., and Nordlie, R. c., 1975, Mammalian carbamyl phosphate: glucose phosphotransferase and glucose-6-phosphate phosphohydrolase: Extended tissue distribution, Biochirn. Biophys. Acta 337:117-125.

Croft, C. J., and Pitt-Rivers, R., 1970, Radioautographic studies on the initial site of formation of protein bound iodine in the rat thyroid gland, Biochern. J. 118:311-314.


Davies, T. F., McLachlan, S. M., Povey, P. M., Smith, B. R., and Hall, R., 1977, The influence of propranolol on the thyrotropin receptor, Endocrinology 100:974-979.

de Duve, C., 1963, The lysosome, Sci. Am. 208:64-72. de Duve, c., 1971, Tissue fractionation. Past and present, J. Cell Bioi. 50:20D-55D. De Groot, L., 1977, Thyroid hormone biosynthesis, in: Endocrinology (V. H. T. James, ed.),

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De Groot, L. J., and Carvalho, E., 1960, Iodide binding in thyroid cellular fractions, J. Bioi. Chem. 235: 1390-1397.

De Nayer, P., and De Visscher, M., 1969, Thyroglobulin synthesis in a thyroid polyribosomal cell-free system, Biochem. Biophys. Res. Commun. 36:639-646.

De Wolf, M., Lagrou, A., Hilderson, H. J., and Dierick, W., 1976, Phospholipases AI and A2 in bovine thyroid, Biochim. Biophys. Acta 424:183-194.

De Wolf, M. J. S., Hilderson, H. J. J., and Dierick, W. S. H., 1977, A note on zonal-rotor centrifugation of bovine thyroid membranes, and the lipolytic enzyme profiles, in: Membranous Elements and Movement of Molecules, Vol. 6, Methodological Surveys in Biochemistry (E. Reid, ed.), Ellis Horwood Ltd., Chichester.

De Wolf, M., Lagrou, A., Hilderson, H. J., and Dierick, W., 1978a, Lipolytic enzymes in bovine thyroid tissue. I. Subcellular localization, purification and characterization of acid phospholipase AI, Arch. Int. Physiol. Biochim. 86:1055-1075.

De Wolf, M., Lagrou, A. R., Hilderson, H. J., Van Dessel, G. A. F., and Dierick, W. S. H., 1978b, Subcellular structure of bovine thyroid gland. The localization of the peroxidase activity in bovine thyroid, Biochem. J. 174:939-949.

De Wolf, M., Lagrou, A., Hilderson, H. J., and Dierick, W., 1978c, Acid triacylglycerollipase from bovine thyroid, Lipids 13:598-604.

De Wolf, M., Hilderson, H. J., Lagrou, A., and Dierick, W., 1978d, Lipolytic enzymes in bovine thyroid tissue. II. Hydrolysis of [3R, 14C] phosphatidylethanolamine by neutral and alkaline phospholipase A activities, Arch. Int. Physiol. Biochim. 86:1077-1089.

De Wolf, M. J. S., Hilderson, H. J. J., Lagrou, A. R., and Dierick, W. S. H., 1978e, Subcellular structure of bovine thyroid gland. VII. A study on the distribution of bovine thyroid plasma membranes by density gradient centrifugation in zonal rotors, Arch. Int. Physiol. Biochim. 86:37-52.

De Wolf, M., Hilderson, H. J., Lagrou, A., and Dierick, W., 1979, Lipolytic enzymes in bovine thyroid tissue. III. Lysophospholipase activity, Arch. Int. Physiol. Biochim. 87:1-18.

Dierick, W., and Hilderson, H. J., 1967, Subcellular structure of bovine thyroid. I. /3-Glucuronidase: A marker enzyme for lysosomal particles, Arch. Int. Physiol. Biochim. 75: 1-11.

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Evans, W. R., 1978, Preparation and characterization of mammalian plasma membranes, in: Laboratory Techniques in Biochemistry and Molecular Biology (T. S. Work and E. Work, eds.), North-Holland, Amsterdam.

Fischer, A. G., Schulz, A. R., and Oliner, L., 1968, Distribution of monoamine oxidase in the thyroid gland, Endocrinology 82:1098-1102.

Friedman, Y., Lang, M., and Burke, G., 1977, Inhibition of thyroid adenylate cyclase by thyroid hormone: A possible locus for the "short-loop" negative feedback phenomenon, Endocrinology 101:858-868.

Glaumann, H., Persson, H., Bergstrand, A., and Ericsson, J. L. E., 1974, Characterization of liver microsomal sub-fractions and the Golgi apparatus with respect to chemical composition and intracellular transport of macromolecules, in: Methodological Developments in Biochemistry (E. Reid, ed.), Vol. 4, pp. 237-246, Longman, London.


Haye, 8., Champion, S., and Jacquemin, C, 1974, Existence of two pools of prostaglandins during stimulation of the thyroid by TSH, FEBS Lett. 41:89-93.

Herveg, J. P., Beckers, C., and De Visscher, M., 1966, Lysosomal hydrolases in calf thyroid, Biochem. J. 100:540-548.

Hilderson, H. J., and Dierick, W., 1973, Absence of respiratory activity in bovine thyroid nuclei, Arch. Int. Physiol. Biochim. 81:921-923.

Hilderson, H. J., and Dierick, W., 1974, Subcellular structure of bovine gland. VI. A study of bovine thyroid nuclei by density gradient centrifugation in a zonal A-XII rotor, Arch. Int. Physiol. Biochim. 82:75-83.

Hilderson, H. J., Dierick, W., and Stockx, J., 1965, Lysosomen in de schildklier, Arch. Int. Physiol. Biochim. 73:871.

Hilderson, H. J., Stockx, J., and Dierick, W., 1970, Subcellular structure of bovine thyroid tissue. II. Localization of RNA, DNA, acid hydrolases and adenosine deaminase, Arch. Int. Physiol. Biochim. 78:509-518.

Hilderson, H. J., Stockx, J., and Dierick, W., 1971a, Subcellular structure of bovine thyroid. III. The significance of sedimentable free enzyme activities, Arch. Int. Physiol. Biochim. 79:337-344.

Hilderson, H. J., Stockx, J., and Dierick, W., 1971 b, Subcellular structure of bovine thyroid. IV. The presence of two phosphatases. The influence of {3-glycerophosphate on the stability of lysosomes, Arch. Int. Physiol. Biochim. 79:345-350.

Hilderson, H. J., Stockx, J., and Dierick, W., 1971c, Subcellular structure of bovine thyroid. V. Gradient centrifugation, Arch. Int. Physiol. Biochim. 79:523-534.

Hilderson, H. J., Lagrou, A., and Dierick, W., 1974, The nuclear lipids of bovine hypertrophic thyroid, Biochim. Biophys. Acta 337:385-389.

Hilderson, H. J. J., De Wolf, M. J. S., Lagrou, A. R., and Dierick, W. S. H., 1975, Subcellular structure of bovine thyroid gland. A study on bovine thyroid membranes by buoyant-densitygradient centrifugation in a B-XIV zonal rotor, Biochem. J. 152:601-607.

Hilderson, H. J., De Wolf, M., Lagrou, A., and Dierick, W., 1976, The hydrolysis of glucose-6-phosphate in bovine thyroid gland, Abstr. Commun. 10th IUB Congress, Hamburg 662, Abstr. No. 16-5-052.

Hinton, R. H., Norris, K. A., and Reid, E., 1970, Isolation of hepatic membrane fragments, in: Separations with Zonal Rotors (E. Reid, ed.), pp. S2-1-S2-16, University of Surrey, Guildford.

Hosoya, T., Matsukawa, S., and Nagai, Y., 1971, Localization of peroxidase and other microsomal enzymes in thyroid cells, Biochemistry 10:3086-3093.

Hosoya, T., Matsukawa, S., and Nagai, Y., 1973, Localization of thyroid peroxidase, in: Endocrinology (R. O. Scow, ed.), Proceedings of the 4th International Congress of Endocrinology, Washington, D.C., June 18-24, 1972, Excerpta Medica, Amsterdam.

Ichikawa, Y., Saito, E., Abe, Y., Homme, M., Maraki, T., and Ito, K., 1976, Presence of TSH receptor in thyroid neoplasms, J. Clin. Endocrinol. Metab. 42:395-398.

Jablonski, P., and McQuillan, M. T., 1967, The distribution of proteolytic enzymes in the thyroid gland, Biochim. Biophys. Acta 132:454-471.

Jansegers, L., Hilderson, H. J., Lagrou, A., and Dierick, W., 1978, Vergelijkende studie over het bereiden van plasmamembranen uit runderschildklier, Arch. Int. Physiol. Biochim. 87:187-188.

Johnston, I. R., Mathias, A. P., Pennington, F., and Ridge, D., 1968, The fractionation of nuclei from mammalian cells by zonal centrifugation, Biochem. J. 109:127-135.

Kellas, 8. L., Austoker, J. L., Beebee, T. J. C, and Butterworth, P. H. W., 1977, Forms AI and All DNA-dependent RNA polymerases as components of two defined pools of polymerase activity in mammalian cells, Eur. J. Biochem. 72:583-594.


Kogl, F., and Van Deenen, L. L. M., 1961, Metabolism and function of phosphatides. Investigation on the action of thyrotrophic hormone on thyro-phosphatides, Acta Endocrinol. 36:9-21.

Kohn, L. D., 1978, Relationships in the structure and function of receptors for glycoprotein hormones, bacterial toxins and interferon, in: Receptors and Recognition (P. Cuatrecasas and M. F. Greaves, eds.), Series A, Vol. 5, pp. 135-212, Chapman and Hall, London.

Kondo, Y., De Nayer, P., Labaw, L. W., Robbins, J., and Rall, J. E., 1968, Properties of bovine thyroid ribosomes and polyribosomes, Endocrinology 83: 1117-1132.

Lagrou, A., Hilderson, H. J., De Wolf, M., and Dierick, W., 1974a, Bovine thyroid gangliosides: Distribution pattern after differential pelleting, Arch. Int. Physiol. Biochim. 82:733-736.

Lagrou, A., Dierick, W., Christophe, A., and Verdonk, J., 1974b, Lipid composition of bovine normal and hypertrophic thyroid, Lipids 9:870-875.

Lecocq, R. E., Cantraine, F., Keyhani, E., Claude, A., Delcroix, C., and Dumont, J. E., 1971, Quantitative evaluation of polysomes and ribosomes by density gradient centrifugation and electron microscopy, Anal. Biochem. 43:71-79.

Lee, T., Swartzendruber, D. C., and Snyder, F., 1969, Zonal centrifugation of microsomes from rat liver: resolution of rough- and smooth-surfaced membranes, Biochem. Biophys. Res. Commun. 36:748-755.

Leighton, F., Poole, B., Beaufay, H., Baudhuin, P., Coffey, J. W., Fowler, S., and de Duve, C., 1968, The large scale separation of pcroxisomes, mitochondria and lysosomes from the livers of rats injected with Triton WR-1339. Improved isolation procedures, automated analysis, biochemical and morphological properties of fractions, J. Cell Bioi. 37:482-513.

Lin, y.-c., Rose, K. M., and Jacob, S. T., 1976, Evidence for the nuclear origin of RNA polymerases identified in the cytosol: Release of enzymes from the nuclei isolated in isotonic sucrose, Biochem. Biophys. Res. Commun. 72:114-120.

Lygre, D. G., and Nordlie, R. c., 1969, Rabbit intestinal glucose-6-phosphate phosphohydrolase and inorganic pyrophosphate-glucose phosphotransferase: Inhibition by phlorizin, Biochim. Biophys. Acta 185:360-366.

Macchia, V., and Meldolesi, M. F., 1974, Cell membrane of thyroid and its altered responsiveness to the hormone TSH, in: Advances in Cytopharmacology (B. Ceccarelli, F. Clementi, and J. Meldolesi, eds.), Vol. 2, Raven Press, New York.

Matsukawa, S., and Hosoya, T., 1979a, Process of iodination of thyroglobulin and its maturation. I. Properties and distribution of thyroglobulin labeled with radioiodine in pig thyroid slices, J. Biochem. 85:1009-1021.

Matsukawa, S., and Hosoya, T., I 979b, Process of iodination of thyroglobulin and its maturation. 2. Properties and distribution of thyroglobulin labeled in vitro or in vivo with radioiodine, 3H-tyrosine or 3H-galactose in rat thyroid glands, J. Biochem. 86:199-212.

Matsuzaki, S., Pochet, R., and Schell-Frederick, E., 1973, A comparison of the subcellular distribution of 5'-nucleotidase, (Na + -K+)-A TPase and adenylcyclase in beef thyroid gland, Biochim. Biophys. Acta 313:329-337.

Mehdi, S. Q., and Nussey, S. S., 1975, A radio-ligand receptor assay for the long-acting thyroid stimulator. Inhibition by the long-acting thyroid stimulator of the binding of radio-iodinated thyroid-stimulating hormone to human thyroid membranes, Biochem. J. 145:105-111.

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Yamashita, K., and Field, J. B., 1970, Preparation of thyroid plasma membranes containing a TSH-responsive adenylcyclase, Biochem. Biophys. Res. Commun. 40: 171-178.

Chapter 6

The Molecular Organization of NADH Dehydrogenase

C. I. Ragan Department of Biochemistry University of Southampton Southampton, HANTS, S09 3TU, U.K.

1. INTRODUCTION

1.1. The Purpose of This Chapter

Interest in the structure of the enzymes of the mitochondrial respiratory chain is a fairly recent development. Previously work has concentrated on the nature of the chemical groups that undergo oxidoreduction, their sequence of operation, and their thermodynamic and kinetic properties. Although this aspect is by no means close to completion, research on the protein structure around these functional groups is becoming increasingly common. I think it has taken a while for us to realize that the respiratory chain is composed of enzymes rather than just prosthetic groups. Peter Mitchell has shown us that we cannot describe the process of oxidative phosphorylation without considering the organization of the constituent enzymes in the membrane. Second, we have had to wait for the introduction of powerful analytical techniques, such as sodium dodecyl sulfate-polyacrylamide gel electrophoresis, before attempting to describe the protein composition and structure of these amazingly complex enzymes.

This chapter is concerned with NADH dehydrogenase, the most complicated of the respiratory chain enzymes. The catalytic properties of this enzyme, both in the membrane and in isolation, have been reviewed many times in recent years (e.g., Singer and Gutman, 1970; Hatefi and Stiggall, 1976), and I shall try to avoid this aspect as much as possible, concentrating instead on the structure of the enzyme and, where possible, on its relationship to function.

267

268 C.I. Ragan

Sections 2 and 3 concern, the protein and Section 4 the phospholipid components of the enzyme. The fifth major section concerns the organization of the enzyme in the inner mitochondrial membrane.

1.2. Definitions and Terminology

The terminology of respiratory chain enzymes is frequently cumbersome. Only cytochrome oxidase has a name that is short, descriptive, and most importantly, used for the enzyme either in the membrane or quite generally for the product of any of the published purification procedures. No such name exists for the enzyme of NADH oxidation. NADH dehydrogenase is the term used for the enzyme in the membrane, but isolated forms are usually named according to their specific properties or specific isolation procedures, e.g., complex I, type I NADH dehydrogenase, and soluble NADH-ubiquinone reductase. I would like to propose that NADH dehydrogenase be used as a generic term for the enzyme in the membrane and in isolation. This presupposes that we know what properties the isolated enzyme should have. Whereas this was a problem some years ago, I do not think that anyone would disagree that the ideal isolated NADH dehydrogenase should retain the capacity for rotenonesensitive reduction of ubiquinone analogues by NADH and should be free from cytochromes and other dehydrogenase activities. The low-molecular-weight fragment of NADH dehydrogenase, which catalyzes the oxidation of NADH by artificial electron acceptors as well as ubiquinone analogues, has long been recognized as having been modified in its properties by isolation. This fragment is generally called type II NADH dehydrogenase, but I would prefer to avoid any possible confusion by abandoning this term and using the term flavoprotein fragment instead. This term is descriptive of both its prosthetic group and its derivation.

The purified form of NADH dehydrogenase that was used for most of the work described in this chapter is the lipoprotein isolated first by Hatefi et al. (1962a) and generally known as complex I or NADH-ubiquinone oxidoreductase. This preparation retains the ability to reduce ubiquinone analogues in a rotenone-sensitive manner and can be recombined with other respiratory enzymes to reconstitute NADH oxidase activity (Hatefi et al., 1962b). Furthermore, when reconstituted into phospholipid vesicles, the enzyme catalyzes oxidoreduction-linked proton translocation (Ragan and Hinkle, 1975). It therefore closely resembles NADH dehydrogenase in the membrane in its most important properties. Unfortunately it cannot be isolated in a completely pure form, and it is always contaminated with small amounts of cytochromes band CI' Soluble lipid-free NADH dehydrogenase preparations can be obtained without any heme contamination (Ringler et al., 1963) and, as has been pointed out by Singer anq Edmondson (1978), these may be more suitable for certain types of research. However, the inability to reduce ubiquinone ana-

NADH Dehydrogenase Structure 269

logues remains a serious drawback. This lack is generally ascribed to the absence of phospholipid, although a lipid-free preparation that retains this activity with altered kinetic characteristics has been described (Baugh and King, 1972). I believe that phospholipids should be regarded as constituents of NADH dehydrogenase, and there is evidence that their absence causes conformational changes in the protein. As the work to be described is on the structure of NADH dehydrogenase, clearly complex I remains the preparation of choice.

All the purified NADH dehydrogenases contain FMN as the sole flavin constituent and comparatively large amounts of nonheme iron and acid-labile sulfide (e.g., Ragan, 1976c). The precise stoichiometry of iron to flavin is not certain, although the enzyme appears to contain five distinct iron-sulfur paramagnetic centers (Orme-Johnson et al .. 1974; Albracht et al .. 1977).

1.3. The Functional Unit

Preparations of NADH dehydrogenase purified by different procedures have been reported to contain 1.1-1.5 nmoles FMN / mg protein (e.g., Ragan, 1976c). The variation in FMN content may be caused in part by the ease with which this prosthetic group can be removed, as it is not covalently attached to the protein. For example, ammonium sulfate precipitation of complex I in the presence of cholate causes a considerable reduction in FMN content (Ragan and Racker, 1973). On the basis of FMN content, the minimum protein molecular mass would therefore be in the range of 600,000-900,000 daltons. Direct measurements of the molecular mass are difficult to make because both the water-soluble lipid-free NADH dehydrogenase (Cremona and Kearney, 1964) and the detergent-soluble lipoprotein enzyme are heterogeneous by ultracentrifugation. Dooijewaard et al. (1 978a) analyzed preparations of complex I by ultracentrifugation and found two main components. One of them had a S ;o,w value of 42 S. The partial specific volume was 0.83 cm3/g, giving a molecular weight of 2.8 X 106, for a spherical molecule. The minimum molecular weight based on FMN content was 1.1 X 106 (rather higher than usually found), from which they concluded that the 42 S component contained two FMN molecules per molecule of enzyme, with phospholipids and residual detergent making the molecular weight up to 2.8 X 106• This conclusion only concerns the isolated enzyme and does not necessarily imply that in the mitochondrial membrane a molecule of NADH dehydrogenase contains two molecules of FMN. However, there are a number of other pieces of evidence which suggest that NADH dehydrogenase may be oligomeric.

The epr spectrum of purified complex I indicates the presence of five paramagnetic species-centers la, 1 b, 2, 3, and 4 (Orme-Johnson et al .. 1974; Albracht et al .. 1977). Orme-Johnson et al. (1974) quantitated the signals by double integration and found that each of the centers 1-4 was present in

270 C. I. Ragan

approximately the same concentration as FMN. As this work was done before center 1 was resolved into two components-l a and 1 b-it can be concluded that each of these centers is present at only one-half the concentration of FMN. Albracht et al. (1977) attempted to sort out the overlapping spectra of complex I by computer simulation and came to different conclusions from Orme-Johnson et al. (1974) about the assignment of certain resonance peaks to individual centers. The details are outside the scope of this chapter, but Albracht et al. (1977) concluded that centers 1 a and 1 b were each present at only one-quarter the concentration of FMN. That the two centers la and 1b were not preparation artifacts was shown by their presence in submitochondrial particles. Moreover, the combined concentration of centers 1a and 1 b was only one-half that of center 2, exactly as found with purified complex I (Albracht et al., 1979).

Further evidence for an oligomeric structure for NADH dehydrogenase came from analyses of the steady state kinetics of complex I by Dooijewaard and Slater (1976a) and Ragan (1978). The former found evidence for at least two interacting NADH binding sites per molecule of enzyme, whereas the latter found that reduction of ubiquinone-l by complex I did not follow MichaelisMenten kinetics. In addition, isolated complex I binds approximately one molecule of the inhibitor piericidin per molecule of FMN at a high-affinity binding site (Gutman et al., 1970). Although there is no evidence for binding sites of differing affinities, the relationship between inhibition of NADH-ubiquinone oxidoreductase activity and concentration of added rotenone, which acts at the same site as piericidin, is not linear. For example, 50% inhibition was found with as little at 0.1 mole of rotenone per mole FMN (Ragan and Heron, 1978). A similar relationship between inhibition and inhibitor binding was shown by the covalent modifier, diphenyleneiodonium (Ragan and Bloxham, 1977; see also Section 3.7.3). This type of behavior is normally attributed to negative cooperativity between the monomeric units of an oligomeric enzyme.

The results described above suggest the following possibilities for NADH dehydrogenase. First, NADH dehydrogenase could be a tetramer with each monomer containing one molecule of FMN and binding sites for NADH, piericidin (or rotenone), and diphenyleneiodonium. Because several of the dehydrogenase polypeptides are present in similar concentrations to FMN, it is most likely that each monomer would have identical polypeptide composition. The four units would be linked by the iron-sulfur centers 1 a and 1 b, perhaps by each monomer providing two cysteinyl residues for bonding to the iron atoms. Thus there would be only one center la or 1 b per tetramer. This proposal is not consistent with the molecular-weight determinations, however, and Albracht et al. (1979) have proposed an alternative in which there are two types of dimeric molecules in equal numbers, one containing center la and the other containing center 1 b. What function this arrangement might serve is


quite unknown. Were it not for the centers 1 a and 1 b stoichiometries in submitochondrial particles, most of the results obtained with isolated complex I might have been explained by partial denaturation of the enzyme during the purification. Certainly, this would account for the rotenone titer, for example. However, there seems to be no good reason for assuming this, now that the same iron-sulfur center stoichiometry has been found both for the isolated and membrane-bound enzyme.

For either of the models proposed, the monomeric units most probably have the same subunit composition (as explained above), and each contains an NADH binding site, as there is probably one NADH binding site per FMN (Section 3.7.1). Therefore, for the structural work to be described in this chapter, I will assume that the enzyme contains only one FMN per molecule with the proviso that this unit, in fact, represents only one-half or one-fourth of the structure. One further complication of a possible oligomeric dehydrogenase is described in Section 5.2. Finally, it should be noted that the epr center stoichiometry is not universally agreed, and a simple 1 : 1 molar ratio of center la or 1 b to FMN is favored by Ohnishi (1979).

2. THE PROTEIN COMPONENTS OF NADH DEHYDROGENASE

2.1. Fragmentation of the Enzyme

NADH dehydrogenase may be broken into fragments of much lower molecular weight by a wide variety of treatments. At one time, this property was a source of great confusion, as one of the fragments is reasonably stable and catalyzes the oxidation of NADH by a variety of electron acceptors including ubiquinone analogues. The catalytic properties of this FMN-containing fragment depend to a degree on the method used for its release from isolated NADH dehydrogenase or the membrane, and as a result a number of such preparations were described and thought to be the true NADH dehydrogenase in soluble form. It was only when Singer and collaborators (Watari et al .• 1963; Cremona et al .. 1963) showed that the fragment could be isolated from purified high-molecular-weight NADH dehydrogenase that the confusion was cleared up. Most of the methods used to solubilize this fragment, such as aqueous ethanol or proteolytic digestion (Watari et al.. 1963; Cremona et al .. 1963), lead to loss of nonheme iron or protein. The use of chaotropic agents allows the purification of this fragment without destruction of iron-sulfur groups (Hatefi and Stempel, 1967) and also allows the purification of another soluble fragment containing no FMN, but substantial quantities of iron and acid-labile sulfide (Hatefi and Stempel, 1969). The extent to which these fragments can be used to help analyze NADH dehydrogenase structure and func-

272 C. I. Ragan

tion will be described in later sections. For the present, the most useful application of chaotropic resolution has been to the analysis of the polypeptide composition of NADH dehydrogenase.

2.2. Fragmentation by Treatment with Chaotropic Agents

Briefly, chaotropic resolution involves incubation of NADH dehydrogenase (usually in the form of complex I) with a variety of agents such as urea, sodium perchlorate, or sodium trichloracetate. The precise conditions of reagent, reagent concentration, time, and temperature have been extensively analyzed by Davis and Hatefi (1969). There is little to choose between the methods. Hatefi and co-workers originally used urea, and more recently sodium perchlorate. The latter causes maximum solubilization of approximately 30% of the total protein (Ragan, 1976a). The residue becomes extremely insoluble during the resolution process and cannot be dissolved in urea or detergents other than sodium dodecyl sulfate. The water-soluble material can be fractionated by ammonium sulfate precipitation to yield the iron-sulfur fragment, which does not contain FMN, and the flavoprotein fragment (Hatefi and Stempel, 1969), which contains 13.5 nmoles FMN jmg protein and 5-6 iron atoms per molecule of FMN (Galante and Hatefi, 1979). Thus the flavin content is increased approximately tenfold over the starting material.

2.3. Polypeptide Composition of NADH Dehydrogenase and Its Subfragments

The polypeptides of complex I have been analyzed by SDS-polyacrylamide gel electrophoresis; similar results have been obtained by a number of workers (Hare and Crane, 1974; Ragan, 1976a; Crowder and Ragan, 1977; Y. M. Galante and Y. Hatefi, unpublished observations quoted in Hatefi et al., 1979; Dooijewaard et al., 1978a; Heron et al., 1979b). Differences between the results can be attributed to variations in gel composition, buffer composition, and molecular weight calibration, and are not significant. Table I shows the molecular weights of the constituent polypeptides as reported in the abovementioned publications. Hare and Crane (1974) and Dooijewaard et al. (1978a) reported fewer components than did the others. However, the resolving power of the gel systems they used was rather low, particularly in the lowmolecular-weight range. Galante and Hatefi (1979) found the polypeptide composition to be very similar to that reported by Ragan (1976a), using the Weber and Osborn (1969) buffer system, or Crowder and Ragan (1977), using a discontinuous buffer system with increased resolution. The final column of Table I shows the polypeptides resolved by two-dimensional analysis using isoelectric focusing in one dimension and discontinuous SDS-gel polyacryl-


Table I Subunit Composition of NADH Dehydrogenase"

Hare and Crane Ragan Crowder and Hatefi et a/. Dooijewaard Heron et al. ( 1974) ( 1976a) Ragan (1977) (1979) et al. (1978a) (1979b)a

74 75 75 75 77 75

53 53 b 53 52.5

56 53

49 48.5 49 42 42 42 45

40 42

37 39 39 36 39 33 33 32 33 29 30 33 28 30

27 26 27 27

26 27 26A'

25 26B 25 22 25

23.5 23.5 23.5 23.5 22A

23 22 22 21 18 22B 22C 21

20.5 20.5 20 19 18 18 19 18A

18B 16.5 16 16.5

15.5A 16 15.5 15.5 13 13 15.5B

15.5C 15.5D

8 8 9 8 5 5 <8 5

"10- 3 X molecular weight according to several studies. "Two subunits of this molecular weight were reported. 'Capital letters after the numbers in the last column distinguish subunits of the same apparent molecular weight which are separated by isoelectric focusing as shown in Figure 3.

amide gel electrophoresis in the second (Heron et al., 1979b). These findings are described at greater length below. Taking account of the different resolving powers of the gel systems used, there seems to be a good measure of agreement between different workers on the polypeptide composition.

Figure 1 shows the polypeptide composition of complex I analyzed on 12.5% (wjv) acrylamide gels by the method of Weber and Osborn (1969). Also shown are the polypeptides of the iron-protein and flavoprotein fragments and the insoluble residue resulting from treatment with sodium perchlorate. Clearly, certain polypeptides are quantitatively extracted by chaotropic agents,

274 C. I. Ragan

53

5

c~ _____ ~~ ______ ~

FIGURE 1. The polypeptide composition of NADH dehydrogenase and its subfragments analyzed by SDS-polyacrylamide gel electrophoresis, as described by Weber and Osborn (1969). Samples were run on 12.5% (w/v) acrylamide gels. (a) Complex I, (b) iron-protein fragment, (c) flavoprotein fragment, (d) insoluble residue from perchlorate treatment. Origin was on the left. Molecular weights (in thousands) are given.

and these have molecular weights of 75,000, 53,000, 30,000, and 27,000. The iron-protein fraction contains the 75,000- and 30,000-molecular-weight polypeptides and smaller amounts of several others of lower molecular weights. The flavoprotein fragment contains the rest of the 53,000-molecular-weight polypeptide, the 27,000-molecular-weight polypeptide, and small amounts of a lower-molecular-weight subunit that was not clearly shown in this experiment. These results are broadly in agreement with others using similar electrophoretic systems. Galante and Hatefi (1979) found that the flavoprotein fragment consisted of three subunits in equal molar proportions with molecular weights of 51,000,24,000, and < 10,000. Dooijewaard et al. (l978b) found subunits of molecular weights 56,000, 28,000, and occasionally one of 12,000. The presence of the latter polypeptide was correlated with an increased iron and acidlabile sulfide content, although apparently this was not true of the preparations of Galante and Hatefi (1979). Dooijewaard et al. (1978b) failed to achieve clear separation of subunits by ammonium sulfate fractionation of the material solubilized by chaotropic agents (in their case, urea). Although this failure was caused in part by the poor resolution achieved on their polyacrylamide gels, it is puzzling that even their flavin-rich fraction, with a polypeptide composition the same as that of the flavoprotein fragment, failed to precipitate with ammonium sulfate at the concentration reported by Hatefi and Stempel (1967) and repeatedly confirmed by myself and my colleagues. Consequently, although


their iron-rich fractions bore some resemblance in polypeptide composition to the iron-protein fragment of Figure 1, other polypeptides were present that are hard to account for. One possible factor may have been the residual detergent concentration in their complex I preparations. Chaotropic resolution works just as effectively on complex I from which residual cholate and deoxycholate have been removed by dialysis. Increasing the cholate concentration renders an increasing proportion of the protein apparently "soluble" after chaotropic resolution. For example, sodium perchlorate (0.5 M) in the presence of only 1 % cholate causes apparent solubilization of 85% of complex I protein (C. I. Ragan, unpublished observations). Normally, the residual detergent concentration in complex I preparations is insufficient to affect the solubilization by chaotropic agents. However, the preparation of Dooijewaard et al. (l978b) may have contained more cholate with the result that extra protein was solubilized and interfered with the subsequent ammonium sulfate fractionation.

The lower-molecular-weight subunits of complex I and the chaotropically resolved fragments are separated rather better using discontinuous gel electrophoresis. Figure 2 shows the polypeptide composition thus obtained. The third

FIGURE 2. The polypeptide composition of NADH dehydrogenase and its subfragments analyzed by discontinuous SDS-gel electrophoresis. Samples were the same as those in Figure 1.

276 C. I. Ragan

subunit of the flavoprotein fragment can be clearly seen, and the 53,000-molecular-weight polypeptide of this fragment can be seen to have a different mobility from the polypeptide of similar molecular weight in the iron-protein fragment. This latter subunit migrates with a molecular weight of 49,000 in the discontinuous system. Some contamination of the iron-protein fraction with flavoprotein fragment subunits is frequently seen. This can be eliminated by gel filtration of the iron-protein fraction on Sephadex G-l 00. One further difference between the continuous and discontinuous gels is that the polypeptides of molecular weights 33,000 and 30,000 comigrate in the latter system, with an apparent molecular weight of 30,000 (Crowder and Ragan, 1977). Although the well-separated high-molecular-weight subunits can be unambiguously identified on gels under different conditions, despite minor variations in apparent molecular weights, this is not so for the smaller subunits. As these subunits are incompletely resolved it is possible that, for example, the group of polypeptides that have an apparent molecular weight of 22,000 in the Weber and Osborn (1969) system do not all comigrate with the same apparent molecular weight in any other gel system. Indeed, in some of the labeling experiments to be described later, this does seem to be the case. To try to minimize confusion, the type of gel system used is stated in all the experiments to be described. Where a polypeptide can be unambiguously identified on any gel system, only a single apparent molecular weight is given, even if this may vary somewhat depending on conditions. This accounts for what at first sight appear to be rather odd molecular weights in certain instances (e.g., comparing the relative mobilities of the 53,000-, 49,000-, 42,000-, and 39,000-molecularweight subunits in Figure 2).

From the intensity of the Coomassie Blue staining of the subunits, a rough estimate of their molar stoichiometry could be made (Ragan, 1976a). Table II summarizes the results obtained. All the well-resolved high-molecular-weight subunits seem to be present in the same concentration as FMN. However, the low-molecular-weight subunits would have given much higher stoichiometries by this method, which suggests that there may be present several incompletely resolved proteins of similar molecular weights. To try to achieve a complete separation of the subunits, Heron et al. (1979b) used two-dimensional analysis with isoelectric focusing in the first dimension and discontinuous SDS-polyacrylamide gel electrophoresis in the other. Some improvements in separation in just the electrophoretic direction were achieved by using I-mm-thick slab gels rather than cylindrical gels. Thus the 22,000- and 20,500-molecularweight polypeptides were resolved into two pairs of closely migrating species.

The results of two-dimensional analysis are shown in Figure 3. As expected, several polypeptides were resolved into more than one component of the same molecular weight but having different isolectric points. Some polypeptides were lost during isoelectric focusing because their isoelectric points

NADH Dehydrogenase Structure

Table II Stoichiometry of NADH Dehydrogenase

Subunits

Molar ratio relative 10- 3 X mol wt to FMN

75 0.95' 53 2.24,·b

42 + 39 2.12' 33 0.90' 30 0.91 ' 27 1.0' 23.5 0.87"

1.04d

15.5A' 1.0'

'Based on the percentage of the total protein calculated from the intensity of Coomassie Blue staining on gels and the FMN content of complex I.

bOn Weber and Osborn (1969) gels, two different subunits comigrate with this molecular weight.

'Based on the composition of the flavoprotein fragment.

'Based on diphenyleneiodonium binding (Section 3.7.3).

'The 15,500 dalton subunit of the flavoprotein fragment is shown as IS.SA in Figure 3.

277

lay outside the pH range obtainable with commercially supplied ampholines. Hence there are still some uncertainties as to the exact number of constituent proteins. Table III summarizes the results obtained by the two-dimensional method. The flavoprotein fragment was still found to comprise only three subunits, whereas the iron-protein fragment was found to consist of eight subunits, three of which had very similar molecular weights.

The multiplicity of polypeptides in complex I makes it difficult to decide which components might be merely impurities. Certainly, the proteins listed in Table III are consistently seen in different preparations of complex I but more direct evidence that they are indeed bona fide subunits has come from immunological studies (Smith and Ragan, 1978, 1980; Heron et al., 1979b). Antisera raised against either complex I or the iron-protein fragment coordinately precipitate all the subunits of the enzyme with the exception of the 42,000-molecular-weight polypeptide. This component appears to separate from the enzyme in the presence of Triton X-I 00, which was added to ensure complete solubilization of the enzyme before immunoprecipitation. This polypeptide can also be removed by ultracentrifugation in the presence of EDT A and dithiothreitol or Triton X-IOO (Dooijewaard et aI., 1978a). However, all these treat-

278

-Isoelectric

b III 53

~ ----~

Q. 26A e - 25 - ~ ..:-E!!22C a: 2 ~18B iijl --l 1

, , BC

Basic

-.-.

focusing

75 -49 ~. 42

30 27·~26B

., 1550

C. I. Ragan

-

8 •

Acidic

FIGURE 3. Two-dimensional analysis of the subunits of NADH dehydrogenase. Complex I subunits were separated in the first dimension by isoelectric focusing (pH 3-10) in the presence of Nonidet P-40 and urea, and then in the second dimension by discontinuous SDS-gel electrophoresis. (a) Photograph of a stained gel. Also shown is a marker track of complex I subunits separated by SDS-gel electrophoresis only. (b) Key showing molecular weights (in thousands) of subunits consistently found in complex l. Subunits of similar molecular weights are further indicated by a capital letter.

ments lead to a loss of rotenone-sensitive ubiquinone reductase activity, so it cannot be definitely concluded that this polypeptide is an impurity. The observation that, with the exception noted, all the polypeptides of the dehydrogenase are coordinately precipitated by antiserum raised against a maximum of eight subunits (the iron-protein fraction) is strong evidence that all the others are true constituents of the enzyme. Several minor components in the two-dimensional gel of Figure 3 have not been labeled because they were not found in immunoprecipitates or have been identified as constituents of other mitochondrial enzymes (Heron et ai., 1979b).

The main conclusion to be drawn from these results is that NADH dehydrogenase is a lot more complicaled than one would have wished or anticipated.


Table III Subunit Composition of NADH Dehydrogenase Subfragments'

103 X mol wt Iron-protein Flavoprotein Insoluble residue

75 V 53 V 49 V 42 V 39 V 33 V 30 27 26A b V 20B V 25 V 23.5 V 22A 22B V 22C V 21 V 20 V 18A 18B V 16.5 V 15.5A 15.5B V 15.5C V 15.5D V 8 V 5 V

"The composition of the various subfragments was determined by two-dimensional analysis.

'Capital letters distinguish subunits of the same apparent molecular weight as shown in Figure 3.

3. THE PROTEIN STRUCTURE OF NADH DEHYDROGENASE

3.1. General Properties of MuItisubunit Enzymes

279

In an enzyme of this complexity it is only possible to talk about structure on a gross scale. Whereas for small enzymes, including some membrane-bound proteins, "structure" refers to the disposition of the amino acid chain, for NADH dehydrogenase and similar multisubunit entities such as the ribosome, we have not got beyond the stage of trying to define the relative disposition of individual subunits. There is an important assumption in the kind of work to

280 C. I. Ragan

be described, which is that each constituent polypeptide is structurally distinct and not interwoven with the others. This means that if it is possible to prove that one residue of a subunit is in a particular location, that the rest of the subunit is likely to be in the same region of the overall structure and not threaded in and out of it. Because the use of structural probes generally involves attachment of the probe or modification at only a limited number of sites on a polypeptide, this assumption is necessary to make any kind of sense of the data. There is no proof that this idea is correct, as no structures of this type have been solved. However, it seems to me that this is likely from what little is known of the biosynthesis of mitochondrial multisubunit enzymes. For example, it is now known that the three larger subunits of the F1-ATPase of yeast mitochondria are synthesized in the cytoplasm as precursor molecules of greater molecular weight than that of the final products (Maccecchini et aI., 1979). These are then independently transported into the mitochondrion, cleaved to the correct molecular weight, and then in some unknown fashion assembled into the final ATPase structure. Clearly, the separate synthesis and transport of each polypeptide allows ample opportunity for the protein to fold into a stable tertiary structure that will probably be only slightly different from that of the final smaller subunit. Thus the polypeptides arrive at their site of assembly into the ATPase already prefolded, presumably existing in the final assembled enzyme in much the same conformational state. The opportunity for polypeptide chains to become extensively tangled up with each other would seem negligible. This may not, of course, be true of those proteins synthesized on mitochondrial ribosomes, as it is possible that these hydrophobic polypeptides are incorporated directly into the membrane during synthesis. However, many of the NADH dehydrogenase polypeptides are presumably synthesized on cytoribosomes, particularly those on the outside of the structure and those that are more hydrophilic.

3.2. The Nature of Chaotropic Resolution

The process of chaotropic resolution itself can give some clues to the structure of NADH dehydrogenase. The mechanism whereby anions such as CIO,;-, SCN-, and CCI3COO- cause solubilization of membrane proteins has been discussed at length by Hatefi and Hanstein (1969). Basically, these anions make water disordered and more lipophilic, thereby increasing the aqueous solubility of nonelectrolytes. The insolubility of the side chains of hydrophobic amino acids in water is the basis of hydrophobic bonding in proteins (e.g., Kauzmann, 1959), and the principal effect of chaotropic anions would be to favor transfer of hydrophobic residues to water. Secondary effects could be exerted on certain types of hydrogen bonds (C=O ... H - Nand C=O ... H -0) which are known to be more stable in polar environments (Klotz and Farnham, 1968), and therefore could be affected by transfer to an aqueous environment.


The interesting point about the effect of chaotropic anions on NADH dehydrogenase is that the fragments that are solubilized by such treatments are soluble in water even in the absence of the anions. Thus the chaotropes seem to act by disrupting hydrophobic interactions in the enzyme and by allowing hydrophilic fragments to escape into aqueous solution. If anything, the unsolubilized material seems to be even more hydrophobic as a result, as it is insoluble in all but the most powerful detergents such as sodium dodecyl sulfate.

The use of the terms "hydrophobic" and "hydrophilic" in connection with these enzyme fragments is rather loose. Proteins that are insoluble in water do not necessarily have a larger proportion of hydrophobic amino acids than those that are soluble. Obviously, what matters is the tertiary structure and the distribution of ionic and hydrophobic groups on the surface of protein. Alterations to the tertiary structure will change the distribution of amino acid residues, and therefore the apparent hydrophobicity (most proteins can be made "hydrophobic" by denaturation). To what extent do the fragments solublized by chaotropic anions retain the native conformation? Although the catalytic porperties are altered, the prosthetic groups are not destroyed, and Dooijewaard and Slater (1976a,b) have provided evidence that the initial stages of NADH oxidation by the flavoprotein fragment are kinetically very similar to those of NADH dehydrogenase. It therefore seems reasonable to assume that the conformation of the chaotropic fragments is not changed greatly by solubilization. Consequently, the use of the terms hydrophobic and hydrophilic to describe their surface properties in the intact dehydrogenase and in isolation seems to have some justification.

The first proposals for NADH dehydrogenase structure were those of Hatefi and Stempel (1969). They noted that dithionite was unable to reduce the chromophores of NADH dehydrogenase unless the enzyme had been treated with chaotropic agents. Similarly, water-soluble iron chelators, such as bathophenanthroline sulfonate, only reacted readily with the iron-sulfur groups of the enzyme after treatment with chaotropes. Taken together with the release of hydrophilic fragments by chaotropic anions, these observations suggested that the hydrophilic flavoprotein and iron-protein fragments were surrounded in the intact enzyme by a hydrophobic sheath of lipids and proteins (Hatefi and Stempel, 1969). The more detailed studies that follow show this to be basically correct.

3.3. Isoelectric Points of the Constituent Polypeptides

The two-dimensional analysis of complex I subunits also provides a further item of information about these constituents, i.e., their isoelectric points. For a polypeptide denatured in urea and a nonionic detergent, the isoelectric point merely reflects the average balance between acidic and basic amino acid residues in the sequence, as the polypeptide chain is unfolded and all residues

282 C. I. Ragan

will contribute. A statistically significant correlation (p < 0.01) was found between the isoelectric point and insolubility after chaotropic resolution (Heron et ai., 1979b). Thus the polypeptides of the hydrophobic reside had a higher mean isoelectric point than those that were rendered soluble. It is possible that the former proteins interact with the phospholipid bilayer of the membrane not only by hydrophobic interactions with the fatty acid side chains, but also by electrostatic interaction with negatively charged phosphate groups. The strong association of cardiolipin (with two phosphate groups per molecule) with NADH dehydrogenase may be an indication of such an interaction (Heron et aI., 1977). This proposal would agree with that of Hatefi and Stempel (1969) for a hydrophobic sheath around the flavoprotein and iron-protein fragments.

3.4. Labeling with Hydrophilic Probes

3.4.1. Diazobenzenesulfonate

Diazobenzenesulfonate has been in common usage for several years as a probe for proteins on the surface of membranes. It is extremely soluble in water, has a fairly wide reactivity (Howard and Wild, 1957; Higgins and Harrington, 1959), and does not penetrate membranes (Schneider et aI., 1972). It has also been used for labeling the surface of multisubunit enzymes such as cytochrome oxidase in which two subunits (I and IV), which were not labeled by diazobenzenesulfonate unless the enzyme had been treated with sodium dodecyl sulfate, were deemed to be buried in the core of the structure (Eytan et ai., 1975). It is not possible to say with certainty whether the protein was not labeled because it was situated in a hydrophobic environment in which the probe was insoluble or whether it was not labeled because its susceptible groups were physically hindered from reaction with the probe. For cytochrome oxidase it seems to be implicitly assumed that both mechanims apply, i.e., that subunits I and IV are both hydrophobic and inaccessible (Eytan et ai., 1975).

Labeling of NADH dehydrogenase with diazobenzene [35S]sulfonate appears to label most of the constituent polypeptides (Smith and Ragan, 1980). However, resolution of the enzyme by chaotropic agents shows that there is indeed some selectivity about the labeling. In Figure 4, the labeling profiles of complex I and its subfragments are shown. Whereas in the whole enzyme the distribution of label appears to be fairly uniform, the soluble fragments from treatment with sodium perchlorate take up relatively little label. Thus the ironprotein fragment is labeled in the 75,000-, 49,000-, and 15,500-molecularweight polypeptides (Smith and Ragan, 1979). The labeling in the 8000-molecular-weight region is caused by a heavily labeled impurity in the ironprotein preparation. The polypeptides of molecular weights 30,000, 22,000, and 18,000 were scarcely labeled at all, although they readily reacted with


FIGURE 4. Labeling of NAOH dehydrogenase subunits with diazobenzene-["S]sulfonate. Complex I was treated with diazobenzene-[3SS]sulfonate and then resolved with perchlorate. The iron-protein and flavoprotein fragments were isolated and, together with complex I, were analyzed by discontinuous SOS-gel electrophoresis. Gels were stained, sliced, and counted for radioactivity. (a) Complex I, (b) iron-protein fragment, (c) flavoprotein fragment.

283

8 "'49 a

6

....... 4 E ci. ~2 :?:' .~ U 4 ~

49 '0 ~ 2 x

<:" 0 ~

2 c

------------~~-------~ o 10 20 30 40 50 60 70 80 90 100110

Slice no. from origin

diazobenzenesulfonate after treatment of the enzyme with chaotropic agents. The same is true of the subunits of the flavoprotein fragment, none of which could be labeled by diazobenzenesulfonate in intact NADH dehydrogenase (Smith and Ragan, 1979).

Thus several of the polypeptides previously shown to be part of the hydrophilic fragments are inaccessible to diazobenzenesulfonate. The most reasonable explanation is that these subunits are buried within the enzyme structure and that the hydrophobic sheath around them prevents the reagent from gaining access to them. Of particular interest is the inability of the reagent to get to the flavoprotein fragment which must necessarily be accessible to NADH in the intact enzyme.

3.4.2. Lactoperoxidase-Catalyzed Iodination

Another probe for proteins exposed on the surface of membranes is the enzyme-catalyzed incorporation of 1251 (Phillips and Morrison, 1971). Because this probe has different chemical specificity to diazobenzenesulfonate, a comparison of the effect of these two reagents is important to establish whether lack of labeling is due to inaccessibility or merely to the absence of susceptible residues.

Radioiodination of intact NADH dehydrogenase (Figure 5) produced results very similar to those obtained with diazobenzenesulfonate labeling

284 c.1. Ragan

10 53

8

E 42 a

Ci6 u ~ 75 >.

~4 ...... ~

.Q 2 u ~

N X , $2

1 c

o 10 20 30 40 50 60 70 Slice no. from origin

FIGURE 5. Labeling of NADH dehydrogenase subunits by lactoperoxidase-catalyzed radioiodination. Complex I was labeled with 1251 and further treated as in Figure 4. (a) Complex I, (b) iron-protein fragment, (c) flavoprotein fragment.

(Ragan, 1976a; Smith and Ragan, 1979). The iron-protein fragment was labeled mainly in the 75,000- and 49,000-molecular-weight polypeptides, whereas the subunits of the flavoprotein fragment were not labeled at all. Furthermore all the polypeptides of these fragments could be labeled if radioiodination was carried out after chaotropic resolution (Ragan, 1976a). A polypeptide of molecular weight 23,500 (using the Weber and Osborn, 1969, system) also appeared to be inaccessible to radioiodination (Ragan, 1976a,b).

Because both hydrophilic probes produce very similar results, it can be concluded that those polypeptides that are not labeled are, in fact, inaccessible by virtue of being buried in the overall structure. Of the II polypeptides solubilized by chaotropic agents (i.e., hydrophilic), between six and eight were not accessible to labeling. On the other hand, of the remaining 15 or so hydrophobic polypeptides, at least 11 were labeled with either reagent. Thus, despite the "hydrophobic" designation, these proteins are on the surface of the isolated enzyme. The degree to which these proteins are accessible when the enzyme is in the membrane is described in Section 5.1.


3.5. Labeling with a Hydrophobic Probe

Iodonaphthylazide is a photolabile reagent introduced by Bercovici and Gitler (1978) for labeling hydrophobic regions of membranes. It is extremely hydrophobic and, in the dark, it partitions into the lipid phase of the membrane. Photoactivation generates the highly reactive nitrene, which can then react with a wide variety of neighboring groups. Thus it is regarded as a probe for those polypeptides that are adjacent to the phospholipid molecules in a membrane. As anticipated, this compound behaves very differently from the two hydrophilic probes just described. The labeling pattern obtained by photolabeling NADH dehydrogenase with iodonaphthylazide is shown in Figure 6 (F. G. P. Earley and C. I. Ragan, unpublished observations).

Very little incorporation into high-molecular-weight subunits (75,000, 53,000, 49,000) was found, with most of the label appearing in several of the low-molecular-weight hydrophobic subunits of molecular weights 22,000, 18,-000, and 15,500. In contrast, prior treatment of the enzyme with SDS allowed iodonaphthylazide to photolabel most of the polypeptides, in particular those of highest molecular weight. Resolution was greatly increased by using thin slab gels and autoradiography to detect the incorporation (Figure 7). Track 1 is of complex I and shows that the major labeled proteins are those of molecular weights 39,000, 22,000, 18,000, 16,500, 15,500, and below (the smallest subunits were not resolved in this experiment). Tracks 2-4 show that supplementation of the enzyme with phosphatidylcholine, rotenone, or diphenyleneiodonium (Section 3.7.3), respectively, did not alter the pattern of labeling.

12 15.5

10 ~8 - 505 t; 6 0.

34

.~

~ 53 Q 12 u

~ 10 +505 FIGURE 6. Labeling of NADH dehydroge- c;>

nase subunits with [1251] iodonaphthylazide. $2 8 75

Complex I, or complex I dissociated with SDS, 6 was photolabeled with [mIl iodonaphthylazide and analyzed by SDS-gel electrophoresis, as described by Weber and Osborn (1969). Labeled phospholipids migrated with the dye 0 20 40 60 80 100 front. Slice no. from origin

286

Origin

22

18 165 155

Track

2 3

3.6. Proteolytic Digestion

4

C. I. Ragan

FIGURE 7. Labeling of NADH dehydrogenase subunits with ['25I]iodonaphthylazide. Complex I was labeled with [I25I]iodonaphthylazide and analyzed by discontinuous SDS-gel electrophoresis and autoradiography. Track 1, complex I; track 2, complex I supplemented with phosphatidylcholine (0.5 J,Lmoles/mg protein); track 3, complex I plus rotenone (l nmole/mg protein); track 4, complex I plus diphenyleneiodonium (200 nmoles/mg protein).

Digestion by proteolytic enzymes is a technique that has been rather widely used for investigating the structures of multisubunit complexes. In trying to relate structure and function, it suffers from the drawback that even if several peptide bonds in a particular polypeptide are hydrolysed, the tertiary structure of the chain may not be altered, and its functional activity may not change (e.g., Sale et aI., 1977). A technical difficulty may arise when trying to analyze which polypeptides are being degraded, because low-molecular-weight degradation products may comigrate on gel electrophoresis with authentic lowmolecular-weight constituents.

Several proteolytic enzymes have been shown to have similar effects on the activity of NADH dehydrogenase (Cremona et al., 1963). Incubation with proteases causes eventual release of the flavoprotein fragment, which itself is extremely resistant to degradation. The stages in protein degradation that lead to release of the flavoprotein fragment were investigated using both trypsin (Ragan, 1976b) and chymotrypsin (Crowder and Ragan, 1977). The NADHK3Fe(CN)6 oxidoreductase activity of NADH dehydrogenase was far more


resistant to proteolytic digestion than was the NADH-ubiquinone oxidoreductase activity. With either protease, the polypeptides degraded during the loss of ubiquinone reductase activity were similar (Figure 8), i.e., those of molecular weights 49,000, 42,000, 39,000, 26,000, 23,500, 20,500, 18,000, and 15,-500. Some differences in the relative rates of degradation were apparent, however. For example, the polypeptide of molecular weight 20,500 was particularly sensitive to trypsin (Ragan, 1976b). At a time when the rotenone-sensitive reduction of ubiquinone was abolished, there was neither alteration of the NADH-K3Fe(CN)6 oxidoreductase activity nor solubilization of the flavoprotein fragment. Indeed, the two larger subunits of the flavoprotein fragment can clearly be seen in the gels of Figure 8. Also, it is noteworthy that at least two of the iron-protein subunits of molecular weights 75,000 and 30,000 were unaffected, although the 49,000-molecular-weight constituent was extensively degraded. Despite this, a modified iron-protein fragment could still be isolated that contained the degraded 49,000-molecular-weight subunit (Crowder and Ragan, 1977).

The release of the flavoprotein fragment required more extensive treatment than that shown in Figure 8. The time course (Figure 9) shows that loss of NADH-K3Fe(CN)6 oxidoreductase activity followed the release of the 53,000-molecular-weight subunit. In looking for what factors might be

49

75

a

39 22 18 15.5 ~

FIGURE 8. Degradation of NADH dehydrogenase subunits with chymotrypsin. Complex I (I mg protein/ml) was incubated with chymotrypsin (20 ILg/ml) at 21 °C. At various times samples were analyzed by discontinuous SDS-gel electrophoresis: (a) 0 min, (b) 50 min, (c) 100 min, (d), 200 min, (e) 300 min.

288

~ 80 rtl C 01 .§ 60

'0 ~ 40 rtl C (])

~ 20 cf

o

C. I. Ragan

•

234 5 6 Incubation time(hr)

FIGURE 9. Time course of the release of the flavoprotein fragment from NADH dehydrogenase by chymotrypsin. Complex I (I mg protein/ml) was incubated with chymotrypsin (100 /lg/ ml) at 21 °C for the indicated times. Samples were assayed for NADH-K, Fe(CN)6 oxidoreductase activity (e) and centrifuged, and both pellet and supernatant fractions were analyzed by discontinuous SDS-gel electrophoresis. (4) Percent degradation of the 30,OOO-molecular-weight subunit in the pellet fraction; (_) percent release of the 53,OOO-molecular-weight flavoprotein fragment subunit into the supernatant. Because the flavoprotein fragment is enzymically active, NADH-K, Fe(CN)6 oxidoreductase activity does not decline to zero.

involved in release of the flavoprotein fragment it was noted that one polypeptide-that of 30,000 molecular weight-was degraded by trypsin or chymotrypsin with a very similar time course (Crowder and Ragan, 1977). During the period of release of the flavoprotein fragment, this was the only polypeptide to undergo extensive degradation.

The results of proteolytic digestion are complicated and the purpose of this article would not be served by introducing too many of them. The changes in the polypeptide structure accompanying loss of NADH-ubiquinone oxidoreductase activity are far from simple and do not lead to any obvious structure/ function relationships. Although several of those polypeptides that are more readily degraded by either trypsin and chymotrypsin are on the surface of the structure according to labeling experiments described above, there are exceptions (e.g., the 75,000-molecular-weight polypeptide, which is resistant to proteolytic attack).

The time course of release of the flavoprotein fragment is perhaps more informative. During the distinct lag phase most of the eventual protein degradation is in process. After this stage, the dehydrogenase structure seems to be opened up to allow digestion of the 30,000-molecular-weight polypeptide, which is therefore implicated in. the binding of the flavoprotein fragment.


Because this polypeptide is a subunit of the iron-protein fragment, these experiments may indicate a physical association between the flavoprotein and ironprotein fragments.

Proteolytic digestion has also been used to illustrate lipid-protein interactions in NADH dehydrogenase. These experiments are described in Section 4.4.1.

3.7. Specific Structure/Function Relationships

3.7.1. The NADH Binding Site

The site in NADH dehydrogenase that binds NADH can be narrowed down to somewhere within the three subunits of the flavoprotein fragment. Dooijewaard and Slater (l976a,b) concluded that the binding of NADH to both NADH dehydrogenase and the flavoprotein fragment was very similar. Thus the rate constants for NADH association and dissociation and NAD+ dissociation were the same in both types of preparation. The preparations differed basically in the kinetics of reduction of the acceptors. Using K3Fe(CN)6 as acceptor for NADH dehydrogenase, a ping-pong bi-bi mechanism with double substrate inhibition was proposed. In the flavoprotein fragment, depending on the acceptor and its concentration, an ordered mechanism was found to operate in addition to the ping-pong pathway, i.e., the reduced enzyme could be oxidized both before and after dissociation of NAD+.

As pointed out by Dooijewaard and Slater (l976a,b), it is unlikely that NADH and Fe(CN)~- would bind to the same site on NADH dehydrogenase. However, the kinetics clearly show competition between the two substrates. Dooijewaard and Slater (l976a,b) proposed that access for either substrate to the active center or centers of the enzyme is provided by a single cleft in the structure, which can be blocked by either NADH or Fe(CN)~-. In the flavoprotein fragment, artificial acceptors including Fe(CN)~- are not so restricted in their access to the enzyme. Indeed, the rate constant for reduction of Fe(CN)~- by the reduced enzyme is more than two orders of magnitude greater in the flavoprotein fragment. The increased access of the acceptors allows their reduction without prior dissociation of NAD+.

These conclusions are quite consistent with the proposals put forward in the previous section, in which it was suggested that the flavoprotein fragment is buried within NADH dehydrogenase. Clearly, access to NADH has to be provided and the nature of the cleft in the structure is such that Fe(CN)~- is the only electron acceptor that can use it. Ubiquinone, of course, is reduced by an entirely different pathway. The specificity of the cleft makes it not surprising that diazobenzenesulfonate, for example, is unable to enter and react with the flavoprotein fragment.

290 C. I. Ragan

3.7.2. The Iron-Protein Fragment

The iron-protein fragment exhibits an epr spectrum after reduction with dithionite which resembles that of center 2 in NADH dehydrogenase (OrmeJohnson et al .• 1974). The redox potential of the chromophore also appears to be quite high, as it is reducible by ubiquinol analogues (Hatefi and Stempel, 1967). Thus it is quite possible that the iron-protein fragment contains center 2, but such a conclusion has to be treated with caution, as only 5% of the iron atoms present can be accounted for in the epr spectrum (Or me-Johnson et al.. 1974), and the redox potential may have altered considerably following solubilization.

3.7.3. Inhibition by Diphenyleneiodonium

Diphenyleneiodonium is a potent and specific inhibitor of NADH oxidation (Holland et al.. 1973; Gatley and Sherratt, 1976). The possibility that it might inhibit by covalent reaction with NADH dehydrogenase was suggested by the known properties of iodonium compounds, which undergo nucleophilic attack in nonpolar solvents as shown in Figure 10. Ragan and Bloxham (1977) found that diphenyleneiodonium reacted preferentially with a polypeptide of molecular weight 23,500, and to a much lesser extent with others of molecular weights 75,000, 53,000, 42,000, 33,000, and 15,500 (Figure 11). Maximum incorporation into the 23,500-molecular-weight polypeptide was one molecule per molecule of FMN, and labeling was abolished by prior treatment with sodium dodecylsulfate or chaotropic agents. Although the polypeptide labeled by diphenyleneiodonium may also be one of those labeled by iodonaphthylazide, the specificity of the reaction and its sensitivity to denaturation of NADH dehydrogenase clearly distinguish the reaction from general hydrophobic labeling. Correlation of inhibition of NADH-ubiquinone oxidoreductase with incorporation of diphenyleneiodonium was more difficult to establish. The concentration of inhibitor required for 50% inhibition (20-26 nmoles/mg protein) was much less than that required for incorporation of 0.5 moles/mole FMN (200-

~----.~ ~fP ~ ~

X- X

FIGURE to. Reaction of diphenyleneiodonium with a nucleophile x~ in nonpolar solvents.


4

--- 23.5 E 5-3 3---:-

E

Z' + ci. DPI 0

> U2 2 >. ..... ro > 0 75 \J U ~ ro

0 x \J <";'J

$2 ~ ~x 10

0 10 20 30 40 70 Slice no. from origin

FIGURE 11. Labeling of NADH dehydrogenase subunits with diphenylene-[ I2SI]iodonium. Complex I was incubated with diphenylene-[ '25Iliodonium (240 nmoles/mg protein) and analyzed by SOS-gel electrophoresis. as described by Weber and Osborn (1969). The large peak of radioactivity at the dye front was caused by unreacted diphenyleneiodonium (OPI+) complexed with SOS.

350 nmoles/mg protein). However, owing to the nonhyperbolic nature of the curve relating inhibition with diphenyleneiodonium concentration, 100% inhibition was not attained until 1 mole inhibitor/mole FMN had been incorporated. A further interesting property of the inhibitor was that rotenone increased the apparent affinity of the 23,500-molecular-weight polypeptide for diphenyleneiodonium, clearly showing that the two inhibitors were not binding at the same site.

The nonlinear relationship between inhibition and inhibitor binding might be a further piece of evidence for the oligomeric nature of NADH dehydrogenase. Alternatively, the covalent modification may be only indirectly related to inhibition. For example, inhibition might be due to binding of diphenyleneiodonium to several hydrophobic regions, only one of which provides a nucleophile for covalent attachment or forms a covalent bond sufficiently stable to survive the analysis. However, this argument would not fit in with the considerable potency of diphenyleneiodonium compared with related ring structures, such as dibenzofuran, or iodonium compounds which less readily undergo nucleophilic attack (Ragan and Bloxham, 1977). The evidence seems strong that diphenyleneiodonium is a good inhibitor because of its chemical reactivity. Assuming that the 23,500-molecular-weight polypeptide is the target for this inhibitor, there is unfortunately little information on its properties. It is not solubilized by chaotropic agents and therefore is probably hydrophobic. It has

292 C. I. Ragan

a basic isoelectric point (Heron et al .. 1979b), but does not seem to be readily accessible to labeling by lactoperoxidase catalyzed iodination. Thus it could occupy an internal location in the enzyme, perhaps involved in binding whichever redox groups are responsible for transferring electrons from within the enzyme to the hydrophobic surface at which ubiquinone reduction takes place.

3.7.4. Inhibition by Azidodibenzofuran

An inhibitor structurally related to diphenyleneiodonium is 3-azidodibenzofuran, which was devised by Earley and Bloxham (1978). Dibenzofuran had been shown to be a specific, but rather weak, inhibitor of NADH dehydrogenase (Ragan and Bloxham, 1977). The attachment of the azido group had little effect on its inhibitory properties in the dark, but potent, irreversible inhibition of NADH-ubiquinone oxidoreductase activity occurred after photoactivation. Light-dependent attachment to protein has been shown, but the polypeptides involved have not yet been identified.

4. THE PHOSPHOLIPID COMPONENTS OF NADH DEHYDROGENASE

4.1. Are Phospholipids Essential?

Treatment of submitochondrial particles with phospholipase A results in a substantial decrease in NADH-ubiquinone oxidoreductase activity which can be partially restored (depending on the initial degree of inactivation) by addition of phospholipids (Machinist and Singer, 1965). This work has led to the proposal that the inability of certain NADH dehydrogenase preparations to reduce ubiquinone analogues in a rotenone-sensitive manner resulted from their lack of phospholipid. Restoration of this activity by addition of phospholipids would prove the point, but this has not been achieved. Partial removal of phospholipid from NADH dehydrogenase, can, under suitable circumstances, lead to reversible loss of ubiquinone reductase activity (Ragan and Racker, 1973). Reversibility seems to be related to stabilization of iron-sulfur center 2, which denatures during phospholipid removal unless protected by reduction (Ragan and Racker, 1973; Ohnishi et aI., 1974). Therefore it appeared that phospholipid might be essential to maintain the structure of one of the catalytic centers. However, Baugh and King (1972, 1974) upset this view by isolating a soluble lipid-free NADH dehydrogenase capable of rapidly reducing ubiquinone analogues and, at least for higher homologues such as ubiquinone-6, in a rotenone-sensitive manner. The kinetic characteristics of ubiquinone reduction were substantially altered and the enzyme was unstable compared with the


lipoprotein complex I. A possible explanation for this behavior will be given in Section 4.3.1. For the present, as it is obvious that NADH dehydrogenase is surrounded by phospholipids in the membrane, I will take it that these have a function in the activity of the enzyme and in the maintenance of NADH dehydrogenase structure. Possible roles for lipids are described in later sections.

4.2. Phospholipid Composition of NADH Dehydrogenase

The total lipid content of Complex I is 0.22 mg/mg protein, more than 90% of which is phospholipid (Hatefi and Stiggal, 1976). Taking a value of 33 J.Lg phosphorus/mg total mitochondrial lipid (Fleischer et al., 1961), this is equivalent to 0.24 J.Lmoles of lipid phosphorus per mg of protein in good agreement with Heron et al. (1977). The phospholipids present in bovine heart mitochondria are phosphatidylcholine, phosphatidylethanolamine, and cardiolipin, which together comprise 97% of the total phospholipid (Fleischer et al., 1967). The relative amounts in Complex I are 38%, 35%, and 21 % of the total lipid phosphorus, respectively (C. I. Ragan, unpublished observations), in agreement with the corresponding values for intact mitochondria which are 40%, 37%, and 20%, respectively (Fleischer et al., 1967).

4.3. Phospholipid Function

4.3.1. Ubiquinone Solvent

Because the natural acceptor for NADH dehydrogenase, ubiquinone-IO, is extremely hydrophobic, the site of reduction would be expected to be in a hydrophobic region of the membrane, i.e., within the lipid bilayer. Thus one obvious role of phospholipid would be to act as a solvent for ubiquinone.

In isolated complex I, phospholipid seems to fulfill a similar function. The rotenone-sensitive pathway of ubiquinone reduction is stimulated by supplementation of the assay system with phospholipids (Hatefi et al., 1960). The degree of stimulation depends on the type of ubiquinone analogue (i.e., its hydrophobicity) and its concentration. The rate obtained with any particular combination of phospholipid and ubiquinone concentrations can be rationalized in terms of partition of the ubiquinone between the phospholipid and aqueous phases of the system (Ragan, 1978). Increasing the phospholipid concentration causes more ubiquinone to enter the lipid phase. Fusion of added lipid with endogenous lipid was a prerequisite for this effect, as expected. Because the endogenous phospholipid of complex I behaves no differently from added phospholipid, it can be concluded that phospholipids are important as a solvent for ubiquinone.

A solvent role was also shown by studies on complex I, whose natural

294 C. I. Ragan

phosphatidylcholine and phosphatidylethanolamine had been replaced by synthetic dimyrstoyllecithin (Heron et al., 1977). The phospholipid undergoes transition between the liquid crystalline and gel phases at 24 0 C (Hinz and Sturtevant, 1972). In lipid-replaced complex I, such a transition in the dimyristoyllecithin was evident from measurements of the fluorescence polarization of diphenylhexatriene (V. Poore and C. I. Ragan, unpublished observations). However, Arrhenius plots of activity were linear between 50 and 35 0 C, just as with the native enzyme. However, if the dimyristoyllecithin to protein ratio was raised sufficiently, then ubiquinone reductase activity declined sharply below approximately 20 0 C, and the Arrhenius plot showed two distinct slopes. Only the reduction of this acceptor was affected (Ragan, 1978).

Clearly, the catalytic activity of the enzyme seems unconcerned by the physical state of the surrounding lipid. The effect on ubiquinone reduction is most likely to be on the rate of diffusion of the ubiquinone analogue in the lipid phase, which only becomes rate-limiting for oxidoreduction when the enzyme is surrounded by a large amount of gel-state lipid (Ragan, 1978).

These results do not imply any specific association between phospholipid and proteins which would lead to phospholipids being regarded as part of NADH dehydrogenase. However, they do show that the NADH dehydrogenase protein cannot be regarded as independent of its proper environment, which is both aqueous for NADH and lipid for ubiquinone.

The preparation of Baugh and King (1972) could reduce ubiquinone-l at high rates, but with a Km two to three orders of magnitude higher than that found with complex I or with submitochondrial particles. It is possible that the removal of phospholipid was accomplished without denaturation of any of the catalytic centers of the enzyme. Thus ubiquinone reduction could take place by the normal pathway, but the absence of lipid would not favor reduction at the hydrophobic site, with a consequent massive increase in the Km. I feel that the evidence is not overwhelming that ubiquinone analogues are really reduced at the correct site by this preparation of NADH dehydrogenase. Even if they are, the evidence given above shows that the absence of phospholipid is unnatural and that soluble NADH dehydrogenases are operating in the wrong environment. They are therefore of limited use for defining the mechanism of ubiquinone reduction in the membrane.

4.3.2. Structural Requirement

Reference has already been made to a possible structural requirement for phospholipids. By this, I mean a specific association of a phospholipid with a protein which is required for the correct functioning of that protein. Evidence for such a requirement was provided by Heron et al. (1977). Phosphatidylcholine and phosphatidylethanolamine can be removed by treatment of NADH dehydrogenase with cholate followed by precipitation with ammonium sulfate.


Activity declines linearly with phospholipid depletion until zero activity is reached at a limiting phospholipid content of 0.05 J.lmoles lipid phosphorus/mg protein. Further treatment with cholate did not reduce this figure, and analysis showed that this phospholipid was all cardiolipin. The difficulty encountered in removing this phospholipid might be an indication of tight binding of the lipid to protein.

Although cholate alone could not remove cardiolipin, it was possible to exchange the cardiolipin for other lipids in the presence of cholate (Table IV). At low cholate concentrations, dimyristoyllecithin replaced all the phosphatidyicholine and phosphatidylethanolamine with little loss of activity. At higher cholate concentrations, cardiolipin was also replaced and activity was lost. Exchange of endogenous lipids for exogenous cardiolipin also caused loss of activity. However, a mixture of cardiolipin and dimyristoyllecithin at high cholate concentrations did not lead to loss of activity, although all types of endogenous phospholipid were exchanged. It appears that both cardiolipin and phosphatidyicholine or phosphatidylethanolamine are required by the enzyme and that moreover, these must be bound to their respective sites, which have decreased affinity for the wrong class of lipid (Heron et al., 1977). Clearly, if phospholipids acted only as a ubiquinone solvent, then any phospholipid would do. The findings described above suggest, therefore, that In addition, phospholipids have a more specific function.

A specific role for cardiolipin is interesting, because it is present in so small an amount compared with the others. A similar concentration is found in purified cytochrome oxidase from which it can be removed or replaced by other lipids without loss of activity, according to Watts et .al. (1978). However, complete replacement has not been achieved by other workers, and it is still an

Table IV Lipid Exchange in NADH Dehydrogenase"

Phospholipid content (/Lmoles lipid P /mg protein)

Cholate Total Endogenous Exogenous Lipid added (%) phospholipid DML CL CL Total CL

0.23 0.049 0.049 DML 0.35 0.28 0.22 0.045 0.045 b

DML 2.5 0.31 0.26 0.019 0.019 DML + CL 2.5 0.30 0.22 0.009 0.033 0.042b CL 2.5 0.17 0.04 0.017 0.072 0.089

"Complex I was incubated with the indicated phospholipid and cholate concentration. Dimyristoyllecithin (DML) was added at a concentration of 6"moles lipid P /mg protein and cardiolipin (CL)from yeast at a concentration of 0.25 "moles lipid P/mg protein. The enzyme was separated from excess lipids by ammonium sulfate

.precipitation and assayed for activity. total phospholipid, DML, endogenous CL, and exogenous (i.e., yeast)CL. 'Only these samples had close to normal NADH-ubiquinone oxidoreductase activities.

296 C. I. Ragan

open question whether or not cardiolipin could have a special function in this enzyme.

The organization of cardiolipin in the membrane is unknown. Work with an antibody to cardiolipin has suggested that the antigenic determinant, the glycerol, and two phosphates in the center of the molecule, are not readily accessible from the surface of the membrane (Guarnieri et al., 1971). Because this corresponds to the head group region, the result is somewhat surprising. It was also proposed that this inaccessibility was attributable to binding to membrane proteins (Guarnieri et al., 1971). Thus it is possible that cardiolipin is not a constituent of the bulk bilayer, but that, in fact, it is more or less permanently associated with integral membrane proteins. The possibility of ionic binding of the head group phosphates to basic hydrophobic proteins has already been raised (Section 3.3).

4.4. Specific Lipid-Protein Interactions

The previous section has concentrated on the lipid structure. This section concerns alteration in the protein structure or function induced by removal of phospholipids.

The functional effects of lipid removal have been described. Treatment of complex I with cholate or phospholipase A leads to loss of ubiquinone reductase activity. Obviously one factor involved is removal of the solvent for ubiquinone, but this is not the whole story, for the redox potential of center 2 is shifted by about -80 mY, and this change can be partially reversed under suitable circumstances (Ohnishi et al., 1974). This center, in view of its high redox potential, is the most likely candidate f()r the reductant for ubiquinone. It seems logical, therefore, to suppose that it is located near to the phospholipid on the outside of the protein and reasonable that its redox properties would be altered by removal of phospholipids and exposure of the center to water. The decrease in redox potential would suggest that it is the reduced form of the center that is most affected by lipid removal. An attractive possibility is that the center moves relative to the rest of the structure on oxidoreduction. Thus the oxidized form is relatively internal when it receives an electron and is pushed out into the lipid to donate the electron to ubiquinone.

Removal of phospholipids also makes the enzyme more susceptible to degradation by proteolytic enzymes (Ragan, 1976b; Crowder and Ragan, 1977). Thus solubilization of the flavoprotein fragment by both trypsin and chymotrypsin was severalfold faster after lipid depletion. In general the overall pattern of polypeptide degradation was similar to that observed with the native enzyme, except that all changes occurred faster-in particular, the rates of degradation of the 75,000- and 30,000-molecular-weight polypeptides.


It would be expected that removal of phospholipids would make the polypeptides that are normally in contact with the phospholipid bilayer more accessible to labeling with hydrophilic probes such as those described earlier. Indeed this was the case and certain low-molecular-weight polypeptides were labeled to a greater extent by lactoperoxidase-catalyzed iodination when phosphatidylcholine and phosphatidylethanolamine had been removed by treatment of the enzyme with cholate (Ragan, 1976b). However, of those subunits that were unlabeled in the native enzyme, i.e., the flavoprotein fragment subunits, the 30,000-molecular-weight subunit of the iron-protein fragment and the 23,500-molecular-weight subunit, only the latter was labeled in the lipid-depleted state (Ragan, 1976b). Thus the inaccessibility of the former groups is caused by masking by other proteins, not by phospholipids.

Phospholipid depletion also had marked effects on the labeling pattern obtained with iodonaphthylazide (F. G. P. Earley and C. 1. Ragan, unpublished observations). After removal of phosphatidylcholine and phosphatidylethanolamine from the enzyme by cholate (Section 4.3.2), incorporation of label into three polypeptides was increased while incorporation into all other proteins was unaffected. Figure 12 is an autoradiograph of an SDS-slab gel,

FIGURE 12. The effects of phospholipid depletion on

Origin

49 42

the labeling of NADH dehydrogenase subunits with 20 ['25I]iodonaphthylazide. Complex I was labeled and analyzed by discontinuous SDS-gel electrophoresis. Tracks (I) and (2) are autoradiographs of complex I and lipid-depleted complex I, respectively.

Track

2

298 C. I. Ragan

which shows increased labeling of subunits of molecular weights 49,000, 42,000, and 20,000 in the lipid-depleted enzyme. The converse experiment in which the phospholipid was increased showed no effects.

The interpretation of these results depends on an understanding of the mode of action of iodonaphthylazide. It is possible that this compound can label hydrophobic regions of proteins directly, even in the absence of phospholipids, and this could account for some of the labeling observed. If, on the other hand, iodon~phthylazide exists preferentially in the lipid phase, then lipid depletion would have the effect of concentrating the reagent in the lipid that still remained. If this lipid was randomly distributed over the enzyme then at least over a certain range of depletion, no change in the labeling pattern would be expected. The increased labeling of certain subunits implies that the residual lipid, which is cardiolipin, is not randomly distributed but is specifically associated with certain polypeptides. As the concentration of iodonapthylazide in the cardiolipin is increased by depletion of other lipids, these proteins become more heavily labeled following photoactivation. Thus it seems that three subunits, one of which is a major subunit of the iron-protein fragment may be binding sites for cardiolipin.

5. ORGANIZATION OF NADH DEHYDROGENASE IN THE MEMBRANE

5.1. Transmembranous Organization

Purified NADH dehydrogenase in the form of complex I can be incorporated into the membrane of phospholipid vesicles where it catalyzes proton translocation linked to oxidation of NADH by ubiquinone (Ragan and Hinkle, 1975). Therefore, just as in the mitochondrial membrane, NADH dehydrogenase is functionally trans membranous (Lawford and Garland, 1972), and the purified enzyme retains all the protein constituents required for proton transport. The structural basis for redox-linked proton movements is still debated. The simplest model for NADH dehydrogenase is that of Lawford and Garland (J 972), who proposed that the enzyme is organized in a "loop" (Mitchell, 1968) consisting of a hydrogen-carrying arm (FMN) and an electron-carrying arm (one or more iron-sulfur centers) as shown in Figure 13. The location in the membrane of the various carriers is not known, although it has been shown by the inhibitory effects of nonpermeant iron chelators that electron transport can be blocked from both the C and M sides of the membrane (Harmon and Crane, 1974, 1976).

The distribution of NADH dehydrogenase polypeptides has been studied by labeling exposed proteins on the surface of mitochondria or inverted sub-


FIGURE 13. Possible electron and hydrogen pathways across the mitochondrial membrane from NADH to ubiquinone (UQ).

FMN

Fe/S

299

NADH + H+

mitochondrial particles with the nonpenetrating reagent diazobenzenesulfonate, and by lactoperoxidase-catalyzed iodination. The labeled enzyme was then separated from other proteins by immunoprecipitation (Smith and Ragan, 1978, 1980). The results obtained with both labels are shown in Figure 14. Some differences between the labeling patterns obtained with the two probes were found. These are attributed to their different chemical specificities. Three polypeptides appear to be transmembranous-those of molecular weights 75,000,49,000, and 30,000. The latter is not the iron-protein fragment subunit but the polypeptide which comigrates with it in certain gel systems (Section 2.3). A subunit of molecular weight 26,000 is exposed only on the M side of the membrane, whereas a subunit of molecular weight 22,000 is exclusively labeled on the C side. In addition there are polypeptides of molecular weights, 15,500 and 8,000 labeled from either side but since there is more than one kind of subunit of these molecular weights, it cannot be concluded that they are transmembranous.

The transmembranous organization of the two larger subunits of the ironprotein fragment may be responsible for electron transfer across the membrane according to the model of Figure 13. Certainly, the observation that several polypeptides are exposed on either side of the membrane would lend support to proposals for the organization of the oxidoreduction pathway into a "loop."

Comparing the patterns obtained by labeling the isolated enzyme with those of Figures 4 and 5, it is obvious that several polypeptides are exposed in the isolated enzyme but not when the enzyme is incorporated into the membrane. Isolated NADH dehydrogenase has perhaps only enough phospholipid to form a single shell around the protein (Heron et ai., 1977), and polypeptides that are normally facing the hydrophobic interior of the membrane may be exposed enough in the isolated enzyme to become labeled. A large number of the lower-molecular-weight hydrophobic polypeptides fall into this group

300

o

30

49

30 22 ,.I

49

49 26 "J

30 ....

8

"

8

20 40 60 80 100 Slice no. from origin

C. I. Ragan

FIGURE 14. Transmembrane organization of NADH dehydrogenase subunits. Mitochondria (a) or inverted submitochondrial particles (b) were labeled with diazobenzene-["S]sulfonate or by lactoperoxidase-catalyzed radioiodination as indicated. After solubilization of the membrane with detergents, NADH dehydrogenase was purified by immunoprecipitation and analyzed by discontinuous SDS-gel electrophoresis.

(Table V), which is consistent with their external location in the enzyme. Indeed, it is largely the same group of subunits whose accessibility is further increased after removal of phospholipids from the isolated enzyme (Section 4.4.1 ).

5.2. Lateral Organization

Purified NADH dehydrogenase appears to react with ubiquinol-cytochrome c oxidoreductase in 1 : 1 molar ratio (based on the FMN-to-cytochrome C j ratio) (Hatefi et ai., 1962b; Fowler and Richardson, 1963; Ragan and Heron, 1978). The binary complex catalyzes NADH-cytochrome C oxi-

NADH Debydrogenase Structure 301

Table V Sidedness and Accessibility of NADH Dehydrogenase Subunitsa

Condition 10-3 X mol wt

Exposed on either side of the membrane (i.e., transmembranous) 75 49 33

Exposed on matrix side only 26 15.5b Sb Exposed on cytoplasmic side only 22 15.5 b Sb Exposed only in the isolated enzyme 39 25 20.5 IS 16.5 Inaccessible in the intact enzyme 53' 30 27' 23.5 15.5'

'Results are based on radioiodination and diazobenzene-["Slsulfonate labeling. Subunit labeling patterns were determined from one-dimensional gels only; therefore subunits of similar molecular masses were not resolved.

'These two subunits may be trans membranous. but because there is more than one subunit of these molecular weights it is not yet possible to decide.

'These are the flavoprotein fragment subunits.

doreductase via endogenous ubiquinone. In the membrane the association is transient allowing any dehydrogenase molecule to reduce any ubiquinone or cytochrome c reductase molecules at random (Kroger and Klingenberg, 1973a,b; Heron et al., 1978, 1979a).

The specific association of the two respiratory complexes obviously implies some kind of recognition between the two, and one or more of the NADH dehydrogenase polypeptides may have specific affinity for sites on the cytochrome c reductase molecule. At present there is no information on the nature of the binding or which parts of the molecule are involved. Any future proposals will have to consider the possibility that NADH dehydrogenase may be oligomeric (Section 1.3) and that ubiquinol-cytochome c oxidoreductase may be a dimer (De Vries et al.. 1979).

6. CONCLUSION

6.1. A Model of NADH Dehydrogenase Structure

The most striking feature of the enzyme is the number of subunits it contains. Because of this I have tried to generalize as much as possible about the effects of the various structural probes described above; rather than list their reactions with each individual polypeptide. The main generalization has been to divide the polypeptides up into hydrophobic or hydrophilic groups depending on their response to chaotropic reagents. If this has seemed arbitrary, I will now try to justify my reasons.

Often, the differences in catalytic properties between the flavoprotein

302 C. I. Ragan

fragment and the intact dehydrogenase have been emphasized. Perhaps it is the similarities that should receive the attention. It is surprising how little the properties change when the flavoprotein fragment is removed from a deeply buried membrane location to solution in water. No other respiratory enzyme has a hydrophilic catalytic unit within its structure, and I would like to propose that the primary purpose of most of the other polypeptides is to maintain a hydrophilic environment for this fragment in the membrane by surrounding it completely and preventing contact with the phospholipid bilayer. I have no idea as to why this is required, but assuming it is required leads to a simple explanation of why the enzyme is so large. The molecular weight of the flavoprotein fragment, calculated from its subunit molecular weights, is 95,000 (Table I). The average molecular weight of all the other enzyme subunits is about 25,000. Assuming that the flavoprotein fragment is spherical, it can be calculated that its surface can be covered by 24 spherical proteins of molecular weight 25,000. This number is surprisingly insensitive to variations in the assumed molecular weights. For example, varying the molecular weight of the surrounding proteins between 20,000 and 30,000 varies the number for covering the flavoprotein fragment from 26 to 22. Similarly, varying the molecular weight of the flavoprotein fragment from 80,000 to 120,000 requires 22-26 covering subunits of molecular weight 25,000. These numbers are remarkably close to the actual number of subunits other than those of the flavoprotein fragment, which is at least 23. The sum of the molecular weights of all the identified subunits is approximately 700,000 (Table I). This summation assumes that only one copy of each is present per molecule of NADH dehydrogenase, but the number cannot be much greater than this as the molecular weight based on FMN content is similar. Thus the agreement between theory and practice is quite striking.

I have not included the iron-protein fragment in the part of the enzyme that needs to be protected from the lipid environment because the two largest subunits are quite exposed in the enzyme. Thus the iron-protein fragment may have an intermediate role in which it conducts electrons from within the protected core to the exterior. It should also be remembered that not all the polypeptides designated as involved in shielding are without a catalytic function, as iron-sulfur centers are also associated with the hydrophobic residue from chaotropic resolution. However, the number of such iron-containing subunits must be small compared with total.

This simple model (Figure 15) is compatible with all the experimental work described in preceding sections. The polypeptides proposed to be involved in shielding the flavoprotein core have many general properties in common, e.g., low molecular weight, high isoelectric point, and hydrophobic nature, and, in addition, are located on the surface of the enzyme facing the hydrophobic region of the phospholipid bilayer. It is the polypeptides that do not fall into


Matrix side

- --"""'--1 Zl> ----n;:;:~;;:;t;~J_--r Iron-protein ~ fragment

Flavoprotein fragment

" FMN

Cytoplasmic side

__ -l--"-__ UQ

303

FIGURE 15. Organization of NADH dehydrogenase in the membrane. The main features of the proposed model are shown. Also indicated are possible hydrogen, electron, and proton pathways for the operation of NADH dehydrogenase as a linear redox loop with a H+ j2e- ratio of 2.

this general category, such as the large transmembranous subunits of the ironprotein fragment that may have a more specialized catalytic function.

6.2. Prospects

The model proposed for NADH dehydrogenase suggests that the enzyme may be rather less complex than is immediately apparent. Most of the constituent polypeptides have been assigned a noncatalytic role in which they merely provide the correct environment for the catalytic subunits. Several of the latter have been identified, e.g., the three subunits of the flavoprotein fragment and the two larger transmembranous subunits of the iron-protein fraction. More information on these proteins, even in isolation, would be valuable.

Conventional methods of mapping the structure of multisubunit enzymes, such as the use of cross-linking reagents, will probably be of limited value. Besides the enormous difficulty of analyzing the products, the subunits most likely to be cross-linked would be those on the surface of the enzyme for which there is probably no catalytic role. Probing the internal catalytic sites is clearly difficult. Neither hydrophobic nor hydrophilic reagents are likely to react to any great extent, and a study of the solubilized fragments, although less desirable, may be the only way.

304 C. I. Ragan

ACKNOWLEDGMENTS

The work from this laboratory was mainly undertaken by Sue Crowder, Fergus Earley, Christine Heron, Stuart Smith, and Veronica Poore. Financial support came from the Science Research Council.

7. REFERENCES

Albracht, S. P. J., Dooijewaard, G., Leeuwerik, F. J., and Van Swol, B., 1977, Epr signals of NADH : Q oxidoreductase-Shape and intensity, Biochim. Biophys. Acta 459:300-317.

Albracht, S. P. J., Leeuwerik, F. J., and Van Swol, B., 1979, The stoichiometry of the ironsulfur clusters la, Ib and 2 of NADH : Q oxidoreductase as present in beef-heart submitochondrial particles, FEBS Lett. 104:197-200.

Baugh, R. F., and King, T. E., 1972, Purification, properties and reconstitutive activity of a DPN H dehydrogenase, Biochem. Biophys. Res. Commun. 49: 1165-1173.

Baugh, R. F., and King, T. E., 1974, A mitochondrial membrane bound NADH dehydrogenase and its reconstitutive activity, Biochim. Biophys. Acta Libr. 13: 143-158.

Bercovici, T., and Gitler, c., 1978, 5-[I25I]-Iodonapthhylazide, a reagent to determine the penetration of proteins into the lipid bilayer of biological membranes, Biochemistry 17:1484-1489.

Cremona, T., and Kearney, E. B., 1964, Studies on the respiratory chain-linked reduced nicotinamide adenine dinucleotide dehydrogenase. VI. Further purification and properties of the enzyme from beef heart, J. Bioi. Chem. 239:2320-2334.

Cremona, T., Kearney, E. B., Villavicencio, M., and Singer, T. P., 1963, Studies on the respiratory chain-linked DPNH dehydrogenase. V. Transformation of DPNH dehydrogenase to DPN H-cytochrome reductase and diaphorase under the influence of heat, proteolytic enzymes, and urea, Biochem. Z. 338:407-442.

Crowder, S. E., and Ragan, C. I., 1977, Effects of proteolytic digestion by chymotrypsin on the structure and catalytic properties of reduced nicotinamide-adenine dinucleotide-ubiquinone oxidoreductase from bovine heart mitochondria, Biochem. J. 165:295-301.

Davis, K. A., and Hatefi, Y., 1969, Kinetics of the resolution of Complex I (reduced diphosphopyridine nucleotide-coenzyme Q reductase) of the mitochondrial electron transport system by chaotropic agents, Biochemistry 8:3355-3361.

De Vries, S., Albracht, S. P. J., and Leeuwerik, F. J., 1979, The multiplicity and stoichiometry of the prosthetic groups in QH, : cytochrome c oxidoreductase as studied by epr, Biochim. Biophys. Acta 546:316-333.

Dooijewaard, G., and Slater, E. c., 1976a, Steady-state kinetics of high molecular weight (typeI) NADH dehydrogenase, Biochim. Biophys. Acta 440:1-15.

Dooijewaard, G., and Slater, E. c., 1976b, Steady-state kinetics of low molecular weight (typeII) NADH dehydrogenase, Biochim. Biophys. Acta 440:16-35.

Dooijewaard, G., De Bruin, G. J. M., Van Dijk, P. J., and Slater, E. C., 1978a, Characterization of type-I NADH dehydrogenase-Polydispersity, molecular weight and polypeptide composition, Biochim. Biophys. Acta 501:458-469.

Dooijewaard, G., Slater, E. c., Van Dijk, P. J., and De Bruin, G. J. M., 1978b, Chaotropic resolution of high molecular weight (type I) NADH dehydrogenase, and reassociation of flavin-rich (type II) and flavin-poor subunits, Biochim. Biophys. Acta 503:405-424.

Eytan, G. D., Carroll, R. c., Schatz, G., and Racker, E., 1975, Arrangement of the subunits in


solubilized and membrane-bound cytochrome c oxidase from bovine heart, J. BioI. Chem. 250:8298-8603.

Fleischer, S., Klouwen, H., and Brierley, G., 1961, Studies on the electron transfer system. XXXVIII. Lipid composition of purified enzyme preparations derived from beef heart mitochondria, J. BioI. Chem. 236:2936-294i.

Fleischer, S., Rouser, G., Fleischer, 8., Casu, A., and Kritchevsky, G., 1967, Lipid composition of mitochondria from bovine heart, liver and kidney, J. Lipid Res. 8:170-180.

Fowler, L. R., and Richardson, S. H., 1963, Studies on the electron transfer system. L. On the mechanism of reconstitution of the mitochondrial electron transfer system, J. BioI. Chem. 278:456-463.

Galante, Y. M., and Hatefi, Y., 1979, Purification and molecular and enzymic properties of mitochondrial NADH dehydrogenase, Arch. Biochem. Biophys. 192:559-568.

Gatley, S. J., and Sherratt, H. S. A., 1976, The effects of diphenyleneiodonium on mitochondrial reactions-Relation of binding of diphenylene [1251] iodonium to mitochondria to the extent of inhibition of oxygen uptake, Biochem. J. 158:307-315.

Guarnieri, M., Stechmiller, 8., and Lehninger, A. L., 1971, Use of an antibody to study the location of cardiolipin in mitochondrial membranes, J. BioI. Chem. 246:7526-7532.

Gutman, M., Singer, T. P., and Casida, J. E., 1970, Studies on the respiratory chain-linked reduced nicotinamide adenine dinucleotide dehydrogenase. XVII. Reaction sites of piericidin A and rotenone, J. BioI. Chem. 245: 1992-1997.

Hare, J. F., and Crane, F. L., 1974, Proteins of mitochondrial cristae, Sub-Cell. Biochem. 3:1-25.

Harmon, H. J., and Crane, F. L., 1974, Topographical definition of new sites on the mitochondrial electron transport chain, Biochem. Biophys. Res. Commun. 59:326-333.

Harmon, H. J., and Crane, F. L., 1976, Inhibition of mitochondrial electron transport by hydrophobic metal chela tors-determination of dehydrogenase topography, Biochim. Biophys. Acta 440:45-58.

Hatefi, Y., and Hanstein, W. G., 1969, Solubilization of particulate proteins and non-electrolytes by chaotropic agents, Proc. Natl. Acad. Sci. U.S.A. 62:1129-1136.

Hatefi, Y., and Stempel, K. E., 1967, Resolution of Complex I (DPNH-coenzyme Q reductase) of the mitochondrial electron transfer system, Biochem. Biophys. Res. Commun. 26:301-308.

Hatefi, Y., and Stempel, K. E., 1969, Isolation and enzymatic properties of the mitochondrial reduced diphosphopyridine nucleotide dehydrogenase, J. BioI. Chem. 244:2350-2357.

Hatefi, Y., and Stiggall, D. L., 1976, Metal-containing flavoprotein dehydrogenases, in: The Enzymes (P. D. Boyer, H. Lardy, and K. Myrback, eds.), 3rd ed., pp. 175-297, Academic Press, New York.

Hatefi, Y., Haavik, A. G., and Jurtshuk, P., 1960, Studies on the electron transport system. XXXII. Reduction of coenzyme Q by DPNH, Biochem. Biophys. Res. Commun. 3:281-286.

Hatefi, Y., Haavik, A. G., and Griffiths, D. E., 1962a, Studies on the electron transfer system. XL. Preparation and properties of mitochondrial DPNH-coenzymc Q reductase, J. BioI. Chem. 237: 1676-1680.

Hatefi, Y., Haavik, A. G., Fowler, L. R., and Griffiths, D. E., 1962b, Studies on the electron transfer system. XLII. Reconstitution of the electron transfer system, J. BioI. Chem. 237:2661-2669.

Hatefi, Y., Galante, Y. M., Stiggall, D. L., and Ragan, C. I., 1979, Proteins, polypeptides, prosthetic groups and enzymic properties of Complexes I, II, III, IV and V of the mitochondrial oxidative phosphorylation system, Methods Enzymol. 56:562-602.

Heron, C, Corina, D., and Ragan, C I., 1977, The phospholipid annulus of mitochondrial

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NADH-ubiquinone reductase-A dual phospholipid requirement for enzyme activity, FEBS Lett. 79:399-403.

Heron, C., Ragan, C. I., and Trumpower, B. L., 1978, The interaction between mitochondrial NADH-ubiquinone oxidoreductase and ubiquinol cytochrome c oxidoreductase-Restoration of ubiquinone-pool behaviour, Biochern. J. 174:791-800.

Heron, C., Gore, M. G., and Ragan, C. I., 1979a, The effects of lipid phase transitions on the interaction of mitochondrial NADH-ubiquinone oxidoreductase with ubiquinol-cytochrome c oxidoreductase, Biochern. J. 178:415-426.

Heron, c., Smith, S., and Ragan, C. I., 1979b, An analysis of the polypeptide composition of bovine heart mitochondrial NADH-ubiquinone oxidoreductase by two-dimensional polyacrylamide-gel electrophoresis, Biochern. J. 181:435-443.

Higgins, H. G., and Harrington, K. 1., 1959, Reaction of amino acids and proteins with diazonium compounds. II. Spectra of protein deriv'atives, Arch. Biochern. Biophys. 85:409-425.

Hinz, H-l., and Sturtevant, 1. M., 1972, Calorimetric studies of dilute aqueous suspensions of bilayers formed from synthetic L-a-lecithins, J. BioI. Chern. 247:6071-6074.

Holland, P. C., Clark, M. G., Bloxham, D. P., and Lardy, H., 1973, Mechanism of action of the hypoglycemic agent diphenyleneiodonium, J. BioI. Chern. 248:6050-6056.

Howard, A. N., and Wild, F., 1957, The reactions of diazonium compounds with amino acids and proteins, Biochern. J. 65:651-659.

Kauzmann, W., 1959, Some factors in the interpretation of protein denaturation, Adv. Protein Chern. 14:1-63.

Klotz, I. M., and Farnham, S. B., 1968, Stability of an amide-hydrogen bond in an apolar environment, Biochemistry 7:3879-3882.

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Lawford, H. G., and Garland, P. B., 1972, Proton translocation coupled to quinone reduction by reduced nicotinamide-adenine dinucleotide in rat liver and ox heart mitochondria, Biochem. J. 130: 1 029-1 044.

Maccecchini, M-L., Rudin, Y., Blobel, G., and Schatz, G., 1979, Import of proteins into mitochondria: Precursor forms of the extramitochondrially made F,-ATPase subunits in yeast, Proc. Natl. Acad. Sci. U.S.A. 76:233-347.

Machinist, 1. M., and Singer, T. P., 1965, Studies on the respiratory chain-linked reduced nicotinamide adenine dinucleotide dehydrogenase. IX. Reactions with coenzyme Q, J. BioI. Chem. 240:3182-3190.

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Ragan, C. I., 1978, The role of phospholipids in the reduction of ubiquinone analogs by the mitochondrial reduced nicotinamide-adenine dinucleotide-ubiquinone oxidoreductase complex, Biochem. J. 172:539-547.

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Ragan, C. I., and Racker, E., 1973, Resolution and reconstitution of the mitochondrial electron transport system. IV. The reconstitution of rotenone-sensitive NADH-ubiquinone reductase from NADH dehydrogenase and phospholipids, J. Bioi. Chem. 248:6876-6884.

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Chapter 7

Structure-Function Relationships of Micrococcus lysodeikticus Membranes: A Bacterial Membrane Model System

Milton R. J. Salton Department of Microbiology New York University School of Medicine New York, N.Y. 10016

1. INTRODUCTION

In the course of his studies on the antibacterial and bacteriolytic properties of various tissues and secretions, Fleming (1922) noticed the complete lysis of some of the colonies of a yellow-pigmented organism growing in the vicinity of the mucus. Fleming (1922) isolated the micrococcus species and gave the name lysozyme to the lytic substance. Glynn (1968) records the history of the origin of the name given to this lysozyme-sensitive organism as follows: "The name Micrococcus lysodeikticus for the yellow bacteria we owe to Sir Almroth Wright who would never use a simple word in English or even Latin if he could invent a more complicated one in Greek." The species designation for this gram-positive organism is indeed a hybrid epithet meaning "lysis indicator" (lysodeikticus). Fleming never described the organism in the traditional style of the bacteriologist, probably because he was deliberately searching for natural antibiotics and was much more interested in bacterial lysis and the antibacterial properties of lysozyme and later penicillin. As a result the use of lysodeikticus as a species of Micrococcus has had a spotty history and, after having been mentioned transiently in Bergy's Manual of Determinative Bacteriology. it is no longer recorded in this August catalog. The high susceptibility to lysis by lysozymes from various tissues and secretions reported by Fleming (1922) was undoubtedly the principal feature of this bacterium to attract attention and has probably resulted in the persistence of its name. The general

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310 Milton R. J. Salton

morphological, biochemical, and physiological properties of this organism certainly lump it with other micrococci and related Sarcina species. Micrococcus luteus is widely used as the officially described species.

It has been the author's experience that the strains (e.g., NCTC 2665) originating from Fleming's isolate are probably the most sensitive of all the gram-positive bacteria to hen egg-white lysozyme. When subcellular biochemistry became the vogue, the high lysozyme sensitivity of M. Iysodeikticus strain NCTC 2665 was a very attractive feature and greatly facilitated the biochemical attack on the nature of both bacterial cell walls and membranes. Indeed, the chemical nature of the unique bacterial substrate for lysozyme intrigued such eminent scientists as the late Ernst Chain (Epstein and Chain, 1940) and Karl Meyer (Meyer et al., 1936), who made some of the earliest observations on the properties of the products of lysozyme action. Subsequent studies demonstrated that the rigid cell-wall structure of M. Iysodeikticus could be completely degraded to soluble products (Salton, 1952, 1956a), and the investigation of the nature of the digested wall fragments gave the first clues that a disaccharide repeating unit (Salton, 1956a) was an important feature of the glycan backbone of what is now universally known as peptidoglycan, mucopeptide, or murein. The remarkable progress that has been made in the ensuing 20-30 years has resulted in the elucidation of the primary structures of bacterial peptidoglycans, as well as their recognition as characteristic prokaryotic wall polymers with a unique set of enzymes involved in their biosynthesis and the exciting discoveries that many of the enzymes so involved are targets for specific antibiotic inhibitors (Ghuysen and Shockman, 1973; Blumberg and Strominger, 1974; Ghuysen et al., 1979).

Early investigations of cell-wall structures, at least those of a number of gram-positive bacteria (Salton, 1956b), suggested that they were not the sites of major biochemical and transport activities of the cell. Consequently attention began to be focused on the biochemical functions of the membranes and the observations that the cell walls of certain gram-positive bacteria could be selectively degraded to relatively small-molecular-weight products paved the way for Weibull's (1953a,b) classic studies of protoplast formation and subsequent isolation of bacterial membranes. Osmotically stabilized protoplasts selectively deprived of their outer cell walls retained most of the biosynthetic and transport capabilities of intact bacterial cells (McQuillen, 1960), thereby indicating that the plasma or protoplast membrane was the major permeability and functional barrier of the cell. The localization of cytochromes in the protoplast "ghosts" (Weibull, 1953b) clearly established the membrane as the site of the electron-transport system of the bacterial cell. It is now widely recognized that the bacterial plasma membrane performs a multiplicity of functions in the prokaryotic cell, in contrast to the eukaryotic cell, where the various functions are compartmentalized in the various membranous organelles such

A Bacterial Membrane Model System 311

as the mitochondria, Golgi, lysosomes, and endoplasmic reticulum (Stanier, 1970; Salton, 1976; Salton and Owen, 1976). Thus the plasma membranes are the site of respiratory chain and energized transport processes, including the coupling factor F]-ATPases and an array of enzymes involved in lipid, carotenoid, and menaquinone biosynthesis (Salton, 1976), as well as being the site of the membrane stages (Ghuysen and Shockman, 1973) of wall peptidoglycan synthesis and assembly and teichoic acid biosynthesis (Baddiley, 1972). Although many of these biochemical functions have been demonstrated with isolated membrane fractions devoid of cell-wall components, some important functions are better retained in wall-membrane complexes from disrupted cells of the type used in peptidoglycan synthesis by Mirelman and his colleagues (Mirelman and Sharon, 1972; Mirelman et al., 1974). However, to establish the compartmentalization of cellular functions in the membranes and for the investigation of the molecular architecture and asymmetry of membrane structures, the isolation of wall-free membranes and the ability to prepare intact protoplasts has had great advantages. For these reasons M. Iysodeikticus has been one of the organisms of choice for investigating the structure-function relationships of a bacterial plasma membrane.

At the time we initiated our studies with M. Iysodeikticus membranes, comparable studies with a gram-negative organism such as Escherichia coli would have been very difficult. Similar investigations on the compartmentalization of enzymes and energized transport functions in gram-negative bacteria had to await the development of suitable techniques for separating the outer membranes of the gram-negative envelope structure from the inner or plasma membranes. This formidable task was achieved by Schnaitman (1970) for E. coli and by Osborn et al. (1972a,b) for Salmonella typhimurium following the key observations of Birdsell and Cota-Robles (1967) and Miura and Mizushima (1969). Despite the success in the isolation of inner and outer membranes of gram-negative organisms, studies of the molecular architecture of the membrane faces are still hampered by the inability to obtain true protoplasts to determine membrane sidedness. The only gram-negative organism that has given authentic protoplasts is the marine bacterium investigated by MacLeod's group (De Voe et al., 1970), although there is one report of the preparation of protoplasts from E. coli by Weiss (1976). In contrast, the application of similar approaches to those used with M. Iysodeikticus has worked successfully with the isolated plasma membrane vesicles of E. coli (Owen and Kaback, 1978). The difficult problem of establishing the sided ness of the outer membrane of gram-negative bacteria is still unresolved, and the nature of the components of the inner (plasma) membrane exposed on its outer face (peri plasmic side) has yet to be extended from the results with the right-side-out plasma membrane vesicles. The prospects of being able to resolve these problems now appear to be feasible by antibody absorption experiments using spheroplasts (M. L. Per-


ille Collins, D. E. Mallon, and R. A. Niederman, unpublished results) of Rhodo pseudomonas sphaeroides. Collins et al. (1979) have been able to absorb out antibodies to the components of the outer face of the inner (plasma) membrane by using lysozyme-EDTA spheroplasts, which permit the passage of immunoglobulins through the outer membrane structure. Such studies together with further probing of the outer membrane asymmetry will eventually bring the state of our knowledge of the membrane architecture of gram-negative bacteria closer to that achievable with a gram-positive organism such as M. lysodeikticus.

2. ULTRASTRUCTURE OF BACTERIAL MEMBRANES

Bacterial cell-surface structures have been studied by all the basic electron-microscopy procedures, including thin sectioning, metal shadowing, negative staining, freeze fracturing, and freeze etching. Immunoelectron microscopy has added a further dimension in specifically identifying surface components by using monospecific antibodies conjugated to particles such as ferritin, which can be identified by using one of the basic techniques of electron microscopy, especially thin sectioning, negative staining, and freeze etching (Shands, 1965; Bayer, 1974; Oppenheim and Nachbar, 1977). We now have an excellent knowledge of the topography of the bacterial surface and the ultrastructural features of the walls and plasma membranes of gram-positive organisms, as well as the complex envelope structures of gram-negative bacteria, with their outer and inner (plasma) membrane layers (Glauert and Thornley, 1969; Costerton et al., 1974; DiRienzo and Inouye, 1978).

The surface profiles of M. lysodeikticus conform to those seen in thin sections of most gram-positive bacteria (e.g., other micrococci, staphylococci, and streptococci). The strains of M. lysodeikticus and M. luteus commonly used are nonencapsulated and possess a relatively thick cell wall (40-50 nm) as the outermost structure (Salton and Chapman, 1962). Immediately below the cell wall and closely apposed to it is the plasma or cytoplasmic membrane exhibiting the electron-dense layering of the double-track structure typical of all membrane profiles. As in other gram-positive organisms, thin sections of M. lysodeikticus fixed by the Kellenberger and Ryter (1958) method exhibit the invaginations of the plasma membrane containing the mesosomal membrane vesicles (Salton and Chapman, 1962). When stable protoplasts of M. lysodeikticus are formed by lysozyme digestion of the cell wall in the presence of an osmotic stabilizer (e.g., 0.8 M sucrose), the thin sections show a single membrane structure as the surface component of the intact protoplast (Salton and Chapman, 1962). Because of the sensitivity of its cell wall to complete digestion into soluble products by hen's egg white lysozyme, no residual wall is


detectable on the protoplast surface. The ability to remove the cell wall selectively and to expose the outer face of the underlying membranes of the protoplast surface of M. lysodeikticus has been of immense value in establishing the antigenic architecture and asymmetry of the plasma membranes of this bacterium. It is this feature that has made the organism such a valuable model in the investigation of a prokaryotic, multifunctional membrane system; various aspects of this feature will be discussed in further detail in subsequent sections. It should also be emphasized that similar potentials for bacterial membrane studies also exist for other gram-positive organisms, such as Bacillus megaterium strain KM, which is also very sensitive to lysozyme, Staphylococcus aureus with its sensitivity to the lysostaphin enzyme, and streptococci with the phage muramidase (see review by Salton, 1976). The staphylococcal walldegrading activity of lysostaphin (an extracellular lytic factor produced by a species of Staphylococcus) is due to an endopeptidase which splits the glycine bridge of Staphylococcus aureus peptidoglycan (Browder et al., 1965). Although lysostaphin preparations also contain an endo-N-acetylglucosaminidase active on the glycan backbone it does not lyse Staphylococcus aureus. In contrast, the walls of certain other gram-positive bacteria are only partially degraded by lysozyme (or other muramidases), and the protoplasts may be encased in a residual wall structure. With most gram-negative bacteria, lysozyme-EDT A treatment results in the loss of the peptidoglycan layer and a portion of the lipopolysaccharide, leaving the bulk of the outer membrane structure to yield a spheroplast, rather than a protoplast. Thin sections of spheroplasts of gram-negative bacteria clearly show the double membrane envelope (outer + inner membranes) with the loss of the metal-stained, electron-dense layer of peptidoglycan (Murray, 1968).

A rather different aspect of the membrane topography and ultrastructure is seen in negatively stained preparations examined under the electron microscope. Although of less value in establishing the architecture of membranes, the negative-staining technique has been especially useful in monitoring the preparation of membranes and distinguishing between plasma and mesosomal membrane vesicles of gram-positive organisms such as M. lysodeikticus (Salton, 1971, 1976) and Staphylococcus aureus (Theodore et al., 1971). The typical appearance of membrane sheets and fragments isolated from M. lysodeikticus is shown in the preparation negatively stained with ammonium molybdate presented in Figure 1. The distinction between the plasma membranes and the mesosomal vesicles is readily seen by comparing the appearance of isolated membranes in Figure I with that of the negatively stained preparations of isolated mesosomal membrane fractions presented in Figures 2 and 3. Moreover, the plasma membrane fragments in the crude mesosome fraction shown in Figure 2 can be readily distinguished from the vesicular structures. The preparation of mesosomal vesicles illustrated in Figure 2 also shows that the vesicles


A

FIGURE 1. Typical appearance of isolated plasma membrane fractions of M. lysodeikticus negatively stained with 2% ammonium molybdate. (A) Electron micrograph showing a large membrane sheet and an abundance of uniform 10-nm particles identified as the F,-A TPase (see arrows) randomly distributed on the membranes. (X 120,000, reproduced at 90%) (B) Electron micrograph showing the F,-ATPase particles peripherally displayed on what appears to be insideout vesicle, as well as the random distribution on a larger membrane fragment. (X 150,000, reproduced at 90%)



can exhibit batch-to-batch differences in appearance, being more tubulovesicular in this fraction and predominantly spherical in the purified vesicles in Figure 3. The reasons for the difference in the morphology of the mesosomal vesicles is unknown. However, Burdett and Rogers (1970) have indicated that factors such as ionic strength of the suspending medium can influence the appearance of mesosomes as seen in intact cells of Bacillus licheniformis. The


FIGURE 2. Electron micrograph of a crude mesosome fraction from M. lysodeiklicus negatively stained with ammonium molybdate. Note the presence of vesicles of varying shapes and flat sheets of plasma membrane fragments. From Salton (1976) .

whole status and indeed the functions of the mesosomes are still very much open questions at the present time, and the possible origins and evolution of this puzzling membrane structure have been extensively reviewed in this series by Ghosh (1974).

The negative-staining procedure has also been valuable in studying the


FIGURE 3. The appearance of a purified fraction of mesosomes from M. lysodeikticus, negatively stained with ammonium molybdate. Note the absence of plasma membrane fragments and lO-nm particles in this preparation. (X 156,000, reproduced at 90%)

structure-function relationships in this membrane system. It can be seen from the negatively stained plasma membrane fractions illustrated in Figure 1 that the membrane is covered with an abundance of small uniform particles of approximately 10-nm diameter. These particles have been identified as the bacterial F1-ATPase components (Munoz et al., 1968; Nachbar and Salton,


1970a; Oppenheim and Salton, 1973). Neither ATPase activity nor the 10-nm particles were detectable when isolated mesosomal membrane fractions were examined (Oppenheim and Salton, 1973). Some larger particles or aggregates of a less uniform nature are also seen on the plasma membranes, but their functions have not yet been established. Although these components were retained on the membranes through the standard three to six consecutive washes for membrane isolation (Salton, 1967, 1976), they were removed from the membranes by the low-ionic-strength or EDT A shock washes, or both, which release activities such as the FJ-ATPase, NADH dehydrogenase, and cardiolipin synthetase (Salton and Nachbar, 1970; DeSiervo and Salton, 1971). It is conceivable that these particles may be peripheral membrane aggregates, the sites of NADH dehydrogenase or cardiolipin synthetase activities, or both, associated with small membrane vesicular structures of about 30-nm diameter (Nachbar and Salton, 1970b; DeSiervo and Salton, 1971). The use of negative staining and electron microscopy in establishing the nature of the 10-nm FJ-ATPase particles will be discussed further in Section 3.6.1 devoted to the FJ coupling factor of the M. lysodeikticus membranes.

The development of the freeze-fracture/freeze-etching techniques has added much to our knowledge of the molecular organization of cell membranes of both eukaryotic and prokaryotic cells. Freeze-fracture studies have provided strong supportive evidence for the existence of substantial regions of bilayer lipid in cell membranes (Branton, 1966; Branton and Deamer, 1972). There is now general acceptance that the fracture plane of membrane structures occurs through the hydrophobic center, thus exhibiting internal convex and concave surfaces of the cleaved membranes. Indeed, the principal fracture plane seen in bacterial cells subjected to the freeze-fracture technique is that of the plasma membrane. Much of the work performed on bacteria has been reviewed in the elegant paper of Holt and Leadbetter (1969), and the current interpretation of the results obtained with both gram-positive and gram-negative organisms has been discussed in some detail by Salton and Owen (1976). The main conclusion from all these studies is that the principal fracture plane is through the hydrophobic center of the plasma membranes of both gram-positive and gram-negative bacteria, with the latter also showing a second fracture plane, albeit a much weaker one, which is probably through the plane of the outer membrane. In all instances the bacterial plasma membranes exhibit in common with membranes of other cell types the characteristic appearance of the convex fracture surface with a dense covering of particles (about 5-10 nm in diameter) and the corresponding concave fracture surface with fewer particles and depressions. The abundance of particles in the convex fracture face of the plasma membrane of intact protoplasts of M. lysodeikticus was shown by Salton (1971), and the concave surface showed the characteristics typical


of plasma membrane structures. Thus the membrane of M. Iysodeikticus as seen by the freeze-fracture technique exhibits a similar ultrastructure, and hence organization, to that of other cell membranes. We have not seen particles corresponding to the 10-nm F]-ATPase on freeze-fracture/freeze-etch preparations of cells and membranes of M. Iysodeikticus. Presumably these could only be readily visualized by examining inside-out plasma membrane vesicles by the electron-microscopic freeze-etch technique, to indicate the ATPase on the outer face of such vesicles.

The freeze-fracture/freeze-etch procedures are believed to preserve the cellular structures in as close a state to the native condition of the cell as possible. It has therefore been something of a surprise that the abundance of mesosomal vesicles seen in fixed and sectioned cells was not seen when the same organisms were subjected to freeze-fracture studies. This led Fooke-Achterrath et al. (1974) to suggest that mesosomes seen by thin sectioning are "artificial technikosomes" and that the "real" mesosomes are smaller and much less abundant. As discussed by Salton and Owen (1976), other investigators have also observed the large discrepancy between the relative abundance of mesosomal vesicles in thin sections of fixed cells compared to the paucity of their detection in freeze-fractured cells. We have also been unable to detect mesosomal vesicles in freeze-fractured cells of M. Iysodeikticus. Higgins et al. (1975) have correlated the kinetics of appearance of mesosomes as detected by freeze fracturing of Streptococcus faecalis cells with the glutaraldehyde crosslinking capacity. All these observations add yet another chapter to the mystery of the origins and functions of the mesosomal vesicles in bacteria. Where freeze fracturing has been performed on isolated mesosomal vesicles, the fracture planes appear to be typical for membrane structures with apparently some particles visible on what is assumed to be the convex face (Owen and Freer, 1972). However, at the present time the ultrastructural studies with mesosomes are not really conclusive and should they turn out to be physiologically significant structures then further work on their ultrastructure by this and other techniques would be appropriate.

In summary, the two membrane components of Micrococcus Iysodeikticus that can be isolated as homogeneous preparations are the plasma membranes and the mesosomal vesicles, and the ultrastructure of the former as determined by the techniques of electron microscopy conforms to the general characteristics of other cell membranes. The plasma membranes of this organism have an abundance of 10-nm particles identifiable as the F]-ATPase, whereas the mesosomal vesicles are largely devoid of these particles. Although the mesosomal vesicles are distinctive entities of fixed cells and are amenable to isolation, their paucity in freeze-fractured cells and their origins and functional significance remain an enigma.


3. BIOCHEMICAL CHARACTERIZATION OF MICROCOCCUS LYSODEIKTICUS MEMBRANES

3.1. Enzyme Distribution

The main cellular compartments of a lysozyme-sensitive organism such as M. Iysodeikticus can be readily separated and characterized for studies of enzyme distribution and localization. These compartments are identifiable as the external cell-wall region, the plasma membrane, the mesosomal membrane vesicles and the cytoplasm or cytosol. Thus the lysozyme-digested, cell-wall compartment and any enzymes associated with this external region of the cell can be separated along with the released mesosomal vesicles from the protoplasts by low-speed centrifugation. In general, gram-positive bacteria do not appear to possess a periplasmic compartment containing the variety of enzymes and binding proteins characteristic of the peri plasmic region found in gramnegative organisms such as E. coli (Rosen and Heppel, 1973). However, an enzyme such as the penicillin-sensitive Do-carboxypeptidase of M. Iysodeikticus was found in the supernatant fraction following stable protoplast formation and removal from the protoplasting medium by centrifugation (Linder and Salton, 1975). Some externally located enzymes, perhaps loosely associated with the outer membrane surface, may therefore be found in the wall-digest fraction upon protoplasting. The protoplasts can be washed gently and then subjected to osmotic lysis to yield upon centrifugation (Salton, 1976) a crude membrane fraction and a cytoplasmic fraction. Early investigations in our laboratory (Nachbar and Salton, 1970a) showed that there was a clear-cut localization of certain activities in either the plasma membrane or in the cytoplasmic compartment and that some enzymes (e.g., catalase) partitioned between the crude membrane fraction and the cytoplasmic fraction. Carotenoids, phospholipids, menaquinones, cytochromes, various dehydrogenases, and ATPase were almost exclusively localized in the plasma membranes, and enzymes such as adenosine deaminase and transaminase (glutamate-oxalacetate) were largely of cytoplasmic origin (Salton, 1971; Salton and Nachbar, 1970). Simple washing of the membranes removed partitioning enzymes, such as catalase and polynucleotide phosphorylase (Nachbar and Salton, 1970a), without releasing any of the characteristic lipid or membrane markers (Salton, 1967, 1971). The release of the latter components of the membrane was regarded as indicative of membrane perturbation and fragmentation, and hence nonsedimentable at the normal gravitational forces (approximately 30,000g for 30-60 min) used to sediment the plasma membranes. The mesosomal membrane vesicles required higher gravitational forces for sedimentation in the centrifuge (Owen and Freer, 1972).

Gel'man and her colleagues (Gel'man et al., 1975) have also selected M.


Iysodeikticus for their studies of the organization of the respiratory chain in the membranes of this organism; their investigations have added significantly to our knowledge of the biochemistry of the bacterial membrane. In addition to the studies of Gel'man et al. (1975) and those in the author's laboratory, a number of investigators have selected M. Iysodeikticus for studying the biosynthesis of various cell-wall and membrane components; the cellular compartmentalization of various enzymes in this organism is summarized in Table I.

Table I Compartmentalization of Certain Enzyme Activities in Micrococcus lysodeikticus

Cytoplasmic enzymes

Membrane enzymes

Wall compartment'

Adenosine deaminase Transhydrogenase Glutamic-oxaloacetic transaminase Nonspecific esterase Lipase Hexokinase Adenylate kinase Isocitra te dehydrogenase Catalasea

Polyynucleotidephosphorylasea

NADH dehydrogenaseb

NADH dehydrogenase Succinate dehydrogenase Malate dehydrogenase Lactate dehydrogenase' ATPase CTP-phosphatidic acid, cytidyltransferase Cardiolipin synthetase Mannan synthetase Mannosyl-l-phosphorylundecaprenol synthetase Teichuronic acid synthetased

D-Alanine carboxypeptidase

'These two enzymes in particular partitioned between the cytoplasmic fraction and the crude, unwashed membranes, but they were readily removed by several washings of the membrane fractions with Tris buffer without any release of membrane enzymes.

"The existence of three enzymes with NADH dehydrogenase activity was reported by Owen and Salton (1977)~one was largely cytoplasmic, one distributed between cytoplasmic and membrane compartments, and one occurred largely in the membrane.

'Localization of lactate dehydrogenase in M. Iysodeikticus membranes was reported by Mitchell (1963) and Gel'man et al. (1975), but was not detected by zymogram staining of immunoprecipitates (Owen and Salton. 1975d) probably due to inactivation by Triton X-IOO and the requirement for reactivation with menadione (Tikhonova. 1974).

dparticulate enzymes involved in teichuronic acid biosynthesis. as studied by Anderson et al. (1972). 'The term wall compartment constituting the protoplasting supernatant fluid has been used in preference to peri plasmic for a gram-positive organism. Data from Linder and Salton (1975).


From an inspection of the activities found in the M. Iysodeikticus plasma membranes (and indeed other bacterial membranes) it is evident that these prokaryotic structures are biochemically multifunctional. Bacterial membranes possess the energizing systems for transport across the membrane, the electrontransport chain components, a number of enzymes involved in phospholipid, carotenoid, poly isoprenoid, and enzymes of the membrane stage of peptidoglycan synthesis. Particulate fractions indicating a membrane origin are also active in the synthesis of the mannosaminuronic acid polysaccharide polymer linked to the wall peptidoglycan of M. Iysodeikticus (Perkins, 1963; Anderson et at., 1972; Hase and Matsushima, 1977). Enzymes involved in the biosynthesis of the membrane succinylated lipomannan are also localized in the plasma membrane structures (Pless et aI., 1975; Powell et al., 1975; Owen and Salton, 1975a). It is of special interest to note that although the isolated mesosome vesicles are greatly enriched in the succinylated lipomannan, they do not appear to be the specific site of synthesis of this membrane polymer (Owen and Salton, 1975a). Indeed, only low levels of mannan synthetase activity as determined by the incorporation of [14C]mannose from GDp-[14C]mannose into mannan could be detected in the mesosomal membranes (Owen and Salton, 1975a). However, when purified [14C]mannosyl-l-phosphorylundecaprenol (a lipid intermediate in lipomannan synthesis) was used as a substrate, a dramatic increase was observed in the ability of mesosomal membranes to catalyze the incorporation of labeled [14C]mannose into mannan (Owen and Salton, 1975a). It would appear that the inability of the mesosomes to synthesize lipomannan was due to the failure to synthesize the lipid intermediate. Whether the incorporation represents addition of terminal sugar residues or complete polymer synthesis has yet to be determined. Thus, despite a four- to fivefold enrichment of the lipomannan in the mesosomal membrane fraction, the enzymes involved in its biosynthesis appear to be largely localized in the plasma membrane fraction of M. Iysodeikticus and not in the mesosomal structures.

As with many membrane systems very few enzymes from bacterial membranes including those of M. lysodeikticus have been purified to anything like protein homogeneity. Integral proteins such as the succinate dehydrogenase, for example, have been refractory to purification despite the development of an affinity chromatographic method utilizing the high-affinity inhibitor oxaloacetate linked to Sepharose 4B (Linder et al., 1975). Controlling the lability of membrane enzymes once dissociated from their lipid-rich domains (and possibly lipid cofactors) is a formidable task in membrane enzyme purification. At best, a number of the activities have been enriched in particulate fractions released selectively from the membranes, e.g., NADH dehydrogenase activity enriched in the supernatant fraction after EDT A treatment (Nachbar and Salton, 1970b) and cardiolipin synthetase in a lipid-rich particulate fraction released by low-ion ie-strength EDT A treatment (DeSiervo and Salton, 1971).


The only M. Iysodeikticus membrane enzyme to be purified to protein homogeneity is the F1-ATPase, the purification and properties of which will be discussed in detail in Section 3.6.1 of this contribution.

3.2. Enzymes Involved in Wall-Polymer Biosynthesis and Peptidoglycan Metabolism

Subcellular preparations of M. Iysodeikticus have been used in studies of the biosynthesis of the two cell-wall polymers of this organism-the peptidoglycan and the teichuronic acid polysaccharide containing mannosaminuronic acid. Mirelman and Sharon (1972) and Mirelman et al. (1974) were the first investigators to use a crude cell-wall preparation of M. luteus (Iysodeikticus) NCTC 2665 as a source of enzyme for studying peptidoglycan biosynthesis. This novel system consisting of wall and associated membrane had distinct advantages over the membrane preparations previously used by these investigators in that it not only catalyzed the polymerization of the sugar nucleotide precursors, but it also attached the newly synthesized peptidoglycan to the preexisting polymer in the cell wall (Mirelman et al., 1974). Transpeptidation reactions were observed in this type of preparation from both M. luteus and S. aureus. The crude cell-wall membrane fractions from these organisms incorporated the nucleotide precursors UDP-N-acetyl-D-glucosamine (UDPGlc- NAc) and UDP-N-acetylmuramyl-L-Ala-D-iso-Glu-L-Lys-D-Ala-D-Ala (UDP-MurNAc-pentapeptide)into preformed peptidoglycan accompanied by the release of C-terminal D-alanine from the Mur-NAc-pentapeptide. The release of the D-alanine by the transpeptidation reaction was completely inhibited by very low concentrations of penicillin, but surprisingly the incorporation of GlcNAc and MurNAc-pentapeptide into preexisting peptidoglycan was also markedly inhibited by penicillin. Mirelman et al. (1974) have therefore suggested that transpeptidation results not only in the interpeptide cross-links subsequent to the transfer by transglycosylation of linear glycan strands to the preexisting wall, but also in the covalent attachment of new strands to older ones. Thus it would appear from these studies that elongation and growth of the preexisting peptidoglycan is the result of the two reactions-the penicillininsensitive transglycosylation and the penicillin-sensitive transpeptidation. In contrast to the crude wall system, isolated membrane preparations resulted only in the formation of un-cross-linked glycopeptide formed both in the presence and absence of penicillin (Mirelman et ai., 1974). That the crude wall, and not the isolated membrane fractions, exhibited transpeptidase activity may be attributable to the presence of the enzyme at the wall-membrane interface or within the wall itself. This latter possibility is not entirely unlikely, because the D-alanine carboxypeptidase of this organism (Linder and Salton, 1975) was indeed found in the wall-lysozyme digest fraction on protoplasting. Moreover,


the existence of wall-associated autolytic enzymes has also been well documented (Higgins and Shockman, 1971). Another possibility that may account for the transpeptidase activity in the crude wall system, and not in the isolated membrane preparations, is that the reaction occurs only when the correct enzyme-substrate complex is formed with preexisting and newly synthesized peptidoglycan, thus presenting all the components in a sterically favorable alignment (Mirelman et al., 1974).

Although the crude wall-membrane fractions studied by Mirelman et al. (1974) had no oo-carboxypeptidase activity, Linder and Salton (1975) were able to detect o-alanine carboxypeptidase activity released into the protoplasting medium. With labeled UDP-muramylpentapeptide as substrate, o-alanine carboxypeptidase activity was detectable in washed plasma membranes of M. Iysodeikticus when the incubation mixture contained either Triton X-I00 or Brij-36T (0.2% w Iv). Purification of the enzyme from detergent solubilized plasma membranes proved unsuccessful, but the soluble form released into the protoplasting supernatant was amenable to partial purification by affinity chromatography on ampicillin-Sepharose (Linder and Salton, 1975). The enzyme was activated by Mg2+ ions, had an alkaline pH optimum, and the Km for substrate was 0.118 mM, thus exhibiting properties rather similar to those of other oo-carboxypeptidases (Ghuysen, 1977). Moreover, the enzyme activity was completely inhibited by low concentrations of ampicillin and penicillin G (50% inhibition levels were 50 nM and 7 nM, respectively). The function of this enzyme in peptidoglycan metabolism in M. Iysodeikticus is at present unknown.

The cell wall of M. Iysodeikticus possesses an associated polysaccharide containing glucose and mannosaminuronic acid, as first reported by Perkins (1963). This teichuronic acid constitutes less than 10% of the wall material. Chemical studies have established that the polysaccharide chains contain 10-40 residues of [6-0-fj-(o-glucopyranosyl)-a(1' -- 4)-N-acetyl-o-mannosaminuronic acid] (Hase and Matsushima, 1970, 1972; Nasir-ud-Din and Jeanloz, 1976). The biosynthesis of the teichuronic acid has been investigated with particulate fractions of M. Iysodeikticus by Anderson and his colleagues (Anderson et al., 1972; Page and Anderson, 1972; Rohr et al., 1977; Stark el ai., 1977). The preparations catalyze the incorporation of approximately equimolar quantities of glucose (Glc) from UDP-Glc and N-acetyl-o-mannosaminuronic acid (ManNAcUA) from UDP-ManNAcUA into polysaccharide provided that the reaction mixture also contains UDP-GlcNAc (Anderson et aI., 1972; Page and Anderson, 1972). A heat-stable factor from the soluble fraction of cell extracts increased the level of incorporation. Two stages in the synthesis of the polysaccharide have been recognized. The first stage involves the assembly of a linkage region on a lipid carrier by the transfer of GlcNAc from UDPGlcNAc to form GlcNAc-carrier lipid. The latter serves as the acceptor of two


residues of ManNAcUA from UDP-ManNAcUA to form (ManNAcUA)2-GlcNAc-carrier lipid for the completion of the first stage of teichuronic acid biosynthesis (Rohr et al., 1977). The (ManNAcUA)2-GlcNAc-carrier lipid acts as an acceptor of the glucosyl and N-acetyl-D-mannosaminuronosyl residues from their respective UDP-sugars in the second stage reaction to form (ManNAcUA-Glc)n-ManNAcUA-Glc-(Man-NAcUA)2-GlcNAc-carrier lipid. The N-acetyl-mannosaminuronosyl transferase and glucosyltransferase have been solubilized by Triton X-IOO. No evidence was obtained for a second carrier lipid in the transfer of the two sugar residues to the linkage-carrier lipid, and indeed the inhibition of synthesis of teichuronic acid intermediates by concomitant synthesis of peptidoglycan or membrane lipomannan suggests that a common lipid carrier (undecaprenyl mono phosphate) is involved (Rohr et al., 1977). On the basis of enzymatic and chemical degradation, Hase and Matsushima (1977) have suggested that the M. Iysodeikticus teichuronic acid is linked to peptidoglycan by the following structure:

Teichuronic acid (1 ..... 3) GlcNAc (1 ..... 6)-muramic acid (peptidoglycan)

The detection and structure of other sugar aminouronic acids in polysaccharides of other gram-positive and gram-negative organisms has recently been reviewed by Tonn and Gander (1979) together with a discussion of the biosynthesis of these and other wall and membrane polymers.

3.3. Lipomannan Biosynthesis

Gilby et al. (1958) were the first to report that membrane fractions isolated from M. Iysodeikticus were rich in carbohydrate (approximately 20%) and that the principal monosaccharide detectable was mannose. The nature of the membrane component remained uncertain until Macfarlane (1962) indicated that it was a membrane-associated polymer of high molecular weight and extractable with phenol from the cellular residues (i.e., membrane) sedimentable after lysozyme lysis of M. Iysodeikticus. Further studies of the mannan by Schmit et al. (1974) led to the conclusion that the "mannan contains neutral polymannose units of approximately 50,000 molecular weight covalently bonded through a base-labile linkage to some other moiety that is negatively charged at neutral pH." Subsequent investigations by three groups showed independently that the mannan was an acidic polymer with esterified succinyl residues, accounting for the negative charge (Owen and Salton, 1975b; Pless et aI., 1975; Powell et al., 1975). Moreover, all three groups showed the presence of glycerol and fatty acids. Structural studies of the mannan by Scher and Lennarz (1969) suggested, on the basis of exhaustive methylation, that mannosyl units were linked at the 3-, 2-, and 6-positions in the


approximate ratio of 2 : 2 : 1. Subsequent studies indicated a polymer of 50-60 mannosyl residues with two branch points and a (1 -->- 1) linkage to a diglyceride (Pless et aI., 1975; Powell et al .• 1975). Linkage of the mannan to diglyceride residues would thus ensure its anchoring in the membrane, and by analogy to the lipoteichoic acid polymers of other gram-positive membranes (Wicken and Knox, 1975) the term lipomannan has been applied to this component and has received wide acceptance.

Of particular interest is the observation by Powell et al. (1975) that none of the three Micrococcus species examined (M. Iysodeikticus, M. jiavus, and M. sodonensis) contained lipoteichoic acids, but lipomannan was found in all three organisms. It was suggested that this acidic lipomannan may perform a MgH ion-binding function similar to that of lipoteichoic acids (Powell et aI., 1975). Although the precise physiological functions of these membrane-bound acidic polymers have yet to be firmly established, their presence appears to be confined to the membranes of gram-positive bacteria, and so far only one particular kind of lipid-linked polymer has been found in the membranes of a given species (i.e., lipoteichoic acid and lipomannan do not appear to occur together). These findings may have some taxonomic and evolutionary significance. One could speculate that in the absence of the physiological advantages of an outer membrane barrier such as that of the gram-negative bacterial cell (Leive, 1974), the gram-positive organisms have evolved acidic, divalent cation-sequestering polymers in the vicinity of enzymes such as the carboxypeptidase-transpeptidase enzymes with rather high MgH ion requirements for their optimal activities. Other functions for the lipoteichoic acids as inhibitors of peripherally located autolysins have been suggested (Holtje and Tomasz, 1975; Cleveland et aI., 1975), and it is possible that these membrane-associated acidic polymers have multiple functions. The succinylated lipomannan is one of the major antigens exposed on the outer surface of the protoplast (Owen and Salton, 1977); it could also function as a determinant of the asymmetry of the plasma membrane and provide a charged hydrophilic interface between membrane and wall and binding sites for some of the more basic proteins, such as the DD

carboxypeptidases, transpeptidases, and autolytic enzymes, believed to be functioning in this region of the cell periphery.

The biosynthesis of the succinylated lipomannan of M. lysodeikticus has not been investigated as extensively as have the wall peptidoglycan and teichuronic acid. Early studies by Scher et al. (1968) and Scher and Lennarz (1969) provided evidence for mannosyl-l-phosphorylundecaprenol as the mannosyl donor for mannan synthesis. Plasma membranes of M. lysodeikticus catalyze the transfer of [14C]mannose residues from GDp-[14C]mannose to yield [14C] mannosyl-l-phosphorylundecaprenol carrier as the principal labeled mannolipid extractable from the membrane preparations (Owen and Salton, 1975a). The nature of the lipid carrier as [14C] mannosyl-l-phosphorylunde-


caprenol was confirmed by mass spectrometry (Owen and Salton, 1975a). Incubation of the purified labeled [14C]mannosyl lipid carrier in a plasma membrane reaction mixture containing Triton X-IOO resulted in the incorporation of mannose residues into the succinylated lipomannan. Although this transferase activity was present in both the isolated plasma membrane preparations and the purified mesosomal membrane fractions, the levels were higher in the plasma membranes. On the other hand, the mesosomes appeared to be unable to synthesize significant amounts of the mannosyl-lipid carrier as determined by incorporation of [14C]mannose from GDp-[14C]mannose into the undecaprenol intermediate. The ratio of labeled carrier lipid synthesized by the plasma and by the mesosomal membranes was about 34 : I. The deficiency of this enzyme activity in the mesosomal membranes clearly indicates that they are not the major sites of synthesis of the lipomannan despite the fact that this component is greatly enriched in these structures (Owen and Salton, 1975a).

The nature of the acceptor of mannosyl residues has yet to be established. Scher and Lennarz (1969) reported that in vitro transfer resulted in the formation primarily of nonreducing terminal mannosyl residues to some acceptor. In the studies of Owen and Salton (1975a) the lipomannan could be labeled with mannosyl residues from GDP-mannose and mannosyl-I-phosphorylundecaprenol carrier, but whether the [14C]mannosyl residues were terminal or within the polysaccharide chains was not established. Further investigations are needed to determine the nature of the acceptor and the possible existence of a linkage component in the biosynthesis of the lipomannan. Although [14C] succinate proved a convenient label for the succinylated lipomannan (Owen and Salton, 1975b), the sequence of its addition to acceptor, linkage unit, or lipomannan has yet to be defined.

As with the deacylated forms of lipoteichoic acids (Markham et al .. 1975; Joseph and Shockman, 1975), there is evidence that deacylated lipomannan is also released into the culture medium during the growth of M. Iysodeikticus (Owen and Salton, 197 5c). The significance of the release of these deacyla ted forms of membrane lipoteichoic acid and lipomannan has yet to be determined.

Membranes of M. Iysodeikticus contain mannobiosyldiglyceride, and the two enzymes responsible for its synthesis have been studied (Lennarz and Talamo, 1966), one of which is membrane bound and catalyzes the first step of the reaction transferring a mannosyl residue from GDP-mannose to 1,2-diglyceride. The second reaction is caused by the action of a soluble cytoplasmic enzyme catalyzing the addition of a second mannosyl residue from GDP-mannose to mannosyldiglyceride, with the formation of mannobiosyldiglyceride. Whether the latter acts as an intermediate in lipomannan synthesis has not yet been determined, but it was detected as a minor labeled product along with the labeled lipid carrier in the plasma membrane extracts (Owen and Salton, 1975a). There would seem to be a good probability that this glycolipid has


some relationship to lipomannan formation, especially as the lipoteichoic acids of a given species usually contain the glycosylglycerides characteristic of the membrane glycolipids of the same species (Baddiley, 1972; Brundish and Baddiley, 1968; Button and Hemmings, 1976).

3.4. Enzymes Involved in Lipid Biosynthesis

The membranes of M. lysodeikticus contain about 20-30% total lipid, with phospholipids accounting for the major portion of the lipid, amounting to approximately 75-80% (Salton and Freer, 1965; DeSiervo and Salton, 1971, 1973). The principal phospholipids are phosphatidylglycerol and diphosphatidyl glycerol (cardiolipin) and minor amounts of phosphatidylinositol and phosphatidic acid (Macfarlane, 1962; DeSiervo and Salton, 1971, 1973). Phosphatidylethanolamine, which is the principal phospholipid of the membranes of many gram-negative bacteria as well as a number of gram-positive organisms (O'Leary, 1967; Salton, 1971), is absent in M. lysodeikticus. The amounts of phosphatidylglycerol and cardiolipin undergo marked changes in the cells throughout the growth sequence, the fall in the relative percentage of phosphatidylglycerol during early to midexponential growth phase coinciding with an increase in the cardiolipin content. Abrupt changes were observed when growth ceases, with the phosphatidylglycerol rising to a maximum and cardiolipin falling to a minimum in the stationary phase (DeSiervo and Salton, 1973). Phosphatidylinositol and phosphatidic acid showed relatively small changes throughout the growth sequence.

Macfarlane (1962, 1964) reported the presence of glycosyldiglycerides and amino acyl phospholipids in M. lysodeikticus. In addition to the phospholipids and glycolipids, lipid-soluble carotenoids, menaquinone-9, and undecaprenol derivatives account for a minor portion (about 5-10%) of the total membrane lipid. Changes in the contents of these throughout the growth cycle have not been documented, nor has their biosynthesis been investigated in this organism apart from the reactions involved in polyisoprenoid synthesis in M. lysodeikticus extracts (Sagami et al., 1977, 1978). Although prenyltransferases have been found in extracts prepared from M. lysodeikticus, the distribution of these enzymes in cytoplasm and membranes has not yet been investigated. Such enzymes involved in carbon-chain elongation in isoprenoid biosynthesis are clearly relevant to the biosynthesis of the membrane menaquinones and undecaprenol components of the membrane.

The fatty acid constituents of the membranes of M. lysodeikticus have been identified by several investigators (Lennarz, 1961; Macfarlane, 1962; Cho and Salton, 1966; Whiteside et al., 1971), and the CI5 branched-chain fatty acid is by far the most abundant, accounting for as much as 80-90% of the total fatty acids. However, as with other bacterial species (Kates, 1964;


Table II Principal Fatty Acid Composition of Lipids Extracted from M. lysodeikticus Plasma

Membranes Isolated from Cells Grown on Complex and Defined Media

% Composition of fatty acids

Peptone water-yeast extract medium

M. R. J. Salton and Defined Cho and Whiteside et J. H. Freer medium of

Fatty acids Salton (1965) al. (1971) (unpublished data) Salton (1964)

Saturated straight-chain 14: 0 4.4 1.3 2.8 Trace 16: 0 0.2 0.5 2.2 1.9 18: 0 Trace Trace Trace

Branched-chain 14: 0 2.0 6.1 2.0 15: 0 85.4 91.3 84.6 45.1 16: 0 5.0 0.7 1.1 9.2 17: 0 2.6 1.7 1.3 10.1 18: 0 Trace Trace 2.0 30.5

O'Leary, 1967), the fatty acid composition is subject to marked changes, depending on the growth conditions. The influence of growth medium on the fatty acid composition of M. lysodeikticus membranes is illustrated in Table II (M. R. J. Salton and J. H. Freer, unpublished data). On the complex peptone-yeast extract medium, the C I5 branched-chain fatty acid accounted for 85-90% of the total fatty acids, whereas in cells grown in a defined medium (Salton, 1964), the C I5 branched-chain fatty acid content dropped to about 50%. Apart from showing that [14C] isoleucine was an excellent carbon source for the branched-chain fatty acids with 14C incorporated specifically into CI5

and C I7 acids (Lennarz, 1961), little work has been done on the enzymes involved in fatty acid biosynthesis and metabolism in this organism.

Phospholipid biosynthesis has been investigated in membrane preparations of M. lysodeikticus by DeSiervo and Salton (1971) and Rosenthal and Salton (1974). The study of cardiolipin biosynthesis (DeSiervo and Salton, 1971) provided the first evidence that the reaction leading to the synthesis of cardiolipin in bacteria differed from that found in other organisms (Cronan and Vagelos, 1972). Evidence obtained with particulate fractions of E. coli suggested that cardiolipin was synthesized by the following overall reaction as proposed by Chang and Kennedy (1967):

Phosphatidylglycerol + CDP-diglyceride -- diphosphatidylglycerol + CMP


However, the investigations by DeSiervo and Salton (1971) indicated that cardiolipin was formed by the following reaction:

2-Phosphatidylglycerol -- diphosphatidylglycerol + glycerol

and that the reaction had no specific requirement for added CDP-diglyceride. This reaction was catalyzed by a lipid-rich particulate fraction released

from the membrane by treatment with EDT A at low ionic strength. The activity in the isolated membranes and the particulate fraction was increased by the inclusion of Triton X-I 00 in the incubation mixture, a property characteristic of a number of membrane enzymes. Subsequent studies by Short and White (1972) with S. aureus membranes and by Hirschberg and Kennedy (1972) with E. coli particulate fractions confirmed the synthesis of cardiolipin by this reaction by establishing its stoichiometry as well as the origin of the glycerol moiety. This route of cardiolipin biosynthesis therefore appears to be unique to prokaryotic cells.

Attempts to purify the enzyme have so far been unsuccessful. Although Triton X-I00 (0.25% w jv) was used in the initial assays for the cardiolipin synthetase, longer periods of exposure of the particulate enzyme to this nonionic detergent resulted in substantial losses of activity (Rosenthal and Salton, 1974). Triton X-I00 has subsequently been replaced by Nonidet P40 (0.05% wjv) in the assay system (Rosenthal and Salton, 1974), but the enzyme has so far proved refractory to purification.

In the course of examining the subcellular distribution of the cardiolipin synthetase activity, it was noted that higher levels were found in the plasma membrane fraction than that detected in the isolated mesosomal preparations (L. Vitvitski-Trepo and M. R. J. Salton, unpublished data). The ratio of cardiolipin synthetase activity in the plasma membrane to that in the isolated mesosomes from cells in the late log phase was approximately 6 : 1. To test the possibility that the plasma membranes may contain some factors required for the reaction that were absent in the mesosomes, the two fractions were mixed and assayed. Much to our surprise the addition of mesosome fractions to the plasma membrane preparations caused a marked inhibition of cardiolipin synthesis, and the level of inhibition was dependent on the amount of mesosomal fraction up to a plateau level (L. Vitvitski-Trepo and M. R. J. Salton, unpublished data). Maximal inhibitory activity of 87% was observed by the addition of the mesosomal membrane preparation to plasma membranes at a protein ratio of 4 : 1. The inhibitory activity of the mesosomes could not be accounted for by its high lipomannan content, as inhibition was not observed with the purified lipomannan. The nature of this inhibitory activity has yet to be determined. It was destroyed by ashing but was only partly inactivated by proteolysis and was moderately resistant to heating at lOWe.


As far as we are aware, this appears to be the first instance of an activity, albeit an inhibitory one, that seems to be localized in the mesosome fraction. It will be recalled that most of the enzymes detectable in the plasma membranes are either greatly diminished or absent in the mesosome fractions (Salton and Owen, 1976; see also Section 4 for discussion of the antigenic architecture of membranes). Whether there is any interchange of material between the mesosomes and plasma membrane has yet to be examined. Perhaps relevant to this question is the intriguing observation by Bureau et al. (1976) that a "periplasmic" protein isolated from exponentially growing cells of Bacillus subtilis stimulates the exchange of fatty acids from the mesosomes to the plasma membrane. The protein does not, however, appear to be associated with the mesosomes, as it was isolated from the supernatant lysate of the wall following mesosome extrusion as carried out by Frehel et al. (1970). The relationships of all these interesting observations to membrane lipid biosynthesis and membrane assembly seem worthy of further pursuit. Furthermore, the recent demonstration that the phospholipids of the outer leaflet of the plasma membrane of protoplasts of M. Iysodeikticus can be manipulated by rat liver exchange protein (Barsukov et al., 1978) opens the way to a new approach that could be useful for studying membrane lipid function and biosynthesis in both mesosomes and plasma membranes. The recent detection of phospholipid transfer activity in cell-free extracts of Rhodopseudomonas sphaeroides (Cohen et al., 1979) is the first report to indicate the existence of prokaryotic lipid exchange proteins. These investigations should add further exciting details of the regulation and transfer of lipids in the surface structures of bacterial cells.

3.5. Electron-Transport Chain Components

Gel'man and her colleagues have made an extensive study of the characterization of the electron-transport chain and its organization in M. Iysodeikficus membranes; much of the individual investigations is summarized in the monograph by Gel'man et al. (1975). As in other bacteria, the cytochromes of M. Iysodeikticus are exclusively localized in the plasma membrane fractions. Earlier suggestions that the mesosomes were the prokaryotic equivalents of the mitochondrial structures, enriched in cytochromes and respiratory chain components (Ferrandes et al., 1966), were subsequently withdrawn, having been based on incompletely separated mesosomal and plasma membranes (Frehel et al., 1971). Cytochromes a2, b, b I , and c have been detected and quantitated; in membranes from stationary phase cells of M. Iysodeikticus the contents were a2, 0.18; b I , 0.48; and c, 0.47 nmolesjmg protein (Simakova et al., 1969; Tikhonova et aI., 1970). To the author's knowledge, none of the M. Iysodeik-


ticus cytochromes has been isolated as a homogeneous protein fraction, a state of affairs not unique to bacterial cytochrome components.

The dehydrogenases found in M. Iysodeikticus membranes include alcohol dehydrogenase, malate dehydrogenase, and two distinct NADH dehydrogenases, D-Iactate dehydrogenase and succinate dehydrogenase (Gel'man et al., 1960, 1975; Kulyash et al., 1978; Mitchell, 1963; Nachbar and Salton, 1970b; Owen and Freer, 1970; Pollock et aI., 1971; Collins and Salton, 1979). As pointed out by Gel'man et al. (1975) the dehydrogenases of bacterial membranes differ in their binding strength with the membrane, some being firmly bound (e.g., malate and succinate dehydrogenases) and others (e.g., NADH dehydrogenase II: Collins and Salton, 1979) may be released under relatively mild perturbation of the membrane and behave more like peripheral membrane proteins. Isocitrate dehydrogenase of M. Iysodeikticus cells was about equally partitioned between the cytoplasmic fraction and the membranes (Nachbar and Salton, 1970a).

Release and solubilization of bacterial membrane cytochromes and dehydrogenases is not always selective and quantitative. Ostrovskii et al. (1968) and Tsfasman et al. (1972) found that 30% of the malate and NADH dehydrogenase complex could be released by EDT A treatment, whereas the rest is extracted by detergents. In a recent study of the extraction of membranes with detergents and chaotropic salts, Collins and Salton (1979) found that the maximal extraction of various components from the membranes usually followed the bulk solubilization of protein. However, certain treatments giving preparations enriched in certain specific activities (e.g., NADH dehydrogenase II) may have advantages for the release of some membrane components, even though the overall yield may be low (Collins and Salton, 1979).

It is likely that many of the particulate fractions obtained from bacterial membranes are heterogeneous. Thus the EDT A extractable fraction of Ostrovskii et al. (1968) contained malate and NADH dehydrogenase activities, and the fraction released by Nachbar and Salton (1970b) was a lipid-rich particle fraction containing two bands in polyacrylamide gel staining for NADH dehydrogenase activity. The lipid-depleted deoxycholate insoluble residue of M. Iysodeikticus membranes isolated by Salton et al. (1968) contained succinate dehydrogenase and cytochromes a, b, and c, but appeared to be stripped of the lO-nm particles subsequently identified as the F1-ATPase (Oppenheim and Salton, 1973). Other types of complexes isolated from M. Iysodeikticus membranes include malate dehydrogenase-NADH dehydrogenase-cytochrome b556 and succinate dehydrogenase-cytochrome b (see Gel'man et al. 1975).

As pointed out by Gel'man et al. (1975) isolation and purification of firmly bound dehydrogenases can be difficult and the detergents needed for solubilization can frequently lead to inactivation of the enzymes. Removal of tightly bound lipid from mitochondrial succinate dehydrogenase and purification of this enzyme to near homogeneity has taken a long time and was even-


tually facilitated by the introduction of chaotropic agents (Hatefi and Hanstein, 1969; Davis and Hatefi, 1971). Although some of the procedures used in the purification of mitochondrial succinate dehydrogenase were applied to the M. Iysodeikticus enzyme, its lability only permitted partial purification even with the use of an affinity chromatography system (Pollock et al., 1971; Linder et al., 1975). The strongly hydrophobic character of such integral membrane enzymes combined with oxygen lability of the succinate dehydrogenase present formidable barriers to their purification.

Menaquinone-9 is the principal lipid-soluble menaquinone found in the membranes of M. Iysodeikticus (Salton and Schmitt, 1967), although several isoprenyl homologues may also be present, as reported by Jeffries et al. (1969). The molecular mechanisms through which they participate in the energy-transducing electron-transport system not only of bacteria, but of chloroplasts and mitochondria as well, has yet to be fully elucidated (Crane, 1977). Quinone is required in order for electron transport to occur, and the lipophilic quinone undergoes rapid oxidation-reduction changes. In the gram-positive organism Mycobacterium phlei the quinone (menaquinone) appears to function in NADH oxidation at a site before cytochrome b (Brodie and Gutuick, 1972). As pointed out by Crane (1977), "the view that lipophilic qui nones can be used to establish transmembrane proton gradient is supported by studies of nonenzymatic-ubiquinone oxidation and reduction in a model lipid-bilayer system, which showed [that] autoxidation of UQ6H2 formed by addition of NADH on one side of a lipid bilayer caused the formation of a proton gradient across the membrane. There is no doubt that the universal occurrence of lipophilic quinones in energy coupling membranes can logically be connected to proton gradient generation across the membrane."

The extensive investigations on the M. Iysodeikticus respiratory chain by Gel'man et al. (1972, 1975) and Fujita et al. (1966) suggest that menaquinone functions in the transfer of electrons from specific flavoproteins and further to oxygen via cytochrome types c and a. The respiratory chain components of M. Iysodeikticus membranes are thus believed to be organized according to the scheme presented below (Gel' man et al., 1975; Tikhonova, 1974).

Malate - FP ~ Lactate - FP ~ NADH - FP .A

Succinate - FP ?

Detergent, uv irradiation HQNO

i Ascorbate + TMPD

NaCN

FP, flavoprotein; TMPD, NNN' N'-tetramethyl-p-phenylenediamine; HQNO, hydroxyquinoline N-oxide; K2, menaquinone.


The respiratory chain of M. lysodeikticus could be inhibited by hydroxyquinoline N-oxide and cyanide, but was resistant to inhibitors of mitochondrial electron transfer, such as rotenone and antimycin. Gel'man et al. (1970) found that detergents were specifically inhibitory for the menaquinone region of the chain, and it was later shown that ultraviolet irradiation acted similarly (Tikhonova, 1974).

In their further important studies of the respiratory chain organization and topography, several types of preparations were used by Grinius et al. (1972) and Tikhonova et al. (1973). Sonication of protoplasts of M. lysodeikticus yielded inside-out vesicles, as does sonication of mitochondria, to give submitochondrial particles. Osmotic lysis of protoplasts gave mixtures of rightside-out and inside-out vesicles. Respiring sonic particles were shown to form a membrane potential (+ inside). Results of experiments designed to localize the membrane-potential-generating sites of the M. lysodeikticus respiratory chain suggested that ferricyanide accepts electrons before the LlJLHv generating point in the malate oxidase system, and after this point in the lactate oxidase component (Tikhonova, 1974).

The term LlJLH+, in accord with the chemiosmotic hypothesis of Mitchell (1961, 1968), refers to the transmembrane difference of the electrochemical potentials of hydrogen ions, and transformation of the LlJLH+ energy can be utilized for the catalysis of A TP synthesis and to drive a number of translocases (Harold, 1972).

Both Triton X-IOO and uv irradiation inhibition of electron transfer from lactate to ferricyanide could be reactivated by addition of menadione. It was concluded, therefore, that naphthoquinone (menaquinone-9) participates in electron transfer between lactate and ferricyanide and that it is not involved in electron transfer between malate and ferricyanide (Tikhonova, 1974). Further experiments with ferricyanide, which does not penetrate coupling membranes, indicated that components of the respiratory chain were located along the outer surface of the sonic membrane vesicles, a finding also in accord with the higher oxygen consumption and dehydrogenase activities observed with sonic membranes than that found for protoplasts (Tikhonova et aI., 1973; Tikhonova, 1974).

From their extensive data on anion transport and arrangement of some of the redox enzymes in M. lysodeikticus membranes, Tikhonova (1974) has been able to suggest the details of the arrangement of the respiratory chain and the topography of the components as illustrated in Figure 4.

More recent investigations of the respiratory chain components of M. lysodeikticus by Tikhonova et al. (1979) indicate the reduction of ferricyanide by endogenous substrates of intact protoplasts when the terminal portion of the chain is blocked. The evidence suggests that at least one component of the


In Membrane Out

Malate

NADH---J~--.!.

Succinate---~ ..

sonication

TMPD

" Ascorbate

FIGURE 4. Diagrammatic representation of the topographical arrangement of the respiratory chain in the membrane of M. Iysodeikticus as proposed by Tikhonova (1974) and adapted from Gel'man et al. (1975). Abbreviations: FP, flavoprotein; MaJ, malate; Lact, lactate; DPI, dichlorophenol-indophenol; HQNO, hydroxyquinoline N-oxide; TMPD, NNNN'-tetramethyl-pphenylenediamine; Succ, succinate; Fe"h, non haem-iron protein.

chain is located on the outer surface of the protoplast membrane, appearing to be functionally situated in the region of the cytochromes, thereby confirming the hypothesis of a transmembrane organization of the respiratory chain in this organism. Identification of the component will be eagerly awaited.


With the elegant detailed biochemical analysis of the respiratory chain of M. Iysodeikticus by the USSR research groups of N. S. Gel'man, D. N. Ostrovskii, and G. V. Tikhonova, the specific interactions between components of the electron transport chain should now be amenable to experimental verification by cross-linking studies to determine nearest neighbor contacts between the individual dehydrogenases and cytochromes. Although the purification of the integral proteins of the electron transport chain has presented difficulties, their recognition by specific antisera and as denatured subunits should greatly facilitate the analysis of their associations with one another, and some of the immunochemical approaches discussed in Section 4 could be utilized in further studying the molecular topography of the respiratory chain.

3.6. Membrane Adenosine Triphosphatase (FJ-ATPase)

Membrane A TPases have been recognized as important components of mitochondria and chloroplasts for some time; this enzyme activity was associated with a particle that could be detached from the membranes by mechanical or sonic disruption of the mitochondria (Racker, 1970). The knobbed, stalklike particles seen on the inner mitochondrial membrane were identified as the site of the mitochondrial ATPase (Racker et al., 1964). The released particulate fractions were homogeneous in the ultracentrifuge and were believed to be a single component. The term "coupling factor 1" (Fj) was given to this component, and these factors were thought to participate in a coupling device that transformed oxidative energy into ATP energy (Racker, 1970). The function of the Fj-ATPases as essential components of energy metabolism in all living cells became more apparent with the development of the chemiosmotic hypothesis by Mitchell (1961), although the precise mechanisms whereby A TP is formed from ADP has yet to be established (Downie et aI., 1979). All bacteria-both aerobic and anaerobic species-so far examined have been shown to possess Fj-ATPases; they have also been found recently in cyanobacteria (Binder and Bachofen, 1979; Owers-Narhi et al., 1979). Their universal role in energy transduction mechanisms in eukaryotic and prokaryotic cells alike is now firmly established.

Abrams et al. (1960) were the first to draw attention to the bacterial ATPases when they reported the association of the enzyme with washed membranes of Streptococcus facecalis. Subsequent studies showed that the ATPase could be released from bacterial membranes by subjecting them to low-ionicstrength "shock" washes (Ishikawa and Lehninger, 1962; Abrams, 1965), a procedure that has received wide application for the release and purification of the ATPases from most bacterial species (Downie et al., 1979).

The elucidation of the structure-function relationships in mitochondrial organelles combined biochemical approaches with electron-microscopic char-


acterization of the ultrastructure of the membranes and particulate fractions released or extracted from these organelles (Stoeckenius, 1970; Racker, 1970). Similar combined biochemical and ultrastructural approaches were also essential for probing and resolving the functions of the "simple" undifferentiated bacterial cytoplasmic membrane, which contained most of the components organized in the separate mitochondrial organelle of eukaryotic cells.

A remarkable similarity of the stalklike particles seen on membrane fragments of Bacillus stearothermophilus to those of inner mitochondrial membranes was observed in negatively stained preparations examined in the electron microscope by Abram (1965). As the bacterial membranes probably performed the functions of mitochondrial organelles, she suggested that these structural units were the prokaryote analogues. The biochemical and ultrastructural evidence that the 10-nm particles of the bacterial membranes were indeed the ATPase (F t ) components of bacterial membranes was provided by the studies of Oppenheim and Salton (1973), using specific antibodies to the purified ATPase of M. Iysodeikticus, which had been conjugated with ferritin for immunoelectron-microscopic identification (see Oppenheim and Nachbar, 1977). The fact that the ATPase could be released from the membranes of M. Iysodeikticus by a low-ionic-strength shock wash provided a unique opportunity for identifying a membrane enzyme as a ferritin-antibody-enzyme complex (Oppenheim and Salton, 1973). Membranes labeled with a ferritin conjugate of antibody to the purified ATPase yielded the ferritin-antibodyATPase complex on shock washing, and the nature of the new molecular species from the membrane was established by reaction with specific antisera to the ferritin and immunoglobulin components of the complex (Oppenheim and Salton, 1973). Moreover, earlier electron-microscopic studies of purified ATPase had established that the preparations contained a homogeneous population of uniform particles of about 10-nm diameter with a rosettelike appearance of six peripheral subunits and a central unit (Munoz et al., 1968a). The appearance of these particles in negatively stained preparations of the pruified ATPase of M. Iysodeikticus corresponded remarkably well to the subunit composition subsequently studied by SDS-polyacrylamide gel electrophoresis (Huberman and Salton, 1979).

In studying the structure-function relationships of the membranes of M. Iysodeikticus, it became apparent that electron microscopy of the membranes and fractions released from the membranes could provide valuable evidence to be correlated with biochemical investigations of membrane components. The appearance of membranes before and after sUbjecting them to the shock-wash treatment, which released the major portion of the ATPase activity from the membrane, could be correlated with the release of the 10-nm particles into the supernatant fraction after removal of the depleted membranes, as shown in Figure 5A-C. Negatively stained preparations show an abundance of the 10-


FIGURE 5. Electron micrographs of negatively stained fractions of M. lysodeikticus plasma membranes (A) before (see also Figure 1) and (8) after being subjected to the shock-wash step for release of F,-ATPase from the membranes. Note the stripped appearance and virtual absence

nm particles on the membrane; upon shock washing, the particles are released into a supernatant fraction that contains the ATPase activity and gives a band of ATPase activity on electrophoresis in polyacrylamide gels, and this major band constitutes the principal protein in this fraction (Munoz et al .. 1968b, 1969). The membrane residues shown in Figure 5B have been virtually


of lO-nm particles. (C) The crude shock wash supernatant illustrating the abundance of lO-nm particles together with some fragments of the membranes.

stripped of these particles. Purification of the F1-ATPase yielded a peak fraction of activity containing lO-nm particles of uniform appearance in the negatively stained preparations seen in the electron microscope (Figure 6). Improved methods of purification have shown the FI-A TPase particle fractions to be homogeneous by several criteria (Huberman and Salton, 1979). This



evidence together with the direct labeling of the ATPase on the membranes by use of monospecific antibody conjugated to ferritin conclusively established the site of the F1-ATPase on the bacterial membrane as the lO-nm particles displayed on the inner surface of the bacterial plasma membranes. Further verification of the site of the F1-ATPase on the cytoplasmic side of the plasma


FIGURE 6. Uniform particles of the F,-ATPase seen in the peak fraction of activity on purification on Sephadex 0-200. The diameter of the particles is about 10 nm. (X 156,000, reproduced at 75%)

membranes of M. lysodeikticus was also obtained by 125I-Iabeling of intact protoplasts and isolated membranes by the lactoperoxidase method (Salton et al., 1972) and by antibody absorption experiments (Owen and Salton, 1975d, 1977). The latter will be discussed in more detail in Section 4. Of significance was the failure to detect the F1-ATPase particles in the mesosome fractions, either by electron microscopy (Oppenheim and Salton, 1973), by antigenic analysis of the mesosomal membranes (Salton and Owen, 1976), or by direct ATPase assay in the presence or absence of trypsin (Oppenheim and Salton, 1973).

3.6.1. Purification and Properties of the M. lysodeikticus F1-ATPase

In its membrane-bound form the F1-ATPase of M. Iysodeikticus was shown by Munoz et al. (l968b) to be almost completely "latent," i.e., having extremely low levels of activity in the absence of trypsin stimulation. Thus the membrane F1-ATPase of this organism behaves similarly to those of other strictly aerobic organisms, such as Azotobacter vinelandii and Mycobacterium ph lei (see review by Haddock and Jones, 1977), exhibiting a high level of


latency, as does the chloroplast ATPase complex CF1. The latter was shown by Vambutas and Racker (1965) to respond to the stimulatory effects of trypsin treatment and subsequent experiments with membrane-bound forms of certain bacterial ATPases also demonstrated similar latency phenomena. In contrast, the membrane-bound ATPase of the facultative anaerobe Streptococcus faecalis appears to be nonlatent by virture of its unresponsivenes to trypsin stimulation (Abrams and Smith, 1974). In contrast, E. coli membrane ATPase activity is stimulated severalfold by trypsin (Carreira et aI., 1973). The molecular basis for the latency of certain membrane-bound ATPase has yet to be fully elucidated, although the € subunit of the F1-ATPases with its known inhibitory activity (Petersen, 1975; Nieuwenhuis and Bakkenist, 1977) would appear to be an excellent candidate for the regulation of the latency when the Fl is complexed with the Fa component in the membrane. Further investigations are clearly needed before the molecular mechanisms for achieving the latent state on the membrane can be understood. The functions of each of the individual five subunits of the FI-A TPases have yet to be fully established and reconstitution experiments performed before their roles in latency can be conclusively demonstrated.

In general, the F1-ATPases released from the bacterial membranes exhibited lower levels of latency than that found for the membrane-bound forms (Munoz et aI., 1969; Salton and Schor, 1974). Moreover, Salton and Schor (1974) found a progressive loss of trypsin-stimulated activity during purification of the shock-wash ATPase and the form released into the aqueous phase by n-butanol extraction of the membranes was completely nonlatent. The reasons for the loss of latency during purification of the soluble M. Iysodeikticus F1-ATPase became apparent in subsequent studies when it was found that the inclusion of the serine protease inhibitor phenyl methyl sulfonyl fluoride (PMSF) in the shock-wash buffer helped maintain a highly latent form of the Fl (Huberman and Salton, 1979). Thus it became possible to obtain homogeneous F1-ATPase preparations exhibiting a sevenfold trypsin stimulation by including 1 mM PMSF in the shock-release step and maintaining the stability of Fl with glycerol during the purification on DEAE-Sephadex (Huberman and Salton, 1979). It would therefore appear that the losses of trypsin stimulation that occurred in earlier studies were the result of endogenous protease activities, and indeed we have found in M. lysodeikticus membranes a protease with a substrate profile resembling that of elastase (M. Huberman and M. R. 1. Salton, unpublished results). Proteolytic degradation of the F1-ATPase of Micrococcus sp. ATCC 398E by autoproteolysis has been reported by Risi et al. (1977). Degradation of several of the various subunits of the Fl was observed, and it could be prevented with diisopropylfluorophosphate (DFP). Uncontrolled proteolysis has thus led to loss of latency in our studies and undoubtedly accounts for much confusion in subunit molecular weights and


stoichiometry manifest from an examination of the literature on bacterial F)ATPases from various organisms and by different investigations.

The principal steps involved in the purification of M. Iysodeikticus F)ATPase are similar to those now in general use for many bacterial F)-ATPases and are based essentially on the earlier observations of Abrams (1965) and Ishikawa and Lehninger (1962). By subjecting washed bacterial membranes to a low-ionic-strength wash (usually referred to as a shock wash) the F)ATPases are released from the membrane. As illustrated in Figure 5A-C, the procedure works particularly well with M. Iysodeikticus, and the great enrichment of the F) particles in the supernatant fraction is dramatically shown electron microscopically in Figure 5C. In earlier studies we had shown that NADH dehydrogenase (Nachbar and Salton, 1970b) was a major contaminant of the shock supernatant when this was performed on membranes washed by the standard procedure we had devised (Salton, 1967). Nachbar and Salton (1970a) found that NADH dehydrogenase activity could be released from M. Iysodeikticus membranes by holding them overnight at 4 0 C in 50 mM Tris-l mM EDT A without significant loss of the ATPase into the Tris-EDT A supernatant wash. Prior treatment in this way was incorporated into the scheme for purifying the ATPase by subjecting the residues after the Tris-EDT A wash to the low-ionic-strength shock-wash step in 3 mM Tris buffer, pH 7.5. As indicated above, this is now done in the presence of 1 mM PMSF to inhibit proteolytic activity that may be released from the membranes, and glycerol is added to maintain the stability of this multimeric enzyme during subsequent purification steps with gradient elution from DEAE-Sephadex columns (Huberman and Salton, 1979) as the final step. The F)-ATPase thus purified was judged homogeneous on the basis of its behavior in the ultracentrifuge, uniformity of particle size, and appearance in the electron microscope; also, it gave a single band in nondissociating polyacrylamide gel electrophoresis stainable either with Coomassie Brilliant Blue or for ATPase activity (Munoz et al., 1969), as well as a single peak in the two-dimensional (crossed) immunoelectrophoresis (Owen and Salton, 1975d, 1977), using antibodies to the isolated membranes.

The two-dimensional, crossed, and rocket immunoelectrophoresis techniques are high-resolution and sensitive immunochemical procedures that have proved extremely valuable in studying membrane enzyme purification. In contrast to immunodiffusion techniques for studying antigen-antibody reactions, the rocket and crossed immunoelectrophoresis methods involve the electrophoresis of antigens into antibodies in agarose gels buffered at pH 8.6, thus permitting anodal migration of negatively charged antigens into immunoglobulins immobile at this pH (Laurell, 1965). As the antigen-antibody complexes coalesce they become immobile in the agarose gel and form rocket-shaped immunoprecipitates. Electrophoresis of the antigens can be performed directly into agarose-containing antibody (rocket immunoelectrophoresis) or the anti-


A

B

c


gens can be separated by electrophoresis in one direction on the slides with agarose alone; the agarose strip containing the electrophoretically separated antigens retained on the slide and the rest of the gel replaced with agarosecontaining antibodies for the second-dimension electrophoresis into the antibody gel (Axelsen et al., 1973). The basic one-dimensional (rocket) and twodimensional immunoelectrophoresis techniques have been applicable to a variety of membrane problems. Variations of the basic methodology such as the fused-rocket technique have been very useful in examining a large number of fractions (e.g., from column separations of antigens) by simultaneous electrophoresis of antigens from adjacent wells into antibody gel. Thus M. Huberman, P. Owen, and M. R. J. Salton [unpublished data quoted by Owen and Smyth (1977)] were able to follow the purification of M. Iysodeikticus F]-ATPase by applying the fused-rocket procedure (Axelsen et al., 1973) to column fractions. In this way, the complete elution profile of the enzyme, detected as immunoprecipitated antigen, was obtained. Moreover, the elution of other membrane components and their separation from the ATPase could be readily seen on the immunoplate (Figure 6; Owen and Smyth, 1977). The availability of antisera to the isolated membranes and to the purified F]-ATPases provides a valuable adjunct to the purification of the F]-A TPase and emphasizes the general utility of these immunoelectrophoresis procedures in enzyme or antigen purification or both, by conventional biochemical methods. Figure 7 A shows the pattern of immunoprecipitates obtained with the crude shock-wash fraction from M. Iysodeikticus membranes for the two-dimensional, crossed immunoelectrophoresis of the antigens into purified immunoglobulins to the membrane antigens. As anticipated, the major immunoprecipitate peak is attributable to the presence of F]-ATPase, and the immunoplate shows the presence of a number of other membrane antigens released by the shock-wash procedure. Figure 7B illustrates the detection of immunoprecipitates of the pooled peak fractions following gradient elution from DEAE-Sephadex. It can be seen that by using antibodies to the membrane it is possible to detect minor contaminants in the peak fraction as immunoprecipitates in the crossed immunoelectrophoresis sys-

FIGURE 7. (A) Crossed immunoelectrophoresis of thc crude shock wash fraction from M. lysodeikticus membranes. (B) The peak fractions of activity following chromatography on DEAE-Sephadex as described by Huberman and Salton (1979). The immunoelectrophoresis and immunoplates were prepared and performed as described by Owen and Salton (1975d, 1977). Antigens in (A, B) were electrophoresed into antimembrane, and the plates illustrate that the ATPase is the major antigen present. Several minor peaks are seen after the purification step. (C) An immunoplate of the purified fraction with anti-ATPase in the reference gel. Trace contaminants are shown by this technique. (A-C) The protein loadings of antigens were 20 ltg, l2ltg, and 7 Mg, respectively. Electrophoresis was performed with anodal migration to the left in the first direction and to the top in the second direction. Plates were processed and stained with Coomassie Brilliant Blue as described previously (Owen and Salton, 1975d, 1977).


tem. Such contaminants probably accounting for less than 5% of the total protein of the fraction can be detected in this very sensitive system owing to amplification of their detection by the reaction of the antigens with immunoglobulins and staining with Coomassie Brilliant Blue. This amplification is especially true for detecting contaminating succinylated lipomannan, a glycolipid that does not stain with Coomassie Blue, but that is readily detectable even in nanogram amounts (Owen and Salton, 1976) as an immunoprecipitate. Thus its detection in the M. Iysodeikticus F) fractions can only be reliably established by these sensitive immunoelectrophoretic procedures, as staining for carbohydrate on SDS gels is relatively insensitive in comparison with the immunoglobulin or concanavalin A systems (Owen and Salton, 1976). The minor protein components are generally not detectable on SDS-polyacrylamide gel electrophoresis slabs, even when run with two to three times the amount of protein applied to the immunoplate illustrated in Figure 7B. When the same purified fraction of F)-ATPase was examined against antiserum to purified ATPase, as seen in Figure 7C, only traces of minor immunoprecipitates were detected. The detection of these minor components also indicates that they were present in the preparation of purified ATPase used as immunogen.

The use of these high-resolution sensitive analytical procedures for determining enzyme and antigen homogeneity is especially important in the light of the recent claims by Munoz and his colleagues (Andreu et aI., 1978) that bacterial and chloroplast F)'s are glycoproteins. The procedures we have used and illustrated above will provide the methods for a critical examination of this problem. A range of lectins in intermediate gels could be used to establish the sugar specificity of the carbohydrate portions and to conclusively demonstrate their glycoprotein nature and possible cross-reactivity. So far we have been unable to detect any reaction of the M. Iysodeikticus F)-ATPase either with concanavalin A or with wheat germ lectin (Owen and Salton, 1975d, 1977).

Earlier investigations in our laboratory (Munoz et al., 1969) and subsequent studies in Munoz laboratory have established that the soluble M. Iysodeikticus F1-ATPase is very similar in its general properties to other bacterial ATPases (Abrams and Smith, 1974; Salton, 1974; Haddock and Jones, 1977). Its activity is stimulated by CaH , and its substrate profile, pH optimum, and reaction to inhibitors are similar to those from other bacterial species (Munoz et aI., 1969; Salton, 1974). As is also the case for other bacterial ATPases, the M. Iysodeikticus FI is insensitive to inhibition with oligomycin, ouabain, and p-chloromercuribenzoate, and it is inhibited by azide and is sensitive to 1-anilinonaphthalene-8-sulfonate and phloretin (Munoz et al., 1969; Salton, 1974). I n contrast to some F 1-A TPases, the M. lysodeikticus F 1 is not particularly cold labile. However, as pointed out by Salton (1974) the sensitivity of the bacterial A TPases to cold has not been as carefully documented over a range of protein concentrations as has been done for the mitochondrial ATPase (Penefsky and Warner, 1965).


Much of the earlier work on the molecular weights of the purified ATPase complex and the subunit composition and their molecular weights must now be reevaluated because of the problems of proteolytic degradation. Although independent estimates of the molecular weight of the M. Iysodeikticus Fj-ATPase by Munoz and his colleagues (Andreu et al., 1973) and by Salton and his colleagues (Schor et al., 1974) yielded molecular mass values of approximately 350,000 daltons, it would now appear certain that these are underestimates of the native F j molecular mass because of possible proteolytic degradation and/or loss of subunits during purification. More recent studies by Huberman and Salton (1979) under conditions of purification controlling proteolysis gave estimates of the molecular mass of the highly latent Fj-ATPase of the order of 400,000 daltons. Moreover, this value is in closer accord with the estimate based on the tentative subunit composition and molecular masses of apparently undegraded subunits.

It is now generally recognized that Fj-ATPases from bacteria as well as mitochondria and chloroplasts possess five subunits-two major subunits a and fl, and three minor subunits /" 0, and ~ (Pedersen, 1975; Abrams and Smith, 1974; Haddock and Jones, 1977). The subunit composition of M. Iysodeikticus ATPases showed considerable variation from preparation to preparation and varied in number from two major ones (a and fl) in the n-butanol-released ATPase to fractions containing the two major subunits together with variable appearance of minor subunits as seen in SDS-polyacrylamide gels (Salton and Schor, 1972, 1974; Munoz et al., 1969; Carreira et al., 1976; Ayala et al., 1977). Indeed, a confusing array of different forms of M. Iysodeikticus ATPase varying from two major and two minor subunits to one form containing one major and two minor subunits has been reported by Munoz and his colleagues (Carreira et al., 1976). These investigators have reported an Fj-ATPase from M. Iysodeikticus with four subunits a, fl, /" and E in which the E subunit has a molecular mass of about 25,000 daltons. The latter is almost certainly a degraded from of the 0 subunit with the E undetected either because of the SDS-polyacrylamide gel system used or because of its possible loss following proteolytic cleavage of subunits.

Carefully controlling proteolytic activity with PMSF permitted isolation of a five-subunit Fj-ATPase from M. Iysodeikticus membranes, the subunit molecular weights of which are summarized in Table III together with the molecular weights of the subunits that have undergone endogeneous proteolytic degradation in the absence of PMSF or altered by direct modification on treatment with trypsin. Uncontrolled protease activity in the Fj fractions can ultimately lead to the complete loss of latency and conversion of a subunit to a' together with degradation of 0 to 0' or lower-molecular-weight components. So far as we have been able to tell from inspection of SDS-polyacrylamide gels, there appears to be little degradation of the a', fl, and/, subunits, although more precise quantitative data will be required to firmly establish this conclu-

348

Table III Molecular Weights of Subunits and

Degraded Subunits of M. lysodeikticus Latent Ft-ATPase"

Subunit Molecular weight

a 60,000 a' 58,000-59,000 (3 54,000

'Y 37,000 0 27,000 0' 22,000

9,000

"Data from Huberman and Salton (1979), and unpublished data on the degraded (proteolysis and in absence of phenyl-methyl sulfonyl fluoride) a' and 0'.

Milton R. J. Salton

sion. Because of the low staining intensity of the f subunit, it will have to be carefully quantitated by other techniques, such as use of radioactively labeled Fl' Our evidence therefore indicates that the M. Iysodeikticus FI-ATPase conforms to the five-subunit FI's of mitochondria, chloroplasts, and other bacterial species; also, the molecular weights of the individual subunits are very similar to those reported for the various bacterial FI-ATPases (Pedersen, 1975; Haddock and Jones, 1977).

The suggestion that the FI is a glycoprotein and that all its subunits are glycosylated (Andreu et al., 1978; Guerrero et al., 1978) has yet to be confirmed; at this stage we have no decisive evidence supporting this interesting claim. None of the lectins so far investigated appears to have an affinity for the ATPase, although several other membrane components apart from the succinylated lipomannan do seem to interact (Owen and Salton, 1977). The relatively large variety of sugars reported to be present in these FI-A TPases by Andreu et al. (1978) would make them quite unique as a class of glycoproteins; if their presence is confirmed, it would suggest that the carbohydrate structures are unusual for a glycoprotein, which might account for their lack of reactivity with lectins.

Apart from the problems of proteolysis confusing the variety and molecular weights of the FI-A TPase subunit polypeptides (Risi et al., 1977; Huberman and Salton, 1979) the stoichiometry of the subunits in all FI-ATPases has been a controversial matter for some time (Pedersen, 1975; Haddock and Jones, 1977). At one stage the evidence from mitochondrial and bacterial studies suggested the following subunit stoichiometries: lX, 3; (3, 3; 'Y, 1; 0, 1; and f,

1 (Senior, 1973; Haddock and Jones, 1977). More recent evidence has suggested lX2(32 together with 'Y2 and f2 subunits (there is uncertainty about the


molar ratio of 0), to yield a dimeric symmetrical molecule (Amzel and Pedersen, 1978; Senior, 1975; Verschoor et al., 1977; Binder et al., 1978). Such a structure is thus incompatible with the earlier suggestions of aJf3{yjoj~j. Reconstitution studies of the E. coli Fj-ATPase have also resulted in a suggestion of an azf32'Y20j-2~2 stoichiometry (Vogel and Steinhart, 1976). Our own investigations with Fj-ATPase released from cells labeled with [j4C]amino acids supplied in the form of algal hydrolystate suggest a stoichiometry of aJf33'YjOj~3 (Huberman and Salton, 1979). Until the amino acid composition of the individual subunits is established and the labeling under the conditions employed is shown to give a uniform labeling of all subunits insensitive to individual cellular pools of amino acids as well as large differences in specific actitivies of amino acids in individual subunits, the stoichiometry determined by this method remains tentative. It should be noted, however, that similar labeling techniques have been used by Bragg and Hou (1975) for determining subunit composition of E. coli and Salmonella typhimurium Fj-ATPases and by Yoshida et al. (1975) for the Fj of the thermophilic bacterium. In all instances similar stoichiometries of a3.83'YjOj~j were obtained. The higher proportion of ~ subunits in the Fj-ATPase of M. lysodeikticus (a3.83'YjOj~3) would, in the view of our laboratory, be compatible with a highly latent ATPase, assuming that the ~ subunit [an inhibitor of hydrolytic activity in other Fj-ATPases, e.g., E. coli (Nieuwenhuis and Bakkenist, 1977; Laget and Smith, 1979)] is the regulatory polypeptide. Further investigations on the precise quantitation of the subunit composition are needed before the stoichiometry can be unequivocally established. At least at the present time these stoichiometries fit the higher molecular weights observed for the bacterial ATPase than would the dimeric structure a2.82'Y202~2.

In summary, then, the purified M. Iysodeikticus Fj-ATPase conforms to the five-subunit (a, .8, "I, 0, and ~) ATPases found in other bacteria, and the stoichiometries tentatively suggested are generally similar to those of other bacterial Fj's, although in M. lysodeikticus the ~ subunit appears to be in a 1 : 1 ratio with the a (or .B) subunit. The principal feature that distinguishes it from most of the bacterial Fj-ATPases is its high level of latency, a property it shares with other obligate aerobes and with chloroplast Fj-ATPase. The only other property reported for M. Iysodeikticus Fj-ATPase, and so far for none of the other bacterial Fj's, is the glycoprotein nature of the molecule and its individual subunits (Andreu et al., 1978; Guerrero et al., 1978). Independent confirmation that this and other Fj's are indeed glycoproteins with covalently linked carbohydrate residues is lacking at the present time.

3.6.2. Attachment and Functions of Coupling Factor (F)) ATPase

With the development of suitable procedures for the purification to protein homogeneity of the coupling factor ATPases, efforts then turned to the reconstitution of the coupled functions (Racker, 1970). It appeared highly probable


that in a multimeric structure such as the Fj-ATPase there would be specific subunits involved in its attachment to the Fo component in the membrane. As in mitochondrial and chloroplast studies, attempts to define the subunit requirements for association of bacterial F j with Fo have involved reconstitution of dissociated subunits or restoration of mutational defects, or both, by addition of wild-type subunits (for fuller discussion see Downie et al., 1979). Early investigations with Streptococcus faecalis ATPase by Baron and Abrams (1971) indicated the role of the polypeptide nectin, which was needed to attach the ATPase complex to depleted membranes. This polypeptide was believed to be one of the minor subunits-the b subunit of the ATPase. Involvement of minor subunits in reattaching the M. Iysodeikticus ATPase was also suggested from the work of Salton and Schor (1972), who found that the n-butanolreleased ATPase (composed of lX and (3 subunits) and ATPase, which had lost trypsin stimulation, were unable to bind to the depleted membranes. As the earlier purification and rebinding studies were not performed in the presence of protease inhibitors, it is possible that the interpretation of the results could have been complicated by endogenous protease activities.

Subunit requirements for reattachment of bacterial F j components have been studied in E. coli and the thermophilic bacterium by purification of individual subunits, as well as reconstitution. Sternweis (1978) prepared b,E-deficient E. coli ATPase and found that although it had normal ATPase activity, it was unable to r~bind to depleted membrane vesicles. Both the band E subunits were required for the binding of b,E,-deficient E. coli Fj to the membranes and the restoration of oxidative phosphorylation. Earlier studies by Bragg et al. (1973) had also indicated the requirement for b in binding F j to Fo by isolating an E. coli F j that lacked the b subunit. Neither b nor E subunits were found to bind directly in significant amounts to the depleted E. coli membranes (Sternweis and Smith, 1977; Sternweis, 1978). Other important reconstitution experiments with E. coli F j subunits by Futai (1977) showed that lX, (3, and 'Y subunits were required for reconstitution of ATPase activity. This finding contrasts with the thermophilic bacterium Fj, for which only (3 and 'Y subunits, or lX, (3, and b, were required for reconstitution of ATPase activity (Yoshida et al., 1977). However, as with the E. coli F j system and the reconstitution of energy-transforming activity (Vogel and Steinhart, 1976; Sternweis, 1978), all subunits of the thermophilic bacterium F j were also required for the reconstitution of activity with the Fo proton channel contained in liposomes (Yoshida et al., 1977). Moreover, the requirement of all five F j subunits was observed for the reconstitution of the 32Pj_A TP exchange reaction (Yoshida et aI., 1977). Subunit combinations such as lX, (3, 'Y, E and lX, (3, 'Y, which possess ATPase activities similar to those of the native F j were unable to give reconstituted H + - translocating vesicles. Thus, for the expression of energy-transforming activities, the complete Fj-Fo complex has to be reconstituted (Kagawa, 1978).

The functions of individual components in the F J and Fo complexes in var-


ious energy-requmng processes including oxidative phosphorylation has become amenable to biochemical genetics through the pioneering work initiated with the isolation of uncoupled (une) mutants of E. coli in Gibson's laboratory (Downie et at., 1979). Defects in both the F,-ATPase and the Fa component have been found, and reconstitution experiments have greatly facilitated the identification of the defective components. Thus the inactive F, from a strain carrying the unc A 401 allele could be reconstituted with an excess of a subunit from a normal F, to give ATPase activity (Dunn, 1978). Other mutants devoid of ATPase activity have been shown to possess abnormal {3 subunits in the F, (Downie et at., 1979). Although mutational defects in the Fa component have been genetically characterized (Downie et at., 1979), the defects in the polypeptide components involved have yet to be elucidated. This will await the further characterization of the Fa components of the bacterial membranes.

The Fa complexes of the energy-transducing systems have been studied primarily by isolating the complete Fo-F,-ATPase complex and identifying the subunits seen in addition to those of the well-characterized F,-A TPase sector. These complexes have been purified from beef heart mitochondria (Serrano et at., 1976) and from several microorganisms, including the thermophilic bacterium (Sone et at., 1975), Mycobacterium phtei (Lee et at., 1976), and yeast and Neurospora mitochondria (Sebald, 1977; Tzagoloff and Meagher, 1971; Ryrie, 1977). Much attention has been paid to the dicyclohexylcarbodiimide (DCCD)-reactive protein, which Altendorf and Zitzmann (1975) isolated from E. coli, suggesting that this hydrophobic protein is part of the Fa sector. The proteolipid nature of the DCCD-reactive protein in E. coli has been reported (Fillingame, 1975, 1976; Altendorf, 1977). In addition to inhibiting the ATPase activity of the membrane-bound F, through reaction of DCCD with the Fa component, it has recently been shown that mitochondrial F,-ATPase has binding sites and can also be inactivated by DCCD (Pougeois et at., 1979).

The Fa of the thermophilic bacterium consists of some eight to nine copies of polypeptides per Fa-F" with three different subunits of molecular masses ranging from 19,000 to 5,400 (Kagawa et at., 1976), the latter being identified as the DCCD-binding protein. It would seem a bit premature to make any firm conclusions about the stoichiometry of the Fa subunits. Kagawa (1978) concluded that available evidence shows Fa to be the H+ channel of the H+ATPase. The recent isolation of the Fa-F, complex of E. coli by Foster and Fillingame (1979) and the characterization of the variety of subunits will open the way to the identification of the functions of the individual components of Fa as well as facilitate the biochemical characterization of mutants with Fa defects. In addition to the F,-ATPase subunits, the E. coli Fa-F, complex showed the presence of three subunits with apparent molecular masses of 24,-000, 19,000, and 8,400, believed to be subunits of the Fa sector of the complex (Foster and Fillingame, 1979). The 8400-dalton subunit was identified as the DCCD-reactive proteolipid.


Although the M. Iysodeikticus F]-ATPase has been purified to protein homogeneity and has been well characterized, reconstitution experiments comparable to those performed with E. coli and the thermophilic bacterium F]A TPases have yet to be done. Such reconstitution experiments will, of course, be important in establishing the basis of latency in this F] and should provide a model for other obligate aerobes with highly latent F]-ATPases. The Fo component of the M. Iysodeikticus membrane has not been isolated, although evidence for a Fo-F] complex from detergent-solubilized membranes from Micrococcus lute us (ATCC4698) has been presented (Schmitt et al., 1978). Indeed, the behavior of a component that stains specifically for ATPase activity in Triton X-I00 extracts of M. Iysodeikticus membranes examined by crossed immunoelectrophoresis (Owen and Salton, 1975d) differed from that of purified soluble F]-ATPase released by the shock-wash procedure and led us to explore the possibility that it may be the Fo-F] complex. However, attempts to purify such a component from the Triton X-I00-solubilized membranes of M. Iysodeikticus did not succeed (M. L. Perille Collins and M. R. J. Salton unpublished results); further studies are needed to test the feasibility of applying the procedures used for the isolation of the complexes from Micrococcus luteus and E. coli.

The M. Iysodeikticus F]-ATPase is an H+ -ATPase of the proton-translocating A TPases involved in energy transduction processes, as proposed by Mitchell (1961,1968). Direct evidence for the function of this H+-ATPase in proton translocation was provided by the use of M. Iysodeikticus membrane vesicle preparations in the studies of Mileykovskaya et al. (1976). Sonicated membrane fragments of M. Iysodeikticus possessing latent ATPase activity were unable to generate a membrane potential difference by hydrolysis of ATP, but could do so by oxidation of suitable substrates (Mileykovskaya et al., 1976). It will be recalled that such vesicle preparations have the inside-out orientation with respect to the protoplast membrane. Trypsin treatment of the membrane vesicles did generate a membrane potential, as measured by the energy-dependent transport of the lipid anion phenyldicarbaundecaborane, and the membrane potential could be inhibited by dicyclohexylcarbodiimide. It was concluded that the membrane ATPase from M. Iysodeikticus exhibits its ability to generate ATP-hydrolysis-coupled membrane potential under conditions of inactivation of the ATPase complex protein inhibitor and can function, at least in vitro, as a reversible H+ -ATPase. The tight binding of the inhibitor in the M. Iysodeikticus F]-ATPase would thus serve as a device for maintaining the latent state and preventing the hydrolysis of A TP produced by respiration.

4. ANTIGENIC ARCHITECTURE OF THE MEMBRANE OF M. LYSODEIKTICUS

The specificity of antibody molecules for their determinant groups on antigens has long provided one of the most valuable means of detecting specific


cell-surface components. The chemical basis of the immunological reactions of capsular polysaccharides, lipopolysaccharides, and protein antigens of cell surfaces has given us much knowledge of the antigenic structure of many types of cells. The extension of immunochemical methodology into the field of cell structure and function has provided new insights into the antigenic architecture and molecular organization of cellular membranes and walls, especially in efforts to explore the nature of the surface structures of the bacterial cell.

With macromolecular structures such as membranes, one of the major problems has been to determine the variety of antigens in these insoluble organelles without having to resort to tedious absorption experiments or solubilization techniques (e.g., SDS dissociation), which destroy biological properties.

In order to determine the antigenic structure and asymmetry of membranes it was clearly necessary to be able to resolve the great variety of antigens expected to exist in a multifunctional structure. This has eventually been achieved both by solubilizing membranes in nonionic detergents, which are less likely to denature the membrane proteins, and by applying high-resolution immunochemical techniques such as two-dimensional crossed immunoelectrophoresis.

Early attempts to establish the antigenic complexity of the plasma membranes from M. Iysodeikticus by conventional agar gel-diffusion and immunoelectrophoresis techniques indicated a surprisingly small number of antigens (Fukui et al., 1971). Only two or three immunoprecipitates were seen in preparations dissociated by sonication at pH 9.0, treatment with 0.3% sodium dodecyl sulfate or Triton X-lOO, or by digestion of the membranes with trypsin, phospholipase A, or phospholipase C (Fukui et al., 1971). Similar results had also been observed with other bacterial membranes, including group A streptococcal membranes (Freimer, 1963) and mycoplasma membranes (Kahane and Razin, 1969). This finding was particularly surprising considering the great variety of enzyme functions performed by bacterial membranes and the number of polypeptides (about 40) seen upon SDS-polyacrylamide gel electrophoresis of dissociated membranes. Absorption studies with intact protoplasts indicated removal of antibodies to one of the membrane antigens; however, antibodies to the membrane ATPase were not absorbed (Fukui et al., 1971). This provided the first suggestion that the plasma membrane antigens were asymmetrically located, a finding that was extended to a much larger variety of membrane antigens when high-resolution procedures became available.

Following the development of the rocket immunoelectrophoresis technique by Laurell (1965),which demonstrated the formation of rockets of immunoprecipitates when antigens were electrophoresed into immunoglobulins, twodimensional immunoelectrophoresis emerged as a high-resolution procedure for analyzing complex mixtures of antigens and antibodies (Axelsen et al., 1973). With the expansion of the variety of techniques that could be used with two-dimensional crossed immunoelectrophoresis (e.g., intermediate gels with lectins), it became evident that this procedure was eminently suitable for the


resolution of the complex mixture of antigens present in cell-membrane structures. Its suitability for studying membrane problems became apparent from the studies of Johansson and Hjerten (1974) with the Tween 20 soluble membrane proteins of Acholeplasma laidlawii and Bjerrum's (1975) investigations of mammalian membrane proteins. The availability of a high-resolution sensitive procedure of analyzing membrane antigens was thus an important prerequisite for investigating the antigenic architecture of cell membranes-our ultimate goal in establishing the functional asymmetry of a prokaryotic membrane system. Furthermore, an essential feature of the crossed immunoelectrophoresis methodology was the retention of biological properties, as manifested by the ability of solubilized antigens to interact with antibodies to undenatured membrane proteins and the retention, in some instances, of enzymatic activities permitting identification of immunoprecipitates as specific enzymes by zymogram staining techniques (Axelsen et al., 1973; Owen and Smyth, 1977). The latter adds a further dimension to the utility of these methods in studying membrane proteins. Hence the crossed immunoelectrophoresis methodology is in marked contrast to the analysis of membrane proteins by SDS-polyacrylamide gel electrophoresis (SDS-PAGE), where most enzymes are completely inactivated and other biological activities are either reduced or lost. This is not to imply that SDS-PAGE is not useful, but it provides a different type of information about membrane proteins. One further advantage of the two-dimensional immunoelectrophoretic analysis is that for a given system of antigens and antibodies it provides quantitative information, the peak areas of individual immunoprecipitates being proportional to the antigen/antibody ratios (Axelsen et aI., 1973).

Other variations of the basic crossed immunoelectrophoresis method, such as the intermediate gel procedure, in which, for example, a lectin or a different antiserum is interposed between the electorphoresed antigens and the reference antibody system, permit the identification of glycoproteins or polysaccharide antigens reacting with lectins or the detection of specific antibodies in the intermediate gel serum. Thus a monospecific antibody incorporated into the intermediate gel can be used to identify a specific component in a complex mixture, because it will react only with its homologous antigen and form a precipitate in the intermediate gel. The rest of the antigens will be electrophoresed through the intermediate gel to form the remaining pattern of immunoprecipitates in the reference antibody gel. In this way ATPase was identified in a complex mixture of membrane antigens by use of anti-ATPase in the intermediate gel (Owen and Salton, 1975d).

The principal limitation of the technique as applied to membrane studies relates to the need to solubilize the membranes for electrophoresis in the agarose gels. Therefore only solubilized antigens will be detected in the immunoprecipitate patterns, although it should be remembered that the agarose gel has an exclusion limit of about 108 daltons and will permit the electrophoresis


of high-molecular-mass antigens through the gel. Any insoluble residues will, however, go undetected in this system. Fortunately M. Iysodekticus membranes are effectively solubilized by 1-4% Triton X-100, the non ionic detergent of choice for most membrane studies (Owen and Smyth, 1977). More than 75% of the M. Iysodeikticus membrane protein is solubilized under these conditions. It is possible that highly hydrophobic integral membrane proteins such as the cytochromes may not be seen under the conditions used for these procedures, and further efforts will be needed to specifically identify them in this type of antigenic analysis. Judicious use of lower ratios of SDS to protein than those used for SDS-PAGE may facilitate the examination of such refractory membrane proteins.

Seventeen distinct antigens were detected in the initial crossed immunoelectrophoresis analysis of the Triton X-100-soluble antigens of M. Iysodeikticus plasma membranes (Owen and Salton, 1975d). Five of the antigens were identified as enzymes by specific zymogram staining procedures. The membranes contained two distinct antigens-the immunoprecipitates of which stained for NADH dehydrogenase together with succinate and malate dehydrogenases and ATPase activities in individual precipitates (Owen and Salton, 1975d). None of the immunoprecipitates stained for more than one enzyme activity. The fact that multiple enzyme activities were not detected in any of the immunoprecipitates argues strongly in favor of the detection of individual enzyme species and the absence of any random membrane enzyme aggregates in the solubilized membrane preparations. In addition to the five enzymatically active immunoprecipitates, the identity of a major membrane antigen-the succinylated lipomannan-was also established by both affinoimmunoelectrophoresis with concanavalin A in the intermediate gel and by coelectrophoresis with the purified lipomannan (Owen and Salton, 1975d).

To maximize the detection of membrane antigens it is necessary to establish the immunoprecipitate patterns at high and low antigen loadings and for different periods of electrophoretic separation of the antigens in the first dimension (Axelsen et al., 1973). Under these conditions an additional 10-11 immuno precipitates were detectable (Owen and Salton, 1977), although they all appeared to be minor antigens. The crossed immunoelectrophoretic analysis of these membrane antigens with lectins as affinity absorbents of antigens containing carbohydrate residues has shown that at least five antigens react with concanavalin A, and all except one are exposed on the cell surface and outer surface of the protoplast membrane (Owen and Salton, 1977). In the expanded analysis of the membrane antigens by crossed immunoelectrophoresis no additional enzymes were detected despite the use of many additional enzyme-staining procedures for various phosphatases, esterases, dehydrogenases, and proteases. The failure to identify additional membrane enzymes in the immunoprecipitate patterns could be attributable to a number of factors including inactivation under the conditions of detergent solubilization and


analysis by crossed immunoelectrophoresis and total inhibition by reaction with specific antibodies. In considering the latter possibility, it is of interest to note that although the specific antibodies to purified M. lysodeikticus ATPase inhibited the enzyme by at least 95% (Whiteside and Salton, 1970), the residual activity was still readily detectable in immunoplates of free ATPase and Triton X-100-soluble membrane forms by zymogram staining (Owen and Salton, 1977).

In addition to the analysis of the plasma membranes, crossed immunoelectrophoresis of purified preparations of M. lysodeikticus mesosome membrane vesicles has been performed and has demonstrated the presence of major amounts of the succinylated lipomannan in the Triton X-lOO-solubilized mesosomes, but only trace amounts or undetectable quantities of the major five identifiable plasma membrane antigens, the two NADH dehydrogenases, succinate and malate dehydrogenases, and ATPase (Salton and Owen, 1976). It was suggested that because of the apparent enrichment of the outer membrane surface components in the mesosomal vesicles and the relative absence of the inner face plasma membrane enzymes the mesosomes represented structures formed from the vesicularization of the outer leaflet of the plasma membrane (Salton and Owen, 1976). Our studies of the antigenic analysis of the mesosomes by the crossed immunoelectrophoresis technique have confirmed and extended the paucity or lack of the important functional components found in the plasma membranes. The immunoplates presented in Figure 8 contrast the patterns obtained for the Triton X-lOO soluble fractions from the plasma membranes (Figure 8A) and mesosomes (Figure 8B).

Asymmetry or sided ness of membrane structures has been recognized from the early proposals of the bilayer model of Danielli and Davson (1935). Glycoproteins and other carbohydrate-rich components (e.g., macroglycolipids) have been identified as typical components of the outer surfaces of eukaryotic cell membranes (Brady and Fishman, 1975; Marchesi, 1975; Gardas, 1976; Dejter-Juszynski et al., 1978). Localization of components rich in carbohydrate residues on the outer face of surface membranes could serve multiple functions of providing the cell surface with hydrophilic components as well as a variety of sugar-mediated receptor or recognition sites, or both. Moreover, such components ensure the asymmetry of the surface membranes, and they may indeed be the major determinants of asymmetry during membrane assembly. In general, the prokaryotic cell has appeared to be relatively deficient in cell-surface glycoproteins, and there is little definitive chemical work establishing their occurrence in bacterial membranes. Perhaps the presence of the cell-wall peptidoglycan-polysaccharide structures in gram-positive organisms and the outer membranes of gram-negative bacteria rich in lipopolysaccharides have substituted for the carbohydrate-rich surface membranes of eukaryotic cells. Although cell-wall antigens of gram-positive bacteria and


A

B

FIGURE 8. The differences in the immunoprecipitate patterns for the Triton X-IOO solubilized fractions from (A) M. Iysodeikticus plasma membranes and (B) purified mesosomal vesicle preparations reacted against antimembrane antibodies, as reported by Salton and Owen (1976). Several of the major immunoprecipitates identified as specific membrane enzymes by zymogram staining (Owen and Salton, 1975d, 1977) are either absent or greatly diminished in the mesosomal pattern of immunoprecipitates.


lipopolysaccharides of the outer membrane of gram-negative organisms have been studied extensively, there has been little information on the asymmetry of the bacterial plasma membrane and few indications as to the nature of the components found on the outer surface of the membrane. The location of lipopolysaccharides on the outer cell surface of Salmonella typhimurium was demonstrated by Shands (1965) using ferritin-labeled antibodies to the lipopolysaccharide. The antigenic analysis of the plasma membranes of gram-negative bacteria has presented problems, for the presence of the outer membrane structure acts as a permeability barrier (Leive, 1974) and has generally prevented large molecules (e.g., immunoglobulins) from gaining direct access to the outer face of the plasma membrane. However, gram-positive bacteria with lysozymesensitive cell walls can be transformed into protoplasts with the outer surface of the plasma or protoplast membrane directly accessible for reaction with antibodies. As indicated earlier (Section 2), this was one of the advantages in selecting M. lysodeikticus as a model system in which to investigate the surface architecture and localization of membrane components. The introduction by Singer and Schick (1961) of ferritin-labeled antibodies for the detection of specific membrane components, has provided an elegant means for determining the localization of cell-surface and cell-membrane antigens by electron microscopy. Oppenheim and Nachbar (1977) have discussed various aspects of immunoelectron microscopy and its decisive use in the identification of the M. lysodeikticus F]-ATPase. One of the unique features of this procedure led to the identification of a membrane enzyme as a ferritin-antibody- F]-ATPase complex that could be released by the usual shock-wash procedure, but after labeling with the ferritin-anti-ATPase conjugate (Oppenheim and Salton, 1973). Localization of wall and membrane lipoteichoic acids of lactobacilli has also been achieved with the ferritin-antibody labeling procedure (Wicken and Knox, 1975).

Despite the appealing features of this method for localizing specific components with electron-dense, ferritin-antibody conjugates, the procedure involves extensive purification of the ferritin prior to conjugation and the retention of sufficient reactivity of the conjugated antibody toward its antigen to give adequate labeling (Oppenheim and Nachbar, 1977). However, the resolution of specific sites with ferritin-labeled antibody is considerably greater than that achieved with fluorescent-labeled antibody, peroxidase-labeled antibody, or cytochemical staining for enzymes resulting in heavy metal deposits to be visualized by electron microscopy. An alternate and highly sensitive procedure for determining the location of cell-surface antigens, albeit not directly visualized by electron microscopy, became possible through the combined use of conventional antibody absorption and crossed immunoelectrophoresis. Antibody absorption studies were, of course, the backbone of serology in the past, but the identity of the variety of antigens reacting in such absorption studies


was not resolvable in a simple fashion. By examining the remaining antibodies to a complex structure such as a membrane surface after sequential absorptions, it is possible in the high-resolution crossed immunoelectrophoresis system to determine which specific antigens are exposed and accessible for reaction with antibodies. Thus Owen and Salton (1975d, 1977) were able to determine which antigens of the M. lysodeikticus protoplast membrane were expressed on the outer surface of its intact plasma membrane. One of the major antigens of the outer face of the protoplast membrane was the succinylated lipomannan. Absorption of antimembrane immunoglobulins with increasing amounts of intact protoplasts resulted in the progressive removal of antibodies to the lipomannan. The eventual depletion of antibodies to this component resulted in the complete disappearance of its immunoprecipitate in the pattern of membrane antigens reacted against absorbed serum, as determined by crossed immunoelectrophoresis. In a typical absorption series in which increasing amounts of protoplasts or intact cells were absorbed against immunoglobulins to isolated plasma membrane preparations of M. lysodeikticus, antibodies to the succinylated lipomannan were readily absorbed, and the disappearance of other immunoprecipitates from the complex pattern for Triton X-lOa soluble membrane antigens also indicated that these antigens were exposed on the outer surface of the cells and protoplast membrane. Of the 27 discrete antigens detected in the membranes, absorption studies established that 12 were located on the protoplast surface, and of these at least five reacted with concanavalin A as did the major surface antigen, the succinylated lipomannan (Owen and Salton, 1977). Five of the major antigens identified by zymogram staining as two antigenically distinct NADH dehydrogenases-succinate and malate dehydrogenases and ATPase-were not detectable on the outer surface, as the peaks of these immunoprecipitates in the crossed immunoelectrophoretic analysis were unaffected even after extensive absorption with protoplasts. However, when isolated plasma membrane preparations with both faces of the membrane exposed were absorbed against antimembrane immunoglobulins, antibodies to these five enzymes, and indeed all other antigens, were completely absorbed as illustrated in Figure 9. The absorption series also serves to illustrate the marked differences in the rate of removal of antibodies to individual antigens, a feature that suggests differences in accessibility of the antigens on the membrane structures (e.g., compare the slower absorption of antibodies to antigens 10 and 20 in Figure 9 with fast removal for antigens 18 and 11).

These investigations have shown that the membrane of M. lysodeikticus is an asymmetrical structure with a number of antigens, apparently with carbohydrate residues, exposed on the outer surface. The other major antigens of the membrane identifiable as enzymes were accessible to reaction with antibodies only when absorption was carried out with isolated membrane preparations exposing both sides of the membrane; they must therefore be expressed


., A B

'. c o

20

FIGURE 9. (A-E) Absorption of antibodies to M. lysodeikticus plasma membrane antigens prepared with antimembrane antisera following absorption with increasing amounts of washed plasma membranes as described by Owen and Salton (1977). Note the rapid disappearance of immunoprecipitate 18 (identified as the succinylated lipomannan) from the pattern and the much slower disappearance of immunoprecipitates 10 and 20. Such differences in absorption could be caused by accessibility of antigens to antibodies. (E) All detectable antigens are expressed on the plasma membranes, as indicated by absence of immunoprecipitates. Immunoelectrophoresis conditions as previously described and immunoplates are from Owen and Salton (1977).


only on the cytoplasmic face (Owen and Salton, 1977). It is perhaps not surprising that important biochemical functions carried out by the membranes would occur on the cytoplasmic face and thus protect the loss of substrates and products to the external medium. However, not all membrane enzymes can be expected to share this asymmetry. Tikhonova et al. (1978) have presented evidence for a component of the respiratory chain of M. Iysodeikticus located on the outside of the membrane, thus supporting a transmembrane organization of the chain.

The crossed immunoelectrophoresis-absorption method as applied to protoplasts and isolated plasma membranes in our studies could not be used for the identification of transmembrane components. This could only be done by absorption studies with homogeneous, sealed inside-out vesicles from this organism. Unfortunately, we have not succeeded in achieving this goal at the present time. However, some of the conventional techniques for chemical labeling or enzymatic modification of such transmembrane components could be used for their specific identification (Carraway, 1975). The chromatophore membrane of photosynthetic bacteria is one of the few naturally occurring membrane systems possessing a normal inside-out orientation with respect to the plasma membrane. By carrying out absorption studies with spheroplasts presenting the right-side-out orientation of the plasma membrane accessible to antibodies and with isolated chromatophores of Rhodopseudomonas sphaeroides with their inside-out orientations, M. L. Perille Collins, D. E. Mallon, and R. A. Niederman (unpublished results) have been able to detect transmembrane components by the crossed immunoelectrophoresis method. The major barrier to doing this with the membranes of M. Iysodeikticus has been the difficulty in obtaining uniform populations of sealed membrane vesicles with the unnatural, inside-out orientation in high yields and on a more predictable experimental basis. As pointed out by Salton and Owen (1976) this has also been a general problem with other membrane systems, and it is not at all clear as to what factors govern the consistent production of uniformly sealed inside-out vesicles in the laboratory. Inside-out vesicles of M. Iysodeikticus have been prepared by sonicating protoplasts (Tikhonova, 1974), but the uniformity of the population was not indicated. Wientjes et al. (1979) have succeeded in producing populations of Bacillus licheniformis membrane vesicles with as high as 80% possessing the inside-out orientation. The vesicles were prepared by slow osmotic lysis of protoplasts of B. licheniformis and the insideout orientations determined by examining the fracture faces of the freeze-fractured preparations. The osmotic lysis by dialysis and use of Tris buffer both favored the inside-out formation. Wientjes et al. (1979) were unable to make definite conclusions as to the ways in which buffer composition and lysis procedure exert their effects. The E. coli membrane vesicles so extensively used in transport studies (Kaback, 1972) since their introduction by Kaback have been


shown to possess the right-side-out orientation for at least 95% of the vesicle population by carrying out the crossed immunoelectrophoresis analysis of absorbed membrane antisera (Owen and Kaback, 1978). Entrapment of many cytoplasmic antigens within these vesicles was also evident. Further studies are clearly needed to establish the parameters involved in the inside-out conversion of the membrane to yield, in an experimentally reliable fashion, the desired homogeneous populations of vesicles. This knowledge would greatly facilitate the studies of membrane asymmetry and function-structure relationships and would also permit a more critical evaluation of the significance of membrane enzyme translocation (or dislocation) reported for some vesicles.

Although the bacterial membrane in gram-positive species is enclosed in a continuous cell-wall structure, the detection of membrane antigens on the external surface of the intact cell was first documented for the lipoteichoic acids of the lactobacilli (Wicken and Knox, 1975). Such a surface expression does not appear to be explained simply on the basis of partial lysis of the wall and exposure of the underlying plasma membrane. Similar results were obtained when intact cells of M. lysodeikticus were absorbed with antimembrane immunoglobulins. Antibodies to all 12 antigens exposed on the protoplast surface were also absorbed by intact washed cells of M. lysodeikticus, although they were absorbed at greatly differing rates (Owen and Salton, 1977). For example, antibodies to the succinylated lipomannan and one other antigen were absorbed some six times more readily than were antibodies to two other surface antigens. Such differences in absorbability of antibodies to antigens on both protoplast and intact cell surfaces suggest marked differences in the accessibility of exposed antigens. Antibodies to the ATPase were rapidly removed on absorption with membranes, again suggesting accessibility, for it is also known to be a peripheral protein readily released from the cytoplasmic face of the membrane by the low-ionic-strength shock wash. An alternative explanation of the marked differences in apparent accessibility of the antigens is that the differences are attributable to antibody avidities. At present we favor the former explanation, as our crossed immunoelectrophoresis studies of the M. lysodeikticus membrane antigens have shown little evidence of "flying rockets" (i.e., immunoprecipitate peaks devoid of feet extending to the base of the antibody reference gels; see Axelsen et aI., 1973), an indicator of low-affinity antibodies. Further investigations will be needed, however, to determine whether accessibility or avidity is the major contributing factor to the substantial differences in rates of absorption to whole cells, intact protoplasts, and isolated membranes with exposed outer and cytoplasmic faces of the membranes. Another possibility is that absorption of antibody molecules to one antigen sterically hinders the binding of immunoglobulins to other neighboring antigens. This possibility would be amenable to testing by examining, for example, the rates of absorption before and after the shock-wash release of the ATPase.


One membrane antigen that was not detectable on the surface in the absorption studies with intact cells and protoplasts was absorbed when ricin and soybean agglutinins were used in intermediate gels with plasma membrane antibodies in the reference gels (Owen and Salton, 1977). These results suggest the possibility that this membrane antigen occurs on the cytoplasmic face of the plasma membrane and that it possesses o-galactosyl and N-acetyl-o-galactosaminyl residues reactive with ricin and soybean lectins, respectively. The existence of such an unusual component on the inner face of the membrane further emphasizes the asymmetry of this bacterial membrane structure.

In addition to the powerful immunochemical techniques available for establishing membrane asymmetry, advances have been made in establishing the asymmetric distribution of lipids in membrane bilayers (Bretscher, 1972; Op den Kamp, 1979). There seems little doubt that phospholipid asymmetry is firmly established in erythrocytes and other mammalian membranes, but in bacteria Op den Kamp (1979) concluded that the data "at best indicate that lipid asymmetry may exist." As Op den Kamp (1979) has emphasized, new methods and improved understanding of the naturally occurring flip-flop movement and transbilayer movements induced by lipid localization procedures will be essential for future conclusions regarding lipid asymmetry. Within the limitations of the existing procedures, Barsukov et al. (1976) suggested that in M. Iysodeikticus membranes there was a preferential localization of phosphatidylglycerol in the outer monolayer and of phosphatidylinositol in the inner layer, with cardiolipin distributed between both leaflets of the bilayer. The evidence available thus indicates that both phospholipids and proteins exhibit an asymmetric distribution in the M. Iysodeikticus membrane. Although the succinylated lipomannan is a major component of the outer face of the plasma membrane, we do not know if its distribution is strictly asymmetrical. The latter would have to await studies with inside-out vesicles, for example.

The remaining aspect of the investigations on the plasma membrane antigens analyzed by crossed immunoelectrophoresis worthy of mention relates to the use of crossed immunoelectrophoresis (CIE) as a powerful tool in determining the compartmentalization of cellular components. CIE with an intermediate gel (Axelsen et aI., 1973) provides a simple, sensitive technique for comparing complex antibody populations in two different antisera reacting against antigens in a single immunoplate. The cellular origins of immunogens (e.g., cytoplasmic versus membrane) in heterogeneous antigen fractions can therefore be readily determined by the intermediate-gel variation of CIE. Thus it was possible to determine the distribution of cellular antigens in the cytoplasmic and membrane fractions of M. Iysodeikticus. When antimembrane serum was placed in the intermediate gel below the cytoplasmic antiserum in the analysis of cytoplasmic antigens, several immunoprecipitates were formed in the intermediate gel, as illustrated in Figure 10. One of these stained for

A ·

B

a

c

a

t b

FIGURE 10. (A) The use of crossed immunoelectrophoresis with an intermediate gel is illustrated for the complex pattern of the antigens of the cytoplasmic fraction from M. lysodeikticus reacted against its homologous antiserum in gel region (a). No antiserum was added to the intermediate gel region (c) in immunoplate (A), but anti membrane serum was added to gel region (b) in immunoplate (B) . (A) NADH dehydrogenase (i,ii), isocitrate dehydrogenase (iii), poly-


NADH dehydrogenase and the other major immunoprecipitate formed with antimembrane antibodies in the intermediate gel had no counterpart in the reference pattern of immunoprecipitates for the cytoplasmic fraction (Owen and Salton, 1977). This component was identifiable as the succinylated lipomannan (by coelectrophoresis with purified lipomannan), which is immunogenic in the membrane-associated form, but was not immunogenic in the purified soluble form. Thus the intermediate gel with antibodies to membranebound antigens successfully detected the soluble non immunogenic form of the lipomannan in the cytoplasmic fraction (Figure 10). A portion of the membrane-associated lipomannan can be readily removed from the membrane, some of which may be solubilized during protoplast rupture in dilute buffer to facilitate the separation of membrane and cytoplasmic fractions.

By crossed immunoelectrophoresis experiments with intermediate gels, three NADH dehydrogenases have been detected in M. lysodeikticus. Of these, one NADH dehydrogenase appears to be largely membrane associated, one appears to be essentially cytoplasmic, and one distributes between both fractions. It will be recalled that one of the NADH dehydrogenases behaves as a typical peripheral protein and is more readily released from the membrane than is the other more tightly bound membrane NADH dehydrogenase antigen (Collins and Salton, 1979).

The cytoplasmic compartment of M. lysodeikticus is a very complex mixture of antigens, as revealed by reaction with anticytoplasmic antibodies in the two-dimensional immunoelectrophoresis reference system (Owen and Salton, 1977). At least 97 distinct immunoprecipitates could be resolved in the precipitin pattern, which markedly differed from that observed for the plasma membranes. The fact that the enzymes catalase, polynucleotide phosphorylase, and isocitrate dehydrogenase can be shown by zymogram staining to be localized in the cytoplasmic fraction attests both to the specificity of zymogram reactions and to the cell fractionation procedures. Moreover, when anticytoplasmic antibodies were examined in the intermediate gel with antimembrane immunoglobulins in the reference gel, the membrane antigen fraction gave only three immunoprecipitates in the intermediate gel, one of which was the NADH dehydrogenase distributed in both compartments. This highly sensitive procedure therefore indicates minimal contamination of the membrane fractions prepared by our standard wash procedure with antigens of the cytoplasmic compartment of M. lysodeikticus. These high-resolution methods thus provide powerful techniques for critical evaluation of cell fractionation and membrane

nucleotide phosphorylase (iv), and catalase (v). Two identifiable antigens (18, lipomannan; 10, NADH dehydrogenase) were fully included as immunoprecipitates in the antimembrane intermediate gel region (b). As in (A), the (a) gel region in (B) contained anticytoplasmic antibodies. From Owen and Salton (1977).


isolation procedures, and they have also been successfully applied to the resolution of the complex envelopes (inner and outer membranes) of the gram-negative organisms E. coli (Smyth et al., 1978) and Neisseria gonorrhoeae (Smyth et al., 1976).

5. SUMMARY AND CONCLUSIONS

The plasma membrane constitutes the surface membrane of the protoplast of the gram-positive organism Micrococcus Iysodeikticus after selective removal of the cell wall with lysozyme; this membrane can be recovered quantitatively on lysis of protoplasts or lysis of whole cells. In addition to the plasma membrane, mesosomal membrane vesicles are the only other membranous structures found in this organism. They are released upon protoplasting and can be isolated as homogeneous vesicle preparations. The plasma membrane has all the ultrastructural features common to cell membranes of both eukaryotic and prokaryotic origins, including the electron-microscopic appearance of membrane profiles in thin sections, the characteristic features of convex and concave freeze-fracture faces of membranes, and their general apperance upon negative staining. In common with all prokaryotic plasma membranes, negatively stained preparations show an abundance of uniform lO-nm particles, specifically identifiable as the F1-ATPase in M. Iysodeikticus. The isolated mesosomal membrane vesicles are readily distinguishable from the plasma membrane structures in negatively stained preparations, having a typical spherical or tubulovesicular appearance and apparent absence of the IO-nm ATPase particles.

The principal functions performed by this prokaryotic plasma membrane include those of active transport of metabolites, membrane energizing, and respiratory functions through its components of the electron-transport chain and energy-transducing ATPase, together with the membrane-stage enzymes involved in peptidoglycan, teichuronic acid, succinylated lipomannan, phospholipid, fatty acid, carotenoid, and menaquinone biosynthesis. The plasma membrane is thus characteristic of a multifunctional, prokaryotic membrane structure. Isolated mesosomal vesicles are either devoid or depleted of virtually all the biochemical activities found in the plasma membranes. The origins and functions of the mesosomes still remain a biochemical enigma, and the only unique property found for this membrane fraction has been an inhibitory action on the plasma membrane-cardiolipin synthetase activity.

The asymmetric nature of the plasma membrane of M. Iysodeikticus has been established by demonstrating the localization of the F1-ATPase on the cytoplasmic face of the membrane with ferritin-labeled anti-ATPase and by [125I]peroxidase labeling of protoplasts and membranes, as well as by two-


dimensional crossed immunoelectrophoretic analysis of absorbed antisera to membranes. The latter technique has involved resolution of Triton X-100-solubilized plasma membrane antigens and specific identification of two membrane NADH dehydrogenases-succinate and malate dehydrogenases-as well as FI-ATPase by zymogram staining of immunoprecipitates and identification of the succinylated lipomannan by coelectrophoresis with the purified lipomannan. All five identified enzymes were exposed as major antigens on the cytoplasmic face of the plasma membrane, being undetectable on the surface of the plasma membrane of intact protoplasts. Of the 27 distinct membrane antigens detectable by the crossed immunoelectrophoresis technique, the lipomannan was a major antigen of the protoplast surface and 11 other antigens, five of which reacted with concanavalin A, were also surface exposed. The isolated mesosomes were enriched in the succinylated lipomannan, and the major five enzyme antigens were undetectable or present only in very small or trace amounts. The asymmetry of these vesicles has not been determined.

Absorption studies suggest that some of the plasma membrane antigens, including the lipomannan and those classifiable as peripheral proteins (e.g., FI-ATPase), are more readily accessible to antibody molecules. No evidence has been obtained supporting the existence of antigens, which may be deeply embedded in the lipid bilayers, remaining unexpressed during immunization.

The plasma membrane of this prokaryotic cell appears to have its counterparts to the eukaryotic cell-surface glycoproteins in that the succinylated lipomannan and five other antigens possessing concanavalin-A-reactive components provide a carbohydrate-rich protoplast membrane surface. There is evidence that a component of the respiratory chain of M. Iysodeikticus also exists on the outer surface of the protoplast membrane, thereby suggesting a transmembrane organization of the chain (Tikhonova et aI., 1978). Exposure of components of the proton channel of the Fo-FI complex on the outer surface has yet to be explored. Evidence for the asymmetrical distribution of lipids in inner and outer monolayers of the bilayer (Barsukov et al., 1978) and the ability to modify the lipid composition with exchange proteins (Barsukov, 1978) will contribute further knowledge regarding the structure-function relationships in this membrane system.

ACKNOWLEDGMENTS

I wish to thank the National Science Foundation for grant support (PCM 78-24385) for the investigations carried out in his laboratory. I also wish to thank Kwang S. Kim for generously providing electron micrographs, Ludmilla Trepo-Vitvitski for the mesosome preparation, Carl Urban for the immunoplates of ATPase, and Peter Owen for the immunoplates in Figure 8. Special


thanks go to Josephine Markiewicz for her expert typing and help throughout the preparation of the manuscript.

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175-190, Plenum, New York. Nachbar, M. S., and Salton, M. R. J., 1970b, Biochim. Biophys. Acta 223:309-320. Nasir-ud-Din, and Jeanloz, R. W., 1976, Carbohydr. Res. 47:245-260. Nieuwenhuis, F. J. R. M., and Bakkenist, A. R. J., 1977, Biochim. Biophys. Acta 459:596-604. O'Leary, W. M., 1967, The Chemistry and Metabolism of Microbial Lipids, p. 93, World,

Cleveland, OH. Op den Kamp, J. A. F., 1979, Annu. Rev. Biochem. 48:47-71. Oppenheim, J. D., and Nachbar, M. S., 1977, in: Immunochemistry of Enzymes and Their

Antibodies (M. R. J. Salton, ed.), pp. 89-124, Wiley, New York. . Oppenheim, J. D., and Salton, M. R. J., 1973, Biochim. Biophys. Acta 298:297-322. Osborn, M. J., Gander, J. E., Parisi, E., and Carson, J., 1972a, J. BioI. Chern. 247:3962-3972. Osborn, M. J., Gander, J. E., and Parisi, E., 1972b, J. BioI. Chern. 247:3973-3986. Ostrovskii, D. N., Zhukova, I. G., and Gel'man, N. S., 1968, Biokhimiya 33:612-617. Owen, P., and Freer, J. H., 1970, Biochem. J. 120:237-243. Owen, P., and Freer, J. H., 1972, Biochem. J. 129:907-917. Owen, P., and Kaback, H. R., 1978, Proc. Natl. Acad. Sci. U.S.A. 75:3148-3152. Owen, P., and Salton, M. R. J., 1975a, Biochim. Biophys. Acta 406:235-247. Owen, P., and Salton, M. R. J., 1975b, Biochem. Biophys. Res. Commun. 63:875-880. Owen, P., and Salton, M. R. J., 1975c, Biochim. Biophys. Acta 406:214-234. Owen, P., and Salton, M. R. J., 1975d, Proc. Natl. Acad. Sci. U.s.A. 72:3711-3715. Owen, P., and Salton, M. R. J., 1976, Anal. Biochem. 73:20-26. Owen, P., and Salton, M. R. J., 1977, J. Bacteriol. 132:974-985. Owers-Narhi, L., Robinson, S. J., De Roo, C. S., and Yocum, C. F., 1979, Biochem. Biophys.

Res. Commun. 90:1025-1031. Page, R. L., and Anderson, J. S., 1972, J. BioI. Chern. 247:2471-2479. Pedersen, P. L., 1975, Bioenergetics 6:243-275. Penefsky, H. J., and Warner, R. C., 1965, J. Bioi. Chern. 240:4694-4702. Perkins, H. R., 1963, Biochem. J. 86:475-483. Pless, D. D., Schmit, A. S., and Lennarz, W. J., 1975, J. Bioi. Chern. 250:1319-1327. Pollock, J. J., Linder, R., and Salton, M. R. J., 1971, J. Bacteriol. 107:230-238. Pougeois, R., Satre, M., and Vignais, P. V., 1979, Biochemistry 18:1408-/413. Powell, D. A., Duckworth, M., and Baddiley, J., 1975, Biochem. J. 151:387-397. Racker, E., 1970, in: Membranes of Mitochondria and Chloroplasts (E. Racker, ed.), pp. 127-

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U.s.A. 74:936-940.

Some Recent Books in Cell Biochemistry and Biology

As in previous volumes of SUBCELLULAR BIOCHEMISTRY, we are including a review section on various texts that may be of interest to our readers. The aim of these book notices is to be as informative as possible and to give the reader a full idea of the range and scope of the publication being reviewed. The books listed below will be discussed in this article.

1. Cell and Membrane Biology

Structure of Biological Membranes edited by Sixten Abrahamsson and Irmin Pascher, Plenum Press: New York and London, 1976,580 pp.

Cell Motility by Howard Stebbings and Jeremy S. Hyams, Longman: London and New York, 1979, 192 pp.

Methods in Membrane Biology, Vol. 8, edited by Edward D. Korn, Plenum Press: New York and London, 1977,368 pp.

Essentials of Cell Biology (2nd ed.) by Robert D. Dyson, Allyn & Bacon: Boston, Mass., 1978, 433 pp.

Cell Biology: A Molecular Approach (2nd ed.) by Robert D. Dyson, Allyn & Bacon: Boston, Mass., 1978, 616 pp.

2. Genetics and Viruses

The Biochemistry and Viruses by S. J. Martin, Cambridge University Press: Cambridge, 1978, 145 pp.

Extranuclear Genetics by Geoffrey Beale and Jonathan Knowles, Edward Arnold: London, 1978, 142 pp.

The Phylogeny of Human Chromosomes by Hector N. SeUllnez, Springer-Verlag: Berlin, 1979, 189 pp.

3. Muscle and CaH Transport

Smooth Muscle (British Medical Bulletin, Vol. 35, No.3) edited by Edith Btilbring and T. B. Bolton, The British Council: London, 1979, pp. 209-316.

375

376 Some Recent Books in CeU Biochemistry and Biology

Calcium Transport and Cell Function (Annals of the New York Academy of Sciences, Vol. 307), edited by Antonio Scarpa and Ernesto Carafoli, New York Academy of Sciences: New York, 1978,655 pp.

4. General Biochemistry

Introduction to Biochemistry (2nd. ed.) by John W. Suttie, Holt, Rinehart and Winston: New York, 1977,434 pp.

Modern Concepts in Biochemistry (3rd ed.) by Robert C. Bohinski, Allyn & Bacon: Boston, Mass., 1979,600 pp.

Biochemistry: The Chemical Reactions of Living Cells by David E. Metzler, Academic Press: N ew York, 1977, 1129 pp.

1. CELL AND MEMBRANE BIOLOGY

The first book discussed in this section is Structure of Biological Membranes by S. Abrahamsson and I. Pascher. It is the proceedings of the 34th Nobel Symposium held in Sweden in 1976. Thirty contributions are included, ranging over a wide number of topics in membrane biology, biochemistry, and physical chemistry. Although it is now more than 4 years since the symposium was held, so high is the standard of the articles that the book's contents are still very relevant to contemporary work on biomembranes.

The book begins with an introductory chapter by S. Abrahamssoll, B. Dahlen, H. Lofgren, I. Pascher, and S. Sundell that shows how the various permitted packing arrangements of membrane lipids can give us an indication of the probable structure of biological membranes. The lipid layer of a surface membrane is visualized as consisting of the liquid region of the hydrocarbon matrix, the structural region of that matrix, the structural region of the polar part of the membrane and, finally, the surface functional region of the polar part. The behavior of the various regions is then related to the physicochemical properties of the lipids which make them up. Next, M. Anon, U. Pick, Y. Shahak, and Y. Siderer discuss proton transport through chloroplast membranes and its relation to energy conservation. The article strikingly demonstrates the close interrelationship between proton transport, light-induced electron transport, and A TP synthesis in chloroplasts, as well as reverse electronflow luminescence. Similar questions are discussed by M. Baltscheffsky in the next chapter on energy transduction in the chromatophore membrane. The organism Rhodospirillum rubrum was used as a model photosynthetic system; the absorbance of membrane-bound carotenoids changes during light-induced energy conversion, and the author discusses the significance of these alterations.

Some Recent Books in CeU Biochemistry and Biology 377

The next article is a treatment of "Monomolecular Films and Membrane Structure," by D. A. Cadenhead, in which the author compares two contrasting physical states of monomolecular films (the "liquid condensed" and the "liquid crystalline" states) with the "gel" and "liquid crystalline" states of hydrated lipid bilayers. D. Chapman and B. A. Cornell then give a brief account of phase transitions, protein aggregation, and membrane fluidity. They discuss the various ways in which lipids can pack and give an interesting model showing naturally induced packing faults in a ball-bearing raft.

G. Dallner then discusses the biosynthesis and transport of microsomal membrane proteins. As the proteins move via the smooth endoplasmic reticulum to the Golgi system, the oligosaccharide element is continually being synthesized; final release into the cytoplasm is via a lipoprotein complex. There follows a short paper by L. L. M. van Deenen, J. de Gier, L. M. G. van Golde, I. L. D. Nauta, W. Renooy, A. J. Verkleij, and R. F. A. Zwaal on the asymmetry of lipids in the erythrocyte membrane and the fusion of plasma lipoproteins with the membrane. The relationships between phospholipid metabolism in plasma and erythrocyte are also discussed. A. Ehrenberg, Y. Shimoyama, and L. E. G. Eriksson then describe methods for evaluating EPR spectra of spin-labeled amphilic molecules in lipid bilayers. The method was applied to orientated and unorientated samples and used to study the influence of cholesterol on the physical state of dipalmitoyllecithin multibilayers.

L. Ernster, K. Asami, K. Juntti, J. Coleman, and K. Nordernbrand then describe the interaction of a protein that inhibits mitochondrial ATPase with submitochondrial particles. The authors suggest that the inhibitory protein may regulate energy transfer between the respiratory chain and the A TP generating system. E. H. Eylar then gives an account of the myelin membrane and of the basic proteins found in it. Their arrangement in the myelin membrane is discussed and their role in human demyelination diseases assessed. This is followed by an article entitled "Regulation of Pancreatic Phospholipase A2 by Different Lipid-Water Interfaces" by M. C. E. van Dam-Mieras, A. J. Siotboom, H. M. Verheij, R. Verger, and G. H. de Haas. This enzyme contains a specific "interface recognition site" that interacts with lipid-water interfaces. The site was studied in various modified phospholipases in which the N-terminal alanine had been substituted by other amino acids or by amino acid chains.

C. R. Hackenbrock then presents a comprehensive account of the molecular organization and fluidity of the mitochondrial membrane. In view of the importance of the subject, it may be of interest to quote in full Hackenbrock's assessment of current views on mitochondrial organization:

The picture that emerges is that of a highly effective concentration of integral proteins partitioned in a polar bilayer phospholipid environment of relatively low viscosity and high fluidity. The polar environment provides for a precise vertical

378 Some Recent Books in Cell Biochemistry and Biology

orientation of the integral metabolically active membrane proteins. The fluid environment provides for lateral translational and rotational mobility of the integral metabolically active membrane proteins which can diffuse laterally, depending on their specific metabolic role, either independent of or in association with other integral proteins.

J. N. Hawthorne then gives an account of the role of triphosphoinositide in myelin and plasma membranes in Ca2+ ion binding. The author also discusses the role of phosphatidyl inositol and phosphatidic acid in the release of neurotransmitters. K.-A. Karlsson then gives an account of sphingolipids in cell-surface membranes. He surveys the ceramide composition of a number of cells, postulates that sulfa tides and acid phospholipids are involved in the action of the Na + -K+ pump, and assesses the role of sphingolipids as surface antigens. J. A. Lucy then describes the use of the hen erythrocyte membrane as a model system for studies on membrane fusion. He suggests that "fusogenic lipids" alter the polar regions of membrane phospholipids and that as a result the membranes become more permeable to Ca2+ ions. This sets off a series of changes that ultimately favor membrane fusion. I. Lundstrom then discusses the structure and electrical properties of lipid-water systems. The lateral conductivity, Raman spectra, and light-scattering properties of an artificiallamellar lipid-water system were examined and the artificial system compared to biomembranes. A. Tardieu, C. Sardet, and V. Luzzati then present the results of X-ray scattering studies of bovine rhodopsin. Using a detergent-rhodopsin complex, they concluded that thin elongated rhodopsin molecules (more than 80 A long) span the flat detergent micelle.

P. BrUlet, G. M. K. Humphries, and H. M. McConnell then discuss the immunochemistry of model membranes containing spin-labeled haptens. They include studies on spin-label hapten resonance spectra and on complement fixation. Next, M. D. Hous)ay, A. Johannson, G. A. Smith, T. R. Hesketh, G. B. Warren, and J. C. Metcalfe present an article on the coupling of the glucagon receptor to adenylate cyclase. They describe various models that have been proposed to explain the relationship between hormone receptors and adenylate cyclase. This is followed by an account of peptide ionophores by Yu. A. Ovchinnikov. In particular, he discusses valinomycin, enniatin B, gramicidin, and their analogues. The article includes detailed biochemical and physicochemical studies of various ionophores and of their interaction with liposomes.

G. D. Eytan, G. Schatz, and E. Racker then describe experiments on the incorporation of integral membrane proteins into liposomes. They give details of the sequential insertion of various mitochondrial multienzyme complexes and compare the results of these in vitro systems with what is known of mitochondrial membrane assembly in vivo. P. R. Cullis, B. de Kruijff, A. E. McGrath, C. G. Morgan, and G. K. Radda then discuss lipid asymmetry and molecular motion in biomembranes with particular reference to the behavior

Some Recent Books in Cell Biocbemistry and Biology 379

of chromaffin granules. They describe various model systems and also give details of the interesting technique in which the decay times of positrons introduced into the lipid matrix are measured. O. Renkonen, M. Pesonen, and K. Mattila then give an account of the oligosaccharides of the membrane glycoproteins of the Semliki Forest virus. They propose a complex structure involving a branched arrangement of N-acetyl neuraminic acid, mannose, and Nacetyl glucosamine groups attached to a peptide framework.

A. M. Scanu then discusses the use of phospholipases as probes for circulating lipoproteins. In particular, the effects of lipolytic enzymes of low-density lipoprotein LDL2 and high-density lipoprotein HDL} are described. Next, S. J. Singer gives an account of the fluid mosaic model of membrane structure. He considers various thermodynamic aspects of membrane fluidity and also discusses the molecular asymmetry of membranes and the mechanism of transport of hydrophilic ligands through membranes. J. C. Skou discusses the coupling of the passive flux of N a + and K + ions to the active transport of these ions and analyzes the role of the "i-site" (inside) and the "o-site" (outside) in Na+ and K+ transport. The relative merits of one-site and two-site models are discussed. W. Stoeckenius, S-B. Hwang, and J. Korenbrot then give an account of proton translocation by bacteriorhodopsin in lipid vesicles. In intact cells it is difficult to examine the relationships between light absorption, proton gradients, and membrane potential. Use of a reconstituted model system overcomes these difficulties and the article assesses the results of studies of these parameters in the model system.

C. Tanford then discusses the state of association of membrane proteins. By using appropriate detergents, membrane proteins may be isolated as a part of a detergent micelle in a form similar to that in the membrane, and then subjected to classic physicochemical techniques. This approach is used to study the subunit structure and organization of cytochrome bs and sarcoplasmic reticulum Ca2+ ATPase. The next chapter on membrane electrostatics is by the late H. Traiible and deals comprehensively with surface electrostatics, the effect of electrostatic forces on membrane structure, ion pulses in membranes, electrostatic regulation of membrane phase separations, and electrostatic coupling between two layers of a membrane. The book ends with a chapter by G. Vanderkooi and J. T. Bendler entitled "Dynamics and Thermodynamics of Lipid-Protein Interactions in Membranes." The free energy of mixing of lipids and membrane proteins is discussed, as well as the athermal entropy of mixing. Hamaker constants are given for lipid-lipid, protein-lipid, protein-protein interactions, and the use of the generalized Guggenheim method is discussed. The book includes a short subject index.

As the above detailed survey shows, Structure of Biological Membranes covers a broad canvas and contains detailed and authoritative chapters by established experts. However, as the reader may well have noticed in the above survey, there has been no clear attempt to group related chapters, so that one


has to leap rather suddenly from one area of membrane biology to another. In fact, many of the chapters are quite closely related, and had the editors departed from the rigid method of listing the articles in simple alphabetic order, a more stimulating and readable book would have been presented. Nevertheless the publication is a most useful addition to the now regrettably overwhelming literature on biological membranes.

The next book to be discussed is Cell Motility by H. Stebbings and J. S. Hyams and contrasts strongly with the rather massive multiauthor text just reviewed, being written by only two authors and only running to 192 rather small pages. This interesting and well-presented book is part of an excellent Longman series edited by I. D. J. Phillips entitled Integrated Themes in Biology, which includes such useful monographs as Mitochondria by P. A. Whittaker and S. M. Danks.

In the preface the authors stress the great current interest in cell motility in all its aspects as a result of the appreciation of the widespread occurrence of microtubules and microfilaments. The book begins with an account of the structure and function of striated muscle, as the most highly specialized system capable of converting chemical energy in the form of ATP into useful mechanical work. The structural organization of striated muscle is described, and the biochemistry of muscle proteins surveyed. The now famous sliding filament theory is briefly but elegantly presented, and there is a short account of the control of contractility. The next chapter deals with microtubules and microfilaments. The morphology of microtubules is discussed in detail and there is a full account of microtubule biochemistry including the heterodimer model, the mode of action of colchicine, the Vinca alkaloids, and other spindle poisons, as well as what we know of microtubule assembly both in vivo and in vitro. The identification and morphology of microfilaments is then described and the role of actin discussed. Evidence for the existence of cytoplasmic myosin is assessed and the general role of contractile proteins discussed. There is an account of actin-membrane associations, and the chapter ends with a critical analysis of the efficacy of cytochalasin B as a specific disrupting agent for microfilaments.

The next chapter deals with cilia, flagella, and axostyles. After an account of ciliary and flagellar movement, there is a full treatment of the structure and biochemistry of the axoneme and of the reactivation of cilia and flagella. The chapter ends with a description of the fascinating axostyle found in some anaerobic flagellates; this is a ribbonlike bundle of an ordered arrangement of singlet microtubules and is an ideal model system for the study of ciliary and flagellar movement. (In passing, it may be mentioned that these elegant and informative observations on the axoneme provide a good example of the impossibility of predicting the scientific usefulness of a given field of study in advance. The organism mentioned is the anaerobic flagellate Saccinobacculus, which inhabits the hindgut of the wood-eating roach Cryptocercus. This appar-

Some Recent Books in Cell Biochemistry and Biology 381

ently obscure organism may, however, provide information of central importance to cell biology.)

The authors then deal with cell movements and contractile proteins. They describe the acrosome reaction in invertebrate sperm, shuttle streaming in slime moulds, amoeboid movements, and brush-border contraction. The next chapter deals specifically with movements within cells. The role of microtubuies in intracellular transport is extensively and critically discussed, and there is an account of protoplasmic streaming and the possible role of microfilaments in this process. The next chapter deals with mitosis and cytokinesis. The structure of the mitotic spindle is described, as well as the distribution of spindle microtubules. The evidence for the presence of actin in the spindle is assessed and there is a useful account of the "sliding microtubule model" as a mechanism of mitosis. The chapter ends with a brief account of cytokinesis and of the contractile ring. The book ends with a short description of contractile movements in ciliates such as Stentor and Vorticella and of various "pendulous" and "gliding" movements of such sporozoans as Selenidium and Gregarina.

There is a short subject index and each chapter is amplv furnished with detailed references and a basic list of suitable review articles and monographs. The text is admirably illustrated throughout with high quality diagrams and electron micrographs. Cell Motility clearly shows that we have now entered a phase of dramatic unification in our understanding of the mechanics of a wide range of biological processes. The terms filament, tubule, actin, and myosin occur again and again throughout the text, leaving one with the overwhelming impression that just as a relatively few amino acids can produce an infinite variety of proteins, so can a few contractile and fibrous elements produce a wealth of contractile and motile activities. Stebbings and Hyams's book is an excellent and stimulating introduction to this exciting new synthetic phase of cell biology.

The next book to be discussed is Essentials of Cell Biology by Robert D. Dyson. This is an undergraduate textbook, and the author states in his preface: "The objective of this book is to present a unified description of cellular structure and function at the introductory level." The first chapter surveys the cell theory, the main structural features of prokaryotic and eukaryotic cells, and techniques for investigating cell structure. Some excellent electron micrographs show the main features of the major cellular elements and illustrate the section on methods. The next chapter is entitled "Membranes and Macromolecules" and in a most original way deals with the chemistry of lipids, proteins, and carbohydrates at the same time as the main features of membranes and macromolecular assembly are discussed. The student is therefore encouraged to view the study of the chemistry of macromolecules as being intimately connected to an understanding of the way in which these macromolecules come together to form organized cell structures.

The next chapter discusses bioenergetics and cellular homeostasis. An


account of basic thermodynamics leads on to an explanation of enzyme action, metabolic feedback, and ATP-coupled reactions. Glycolysis is treated as an example of an integrated metabolic pathway and there is a brief account of biological oxidation. Thus thermodynamics, enzymology, and metabolic pathways are not presented as separate topics, as is so often the case, but are examined in an interlinked and intellectually satisfying manner. Genes and their regulation are the next major topic, and here we have accounts of general genetic concepts, the chemical nature of the gene, DNA structure, the nuclear envelope, chromosomes, the genetic code, and transcription. Again, the various subsections interlock naturally. From transcription the author proceeds to an account of protein biosynthesis, and via a description of the endoplasmic reticulum and Golgi apparatus, to a survey of the main aspects of secretion.

The plasma membrane is then discussed in a chapter that includes accounts of cellular recognition, membrane transport, active transport, endocytosis, lysosomes, membrane turnover, and biosynthesis. Mitochondria, chloroplasts, and peroxisomes are discussed in one chapter that includes accounts of the Krebs' cycle, electron transport, oxidative phosphorylation, photosynthesis (light and dark varieties), peroxisomal metabolism, and organelle evolution.

Excitability, contractility, and motility are discussed next in a chapter that includes accounts of muscle contraction, nerve, and muscle excitability, excitation-contraction coupling, cilia, and flagella. Next follows a chapter on DNA replication, mitosis, meiosis, and cytokinesis. The final chapter gives an account of cellular differentiation. There is an excellent glossary of terms at the end of the book, followed by a medium-size subject index. Each chapter contains rich illustrations with excellent structural diagrams and well-chosen electron micrographs and ends with a well-set-out reference list subdivided according to subject. All in all, reading Essentials of Cell Biology is a most pleasurable experience. Its most admirable feature is the way in which the "classic" sections on glycolysis, the Krebs' cycle, enzymology, and protein structure are neatly woven into the descriptions of organelle structure and function. Let us hope that students who have based their studies on this book will come away with as integrated a view of the cell and its biochemical constituents as its author has.

Dyson's other book (Cell Biology: A Molecular Approach) was reviewed in detail in Sub-Cellular Biochemistry, Volume 3 (pp. 371-372), and the review ended with the statement that it is " ... a most excellent contribution to the educational literature in the field and can be recommended without hesitation to all cell biologists." We have now received the second edition for review, and it is similar to the first, only better. The author has taken into account many readers' suggestions and has more on membrane structure and function, new material on contractility and motility, plus some new clinically


relevant material. There is no doubt that Essentials of Cell Biology and Cell Biology: A Molecular Approach represent two major landmarks in the evolution of student texts in cell biology.

The final book to be considered in this section on cell and membrane biology is Volume 8 of Methods in Membrane Biology edited by E. D. Korn. The first article is by P. Zahler and V. Niggli and surveys the use of organic solvents in membrane research. The authors discuss the difficulties of working with amphipathic proteins, as a medium suitable for the polar regions is unsuitable for the nonpolar parts of the molecule, and vice versa. The "ideal" liquid medium for the fractionation of membrane proteins would tolerate both the charged and hydrophobic regions without causing extensive secondary unfolding of the molecule. In many ways detergents answer this need-unfortunately their use results in the isolation of detergent-membrane protein complexes, and it is often difficult to distinguish which properties are attributable to the protein and which to the detergent. Zahler and Naggli survey the wide variety of organic solvents that have been used to overcome these difficulties and give useful details of their physical properties. They also describe the application of the most commonly used solvents to membranology. They include a useful appendix that describes extraction methods for total lipids, serum phosphatides, and membrane proteins. The chapter thus usefully brings together data on a technique that is gaining increasing importance in the investigation of hydrophobic interactions in membranes.

The next article is by R. A. Klein and P. Kemp and surveys in a most comprehensive way recent methods fdr the elucidation of lipid structure. As our understanding of the molecular organization of membranes improves, it becomes more important to have a precise knowledge of the chemical structure of membrane lipids. This article shows how powerful contemporary analytical techniques have been applied to the investigation of lipid structure. The authors survey separation techniques, methods for lipid identification, the analysis of stereoisomers, mass spectrometry, and proton and carbon-I 3 nuclear magnetic resonance spectroscopy. There is also a useful section on artifacts and contaminants. The next chapter by M. Kates is entitled "Synthesis of Stereoisomeric Phospholipids for Use in Membrane Studies." Chemical studies on lipids clearly depend on the supply of uncontaminated lipids of known composition and stereochemical configuration. Regrettably, lipids isolated from natural sources often do not meet these requirements, so that the use of lipids chemically synthesized by unambiguous routes is important in lipid (and hence membrane) research. Procedures for the synthesis of a wide range of phospholipids are given, and there is also information on the synthesis of alkyl and alk-I-enyl ether analogues of phospholipids (e.g., various plasmalogens). One is struck on reading Kates's article by the great variety of natural phospholipids and the superficiality of the view that only phosphatidyl choline, serine, and ethanol-


amine are of interest. The article should help considerably to improve the precision of studies on the chemical constitution of membrane phospholipids.

The last chapter of this useful volume is by B. J. Gaffney and S-c. Cben and deals with spin-label studies of membranes. The authors begin with a historical treatment of the application of electron paramagnetic resonsance spectroscopy to the study of biomembranes; the various nitroxides used are described and data are presented of diffusion constants of lipids in membranes. There is a useful theoretical section on the analysis of paramagnetic spectra and of the effects of molecular motion on these spectra. The application of these techniques to the study of lipids and proteins in membranes is then surveyed and the results compared to those obtained by other physical techniques such as freeze-fracture electron microscopy and X-ray diffraction.

Volume 8 of Methods in Membrane Biology thus continues the tradition of this series of providing detailed, useful, and expert surveys of methodology in membrane research.

2. GENETICS AND VIRUSES

The first book to be considered in this section is The Biochemistry of Viruses by S. J. Martin. This is a short introductory textbook" ... designed to provide a rapid overall picture of virology at the molecular level," and it succeeds admirably in its task. It is written in essay style, and the various diagrams, structural formulas, and plates are well chosen and clear. After a brief history of virology, the author discusses the classification of viruses. Methods of quantitatively assaying viruses are then described and an account is given of various methods of purification of viruses. The architecture of viruses is then discussed and the author explains how the fact that they appear to be formed by self-assembly greatly limits the range of morphologies encountered in viruses. Essentially two main repeating patterns are used-helical symmetry, which gives rise to rod-shaped viruses, and icosahedral symmetry, which gives rise to spherical or spheroidal viruses. The author surveys the molecular organization of turnip yellow mosaic viruses, picornaviruses, including adenovirus, complex viruses, such as paramyxoviruses, and finally the bacteriophages.

The next chapter deals with the "Strategy of Virus Infection" and describes the physical processes involved (e.g., the attachment of the virus to the host, the penetration of infective material, and the "uncoating" of the virus); the main biochemical features of viral infection are also given. The process of infection is described by the author in an excellent turn of phrase as a "biochemical coup d'etat," and the chapter shows in full detail the elegant devices whereby the imposter takes over the informational and replicative


machinery of the host and subverts it for the manufacture of viral particles. The author, with another graphic expression, calls viruses "Invaders of the Genosphere" and draws analogies between their catastrophic effects and those of man, who he describes as "Invader of the Biosphere." The last chapter discusses the evolution of viruses, the development of vaccines and research into antiviral chemotherapeutic agents. The possible benefits, but also the potential hazards, of genetic engineering involving viruses are discussed .

. All in all, The Biochemistry of Viruses is pleasant and informative reading with a stimulating seasoning of humor and philosophy. It is a most useful introduction to the world of viruses and successfully captures the author's sense of enthusiasm and wonder at these sophisticated biological hijackers.

The next book in this section is Extranuclear Genetics by G. Beale and J. Knowles. This excellent monograph is aimed at a wide audience, merely assuming that the readers have an elementary knowledge of molecular biology and basic genetics. The preface contains the all-too-common "exclusion clause" viz.:

Organelle genetics, and even more the genetics of bacterial plasm ids, are such rapidly advancing areas of research that we cannot hope our account is completely up-to-date even at the time of writing . ... [italics added]

Thus even as we write we are out of date. Worse, we become out of date as we read! If we have to journey through hundreds of pages of multiauthor, jargonreplete, pseudolegalistic prose that makes up so much of the contemporary "literature" our out-of-datedness increases with our sense of confusion and frustration. How refreshing, therefore, to read a book that has a mere 120 or so pages, with sufficient but not excessive tables and figures, and-glory of glories-is written in comprehensible English, not moleculobiological newspeak.

The introduction sketches the main features of non-Mendelian or extranuclear inheritance and reminds us forcibly how until recently genetics had been dominated by the" ... extremely rigid, mechanistic concepts of classical geneticists .... " Thus the authors quote Morgan as having written in 1926: "The cytoplasm may be ignored genetically." It is not generally realized, with respect to this remark by Morgan, that there was considerable evidence for non-Mendelian "aberrant" genetic systems early in 20th century, and it is quite false to regard the development of the science of cytoplasmic inheritance as having occurred after the discoveries of mitochondrial and chloroplastal DNA.

The authors then deal with the mitochondrion, briefly surveying the structure and properties of mitochondria and then giving details of the mitochondrial biosynthetic apparatus. There is an account of the mechanism of repli-


cation of mitochondrial DNA and of the principal phenotypic effects of changes to mitochondrial genes (petiteness in yeast, poky variants in Neurospora, and more recently various drug-resistant phenomena in a variety of cell types). There is a full section on recombination of mitochondrial genes and also of the recent advances in the mapping of the mitochondrial genome. The particular complexities that arise from two genetic systems in mitochondrial biogenesis are then discussed, and the chapter ends with a short account of the kinetoplast and kinetoplast DNA in trypanosomes and similar organisms. The next chapter on the chloroplast follows the same general design as the chapter on mitochondria. The main features of the findings with the two organelles are strikingly similar-each has its own limited biosynthetic apparatus, each relies on the interaction of two genetic systems, and the major gene products in both organelles are ribosomal RNAs and hydrophobic membrane-bound proteins. There are differences in detail, however (e.g., the chloroplast can synthesize a soluble protein, fraction I protein large subunit, and appears to have greater coding capacity than the mitochondrion); nevertheless the similarities between the two systems are very thought-provoking.

There follows a chapter on bacterial plasmids. In view of the absence of a defined bacterial nucleus enveloped in a limiting envelope or membrane, it is not immediately obvious that plasmids should be considered in a book entitled Extranuclear Genetics. However, the authors justify this on the grounds that one can regard the main mass of bacterial DNA as constituting a chromosome (albeit not partitioned in its own enevelope), and the plasm ids can be looked on as "accessory" genetic elements. The authors stress that the plasmid is an excellent model for the study of extrachromosomal phenomena. After a brief description of F, R, and Col plasmids, there are full accounts of the replication of plasmid DNA, plasmid transfer, recombination, and mapping. There is also a useful section on artificial plasmids and on genetic engineering using plasmid vectors.

The next chapter is entitled "Endosymbionts and Viruses as Agents of Extranuclear Heredity" and contains some fascinating examples of external agents that can influence extranuclear inheritance. The examples given include, inter alia, the kappa particles of Paramecium aurelia; a rickettsial ike microorganism apparently responsible for incompatibility between certain mosquito strains; various spiroplasmalike symbionts and their associated viruses that appear to be involved in the production of abnormal sex ratios in the offspring of certain species of Drosophila; the "sigma" virus, which confers CO2 sensitivity on some strains of Drosophila melanogaster; and finally RNAcontaining viruslike particles in "killer" strains of Saccharomyces cerevisiae and the fungus Ustilago maydis. The authors stress that these agents, originally thought to be examples of an unusual biological phenomenon, probably represent a widespread process whereby extraneous elements interfere with the


genetic system of a host. Thus kappa was at first thought to be a special case in P. aurelia. It now transpires that almost every species of ciliate as well as many other protozoa have symbiont-bearing individuals in their midst. To reinforce this picture of the diversity of extrachromosomal systems, the next chapter lists some miscellaneous examples of extranuclear inheritance which have not yet been associated with a known organelle or with a defined physical agent such as a virus. These examples include various incompatibility phenomena in fungi, cytoplasmic supressors in yeast, cytoplasmic male sterility in plants, and various features of nucleocytoplasmic interaction in protozoa.

The final chapter briefly surveys the main arguments and discusses various theories of organelle evolution, the origin of eukaryotes, and their relationship to prokaryotes. There is a manageable well-selected reference list and a brief subject index. In general, Extranuclear Inheritance is a most useful monograph that should stimulate awareness of many interesting, important, and neglected phenomena in cell biology, and should help us see the genetic role of the nucleus in its proper perspective.

The third book to be discussed in this section is The Phylogeny of Human Chromosomes by H. N. Semlnez. Age-old problems can be looked at with gleaming new tools-part of the fascination of this stimulating little book is to see how modern cytological and biochemical techniques have been brought to bear on the problem of the origin of human chromosomes and hence to the fundamental question: whence came man? The first section of the book deals directly with the question of the origin of man, surveying the fossil record and presenting current views on the classification of the Hominoidea. According to Goodman (1975), the reader may be pleased to know, he or she belongs to the population of living beings classified under Homo sapiens, belonging to the superfamily Hominoidea, the family Hominidae, and the subfamily Homininae. In our subfamily are included the chimpanzee and the gorilla.

The next section deals with cytotaxonomy and the evolution of man and the great apes. Their chromosomes are compared in detail, both in number and also via the various banding methods (e.g., G-, R-, and Q-banding). There is a detailed account of chromosomal aberrations (heteromorphisms) in man and the great apes and also of chromosomal rearrangements by inversion and tel om eric fusion. From this analysis one can suggest an ancestral chromosomal complement of the Hominidae, details of which are given. The section ends with a short account of the relationship between speciation and chromosomal rearrangement.

The third and largest section is oriented toward biochemistry and molecular biology and deals with topics such as DNA sequence studies, satellite DNAs, palindromes, DNA replication, eu- and heterochromatin, and so forth. The new approaches have increased our understanding of the chromosome in molecular terms, as distinct from the more "bulk" morphological and staining


studies of classic chromosomology. It is difficult to tell from Seuanez's account, however, to what extent the new techniques have solved the problem of man's origin. One has the impression that the new methods have often merely raised new and more complicated questions. This is perhaps best summed up by a direct quotation from the last paragraph of the book:

... I feel it is necessary to emphasize how far we still are from having a clear idea of how our own species evolved. With the development of new techniques in the years to come let us hope that substantial information will be obtained which will enlarge the limited understanding we already have of our own nature and origins. Until then, and perhaps even then, the basic question of What is man? will remain still unanswered.

There is little doubt that The Phylogeny of Human Chromosomes is a most useful tool in the pursuit of the answer to this intriguing question.

3. MUSCLE AND Ca2+ TRANSPORT

Recent years have shown an increasing appreciation of the role of CaH in muscle function and it is therefore appropriate to discuss books on muscle and on CaH transport in the same section.

Smooth Muscle edited by E. Biilbring and T. B. Bolton is a part of Volume 35 of the British Medical Bulletin and is directed mainly at "physiologists, pharmacologists, and clinical pharmacologists." Nevertheless, much of the material in it would certainly be of interest to readers of SUBCELLULAR BIOCHEMISTRY. The first article by G. Gabella discusses smooth muscle cell junctions and the structural organization of smooth muscle in relation to its contractile ability. He deals with all aspects of smooth muscle organization, including the intercellular materials (collagen, elastin, and possibly mucopolysaccharides), the arrangements of the dense bands, small cell membrane invaginations called caveolae, intermediate junctions, gap junctions (nexuses), links between cells, and the intercellular material and, finally, the physical arrangement of myofilaments. The article, although only five pages long, is an excellent survey of the mean features of the organization of smooth muscle.

S. V. Perry and R. J. Grand then survey what is known of the biochemistry of smooth muscle contraction. They discuss our knowledge of contractile proteins in smooth muscle and deal with other constituents, such as dense bodies, 10-nm filaments, a-actinin, the M protein, and filamin. They also discuss the regulatory systems of smooth muscle (including the role of troponinlike systems, myosin phosphorylation, and calmodulin), as well as the contractile process. The authors compare the state of our knowledge of smooth muscle biochemistry with that of skeletal muscle. A. F. Brading then discusses the maintenance of the ionic composition of smooth muscle and gives an account


of the Na-K pump, Na-linked ion movements, and the regulation of Ca2+ and Cl- levels. A model for Na + exchange is presented, and this is discussed in relation to calcium movements, plasma membrane vesicles, and the general functioning of the sarcoplasmic reticulum.

Other chapters in Smooth Muscle deal with various aspects of membrane physiology, blood supply, innervation, peristalsis, drug receptors, cholinergic and adrenergic mechanisms, as well as the mode of action of prostaglandins. In their introduction, the editors survey the achievements of recent research in the subject, but also stress the outstanding problems. Many of these arise from the fact that there appear to be considerable differences in the properties of smooth muscle depending on where it occurs. There is clearly much work to be done in this field, with many of the tools successfully fashioned in the investigation of skeletal muscle. This publication makes a useful contribution to such an endeavor by bringing together under one cover specialized chapters of manageable size that cover all the major aspects of smooth muscle structure and function.

Calcium Transport and Cell Function, edited by A. Scarpa and E. Carafoli, is a large tome running to morc than 650 pages. It consists of the published proceedings of a conference of the same title held by the New York Academy of Sciences in late 1977. As there are nearly 60 papers in the book, plus a report of the discussion that followed, it is not possible to give an account of each topic covered. Instead, the main sections of the book will be surveyed and some papers of particular relevance to cell biochemistry discussed in a little greater detail.

The book begins with some introductory remarks by A. Scarpa and with an account by D. W. Urry of the basic chemistry of calcium and of its interaction with membranes. Part I then deals specifically with the measurement of calcium and includes accounts of microprobe X-ray analysis of calcium, calcium-sensitive electrodes, the use of photoproteins such as aequorin and obelin, metallochrome indicators, and finally "Extended X-ray Absorption Fine Structure" (EXAFS) studies. One is struck by the range and sophisti,cation of techniques now available for the study of calcium in biological systems-it is an interesting question as to whether our present appreciation of the central importance of calcium in biology is a result of the development of these techniques, or vice versa.

The next part deals with the interaction of calcium with subcellular organelles. H. J. Schatzmann and H. BUrgin begin with an account of calcium in human red blood cells, which includes a detailed discussion of the Ca-pump ATPase. A. N. Martonosi, T. L. Chyn, and A. Schibeci then give a short survey of calcium translocation in the sarcoplasmic reticulum and present a hypothesis for the regulation of the synthesis of Ca2+ ATPase mRNA by nuclear repressor proteins with a high affinity for Ca2+. A. L. Lehninger, B.


Reynafarje, A. Vercesi, and W. P. Tew then discuss the transport and accumulation of calcium in mitochondria. They give an account of high-affinity Ca2+ binding sites and of the possible participation of phosphocitrate in the storage of calcium phosphate in mitochondria. L. Moore and I. Pastan then describe energy-dependent calcium uptake in fibroblast microsomal fractions and discuss the interaction of calcium pumps in the plasma membrane, mitochondria, and endoplasmic reticulum. M. P. Blaustein, R. W. Ratzlaff, and N. K. Kendrick, then give an account of intracellular calcium regulation in presynaptic nerve terminals and survey the properties of nonmitochondrial A TP-dependent calcium storage systems in disrupted nerve terminals.

After Part II there is a report of a panel discussion on the previous two parts and this includes several contributions on the sarcoplasmic reticulum in relation to calcium metabolism and on mitochondrial transport, accumulation, and storage of calcium. Part III deals with the regulation of intracellular calcium and includes accounts of regulation in giant axons, mitochondria, and barnacle muscle fibers, as well as discussion of the role of calcium in inter- and intracellular communication.

Part IV discusses the hormonal control of calcium metabolism and includes an account of the action of the widely used ionophore A23187. There follows a panel discussion on Parts III and IV, which includes papers on the calcium channel in the sarcoplasmic reticulum, calcium transport in bone and intestinal cells, the physiological regulation of calcium metabolism, and the role of prostaglandins. The next part deals specifically with the role of calcium in muscle contraction and the final part is concerned with calcium in vision and secretion. The final panel discussion includes some more material on muscle action, vision and the role of calcium in secretion.

As may be judged from the above account, research into calcium now touches on a very wide range of fundamental biological processes. Calcium Transport and Cell Function therefore does a useful service in bringing together under one cover a representative sample of current experimental approaches to the topic. The book demonstrates the range of sophisticated techniques now being used and is a useful source of reference for those wishing to enter this rapidly expanding field of research.

4. GENERAL BIOCHEMISTRY

The next three books to be discussed (by J. W. Suttie, R. C. Bohinski, and D. E. Metzler, respectively) are all educational texts and illustrate the various approaches that may be adopted to the teaching of general biochemistry.

Introduction to Biochemistry, by J. W. Suttie, is an introductory text


designed around a one-semester elementary biochemistry course taught at the University of Wisconsin. The author has been careful to restrict the amount of material in the book to that which a student can reasonably be expected to assimilate in one semester. The main division of the book is into (1) The Chemistry of Biological Material, (2) Dynamics and Energetics of Biochemical Systems, (3) Energy Production in Biochemical Systems, (4) Energy Utilization in Biochemical Systems, and (5) Metabolic Control. After an account of pH and buffers, the main features of cell structure are presented, and this is followed systematically by chapters on carbohydrate, lipid, protein, and nucleic acid chemistry. An account of the structure and function of enzymes is followed by a chapter on biochemical energetics. A description of methods for studying intermediary metabolism is then followed, equally logically, by chapters on carbohydrate, lipid, protein, and nucleotide metabolism and on the tricarboxylic acid cycle. The section on energy utilization has chapters on photosynthesis, and the biosynthesis of carbohydrates, lipids, and nitrogencontaining compounds, including nucleic acids. The last part surveys the major metabolic pathways and discusses the various ways in which metabolism is controlled.

The student is thus led systematically through the subject and the text is well illustrated with clear diagrams, set problems and short reading lists. The book is clearly a well-thought-out and thoroughly tested teaching text and one would be surprised if students had serious difficulties with it. In contrast to Dyson's books discussed above (which one may call "cell orientated"), Introduction to Biochemistry belongs to the genre of what, for lack of a suitable term, may be called "Carbohydrate-Lipid-Protein-Nucleic Acid (CLPN) orientated" texts. In such books, the student is first asked to divide living matter into these four great chemical subdivisions and is then systematically taught their structure, how they are degraded, and how they are synthesized. The usual cement that binds together the four categories is bioenergetics. In such texts, cell organization is generally included as a subsection of one of the parts and is not treated as the core of the subject. Only the future will tell us whether the cell-structure-orientated or CLPN-orientated approaches are more fruitful-certainly Suttie's well planned book is an excellent example of the latter approach.

Modern Concepts of Biochemistry, by R. C. Bobinski, is a larger text. It may be used for one-semester courses, but it has sufficient material for use in longer courses. After an interesting introduction in which the author marks out the great range of studies included under the simple term "biochemistry" and also includes some salutory philosophy about the complexity of life, the first chapter deals with cellular organization; the structure of the major organelles is surveyed. There then follows a chapter on methodology, and this is followed by a treatment of pH, buffers, hydrogen bonding, and hydrophilic interactions.


Three chapters follow on amino acids and peptides, proteins, and enzymes, respectively, after which follows an account of nucleotide and nucleic acid chemistry and RNA, DNA, and protein biosynthesis. Carbohydrates, lipids, and biomembranes are then discussed, and this leads on to chapters on the citric acid cycle, oxidative phosphorylation, and photosynthesis. The last two chapters deal with lipid metabolism and the metabolism of nitrogen-containing compounds, and the book is completed with appendixes on isoenzymes in clinical diagnosis and on human genetic disorders.

Each chapter is richly amplified with diagrams, structural formulas, tables of data, and literature guides. The general impression is of a most readable and well-thought-out text. Although concepts are presented wherever possible, parts of the book are rather replete with factual material. For example, there is a full-page diagram of the metabolic pathways for squalene biosynthesis, and some students may have difficulty in deciding what is essential and what is merely reference material. Nevertheless Bohinski's book is an excellent attempt to reveal a unifying thread in biochemistry and to deal with the manifold problems associated with teaching a subject that is in a phase of rapid change and development.

The third of the educational texts considered here, Biochemistry: The Chemical Reactions of Living Cells, by D. E. Metzler, stands in a class of its own. The aim of the book is best explained by a direct quotation from the preface:

... rather than dividing biochemistry into segments centered around specific chemical compounds such as proteins, nucleic acids, lipids and carbohydrates, I treat chemical reactions of cells as a primary theme. While stressing biological concerns, I try to trace all physiological phenomena back to the underlying chemistry.

The book therefore stands at the opposite pole to those by Dyson discussed above. Instead of examining living processes as a function of cellular organization, they are looked on as the product of highly specialized, integrated, and spatially organized chemical reactions. A cursory inspection of the contents of the book shows how cleverly the author has organized his material in pursuit of this aim. Clearly, the "chemocentric" analysis of cellular function is a most rewarding exercise. Nevertheless there is a nagging doubt in the reviewer's mind as to whether such an analysis, even in full and minute detail, will provide us with a satisfactory answer to many of the current problems of cell biology. Is the cell merely a complicated machine made up of thousands of chemically interlocking parts, or has it some higher properties, not necessarily predictable from a knowledge of these parts?

Metzler's book is on a much larger scale than the two books just discussed, running to more than 1100 pages. Because of the originality of the author's approach, the main chapter divisions bear no relationship to the classical layout


of so many biochemical textbooks. The author first sets "The Scene of Action." This includes an account of the main features of prokaryotic and eukaryotic cell structure and of the evolution of complex organisms. The next chapter begins with a discussion of the structure of small molecules and leads on to an account of the chemistry of amino acids, proteins, mono- and polysaccharides, nucleic acids, and lipids. The chapter includes a discussion on ions of biological importance and of studies on the chemical composition of cells. There is then a detailed treatment of the energetics of biochemical reactions and this leads to a discussion of the forces that act between molecules of biological importance. There is also an account of the self assembly of complex macromolecular systems, including bacteriophage.

"Membranes and Cell Coats" is the title of the next chapter in which the structure and function of a wide range of biomembranes are surveyed. Enzymes are then discussed, and a full account is given of enzyme kinetics, the various proposed mechanisms for enzyme action, and the regulation of enzyme activity. The next chapter surveys the types of reaction catalyzed by enzymes, classifying these reactions in strictly chemical terms. Coenzyme structure and mode of action are then discussed and this leads onto an account of the "organization of metabolism." Catabolic pathways are described first; then there is an account of electron transport and oxidative phosphorylation, plus a description of various oxygenases and hydroxylases. Biosynthesis is discussed in a generalized manner (i.e., "How molecules are put together"), and there follows a chapter on some selected specific pathways in carbohydrate and lipid metabolism.

Typical of the originality of the author's approach, and of the way he has tried to break out of the straitjacket of conventional texts, is the fact that the treatment of photosynthesis is included in a more general chapter entitled "Light in Biology." This includes an account of the electromagnetic spectrum, circular dichroism, optical rotatory dispersion, fluorescence, phosphorescence, and the biochemical processes of vision. Thus the reactions of photosynthesis are perceived by the student as a special case of the more general problem of the interaction of biological molecules with the whole spectrum of electromagnetic radiation. There is a chapter on the metabolism of nitrogen-containing compounds followed by one on biochemical genetics, nucleic acid synthesis, and protein synthesis. The final chapter is on growth, differentiation, and chemical communication between cells.

An original feature of the book is the inclusion of special "boxes" interspersed throughout the text, and printed over a green background. These are not chapter synopses, but contain ancillary material on topics such as vitamins, essential elements, metabolic diseases, antibiotics, and physiological chemistry. By choosing appropriate patterns of boxes, as suggested by the author, students studying special aspects of biochemistry can amplify the main material in the


text proper with the details in the boxes. The nonspecialist student aiming at a fundamental treatment of the subject can ignore the boxes altogether. The text has a high standard of presentation, and each chapter includes useful references and a series of helpful study questions. The book includes an appendix on the construction of molecular models as well as a detailed and fully comprehensive subject index.

Biochemistry: The Chemical Reactions of Living Cells is thus an original and stimulating textbook. Anyone who has taken the trouble to work through it systematically would have a fundamental understanding of the chemical reactions that occur in living systems. The originality in the way the material has been arranged should also help break down artificial barriers between different parts of the subject and demonstrate that all biological processes have a common underlying theme that is derivable from the chemical properties of the various interacting molecular species that make up living matter. There is thus no doubt that Metzler's book is a most significant contribution to the educational biochemical literature.

D.B.R.

Index

Acetabularia, primary pre-rRNA, 18 Acheta domestic us

extrachromosomal rRNA genes, 5 rRNA genes, 19

Acid phenylphosphatase, in thyroid subcellular fractions, 242

Acid phosphatase, in thyroid subcellular fractions, 224

Actinomycin D, effect on synthesis of ribosomal proteins, 52

Adenylate cyclase in thyroid plasma membranes, 219, 246 TSH-sensitive, 249

A granules, in thyroid epithelial cells, 150 Albumin

biosynthesis, and catalase biosynthesis. 202

in hepatoma, 127 secretion, 119

Alcohol, effect on polysome binding, 126 Alkaline phosphatase, in thyroid subcellular

fractions, 222 a-Amanitin, effect on mRNA synthesis, 50 Aminoacyl tRNA, binding to A site of

peptidyl transferase, 16 Amphibia, rRNA gene amplification, 5 Antibody absorption, in spheroplasts, 312 Antigens

as markers, 216 in Micrococcus lysodeikticus membranes,

353-365 Antiserum, to NADH dehydrogenase, 278 Anucleolate mutants, in Xenopus laevis, 9 Ascites cells, pre-rRNA secondary structure,

33 A site, rRNA-tRNA binding, 16 A-T base pairs, in 5 S rRNA, 14

ATPase antigens, 362 latency, role of f subunit, 342 in Micrococcus lysodeikticus membranes,

336-352 fine structure, 337

in mitochondrial stalked bodies, 334 in Streptococcus faecaUs membranes, 336 in thyroid subcellular fractions, 222

Azidodibenzofuran, effect on NADH dehydrogenase, 292

Bacillus megaterium, effect of lysozyme, 313 Bacillus stearothermophilus, stalked bodies,

337 Bacillus subtiUs. "peri plasmic" protein and

fatty acid exchange, 331 Bacterial membranes

freeze fracture studies, 318 plasma membrane function, 310 ultrastructure, 312-319

Benzidine, peroxidase substrate, 233 B granules, in thyroid epithelial cells,

150 Bilayer model, asymmetry, 356 Biogenesis

of eukaryotic ribosomes major stages, 45 posttranscriptional control, 51-58 regulation, 44-58 review literature, 1 transcriptional control, 57-58

of glyoxysomes, 171-203 of peroxisomes, 171-203

Blobel and Sabatini, 144 Branched-chain fatty acids, in Micrococcus

lysodeikticus membranes, 328

395

396

Capsular polysaccharides, immunology, 353 Carcinogenesis, changes in endoplasmic

reticulum, 124 Carcinogens, binding to

cytosol, 124 microsomes, 124 mitochondria, 124 nuclei, 124

Cardiolipin biosynthesis, 330

inhibition by Micrococcus Iysodeikticus mesosomes, 330

in bovine heart mitochondria, 243 mitochondrial marker, 227 role in cytochrome oxidase activity, 295 role in NAOH dehydrogenase activity, 295 synthetase, release from Micrococcus

leisodeikticus. 318 Cartenoids, in Micrococcus Iysodeikticus

plasma membrane, 320 Castor bean endosperm, glyoxysomal mRNA,

187 Castor oil plant, triglycerides, 184 Catalase

biosynthesis, 186 absence of molecular weight changes, 202 and albumin synthesis, 202

in Micrococcus Iysodeikticus cytosol, 365 in peroxisomes, 174 in thyroid subcellular fractions, 225

Cell cycle immunoglobulin biosynthesis, 154 number of rRNA genes, 46

Cell surface glycoproteins, 367 Salmonella typhimurium. 358

Cell wall polymers, biosynthesis, 281 effect of penicillin, 281 in Micrococcus Iysodeikticus. 281-285

Chaotropic agents, effect on hydrophobic interactions, 281 multisubunit enzymes, 280, 281 NAOH dehydrogenase, 272

Chloroplastal ONA, coding for large subunit of ribulose diphosphate carboxylase, 196

Chloroplasts, biogenesis, 196 Cholesterol

in endoplasmic reticulum, 225 in thyroid plasma membranes, 219

Chromatophores, Rhodopseudomonas sphaeroides. 312

Index

Chromophores, in NAOH dehydrogenase, 290

Clofibrate, and peroxisome proliferation, 178 Cloning, 5 S rONA, 31 Clustering, rRNA genes, 3, 4 Colicin E3, effect on protein synthesis, 95 Collagen

effect of prohydroxylase, 146 synthesis by bound polysomes, 127

Colymbetesfuscus. extrachromosomal rRNA genes, 5

Compartmentalization, endoplasmic reticulum, 149-151

Complex I, 268 discontinuous gel electrophoresis, 275 effect of cholate, 296 effect of phospholipase A, 296 epr spectrum, 269

possible effects of denaturation, 271 lipid replacement studies, 294 SOS-polyacrylamide gel electrophoresis,

272 steady state kinetics, 270

Concanavalin A, and Micrococcus Iysodeikticus antigens, 359

Condensing vacuoles, 118 Conformation, rRNA, 13,82 Coomassie Blue stain, NAOH dehydrogenase

subunits, 276 Cooperative effects, in ribosome

reconstitution, 98 Core glycosylation, 193 Cotranslational processes, 193 Crossed immunoelectrophoresis, 353

of NAOH dehydrogenase from M. Iysodeikticus. 365

and SOS gel electrophoresis, 354 Cycloheximide

effect on protein synthesis, 50 and nucleolar RNA polymerase turnover,

49 Cytochrome a" in Micrococcus

Iysodeikticus. 331 Cytochrome b. in Micrococcus Iysodeikticus.

331 Cytochrome b l , in Micrococcus Iysodeikticus.

331 Cytochrome b" biosynthesis, 193

Index

Cytochrome c, in Micrococcus lysodeikticus, 331

Cytochrome oxidase role of cardiolipin, 295 in thyroid subcellular fractions, 219

latency, 217 Cytochrome P-450, in endoplasmic reticulum,

122 Cytosol

carcinogen binding, 124 ribosome migration, 57 thyroid lactate dehydrogenase, 221

D-Alanine carboxypeptidase, and UDP-muramylpentapeptide, 324

"Dangling" messenger, 141 Danielli and Davson model, 358 Do-Carboxypeptidase, 320

in Micrococcus lysodeikticus, 320 Dehydrogenases, in M. lysodeikticus plasma

membranes, 320 Deoxyribonuclease, from thyroid, 217 Diazobenzene sulfonate, labeling of NADH

dehydrogenase, 324 Differential pelleting, thyroid homogenates,

218-222 Differential replication, rRNA genes, 16 Dimyristoyl lecithin, and complex I, 294 Di-Na-phenylphosphatase, from thyroid,

217 Diphenyl iodonium, and NADH

dehydrogenase, 290 Diphtheria toxin, membrane transport, 197 Discontinuous gel electrophoresis, 275 Dithionite, reduction of NADH

dehydrogenase chromophores, 281 "Domains," in ribosomal subunits, 86

RNA-protein interactions, 87 Drosophila, plasmid rRNA genes, 30 Drosophila melanogaster

rRNA genes, 8 5 S rRNA, 14

Dytiscus marginalis, rRNA genes, 19 extrachromosomal, 5

EDTA, and membrane-ribosome interactions, 13 7

Electron shuttles, role of peroxisomes, 177 Electron transport chain, in Micrococcus

lysodeikticus. 331-334

Endonucleases action on pre-rRNA, 35 in nucleoli, 42

Endoplasmic reticulum changes during carcinogenesis, 124 cholesterol content, 225 cisternae, 117 compartmentalization, 149-151

397

continuity of rough with smooth, 123 continuity with peroxisomal membrane, 189 cytochrome P-450, 122 functional aspects, 121-125 glucose-6-phosphatase, 228 integral proteins, 129 lipoprotein globules, 120 luminal face, 129 membrane composition and structure, 119-

121 membrane symmetry, 120 "microenvironments," 148 monooxygenase, effect of drugs, 123 mRNA binding, 148 NADH-cytochrome P-450 reductase, 122 peripheral proteins, 129 phospholipid synthesis, 188 posttranslational modifications, 149-151 protein synthesis, 125-137

compartmentalization, 149-151 ratio of smooth to rough, 122 role in protein synthesis, 117-155

Enzymes, as markers, 216 epr spectroscopy, NADH dehydrogenase, 269 Escherichia coli

cardiolipin biosynthesis, 330 Fo-F,-ATPase complex, 351 membrane vesicles, 362 ribosomes, 81-104 23 S rRNA, nucleotide sequence, 83 transport, 362

Ethionine, effect on polysome spirals, 141 Etiolated leaves, peroxisomes, 186 Eukaryotes

gene expression, 59 RNA polymerases, 16

Eukaryotic ribosomes biogenesis, 213-251

posttranscriptional control, 51-58 regulation, 44-58 review literature, 1 transcriptional control, 46-51

398

Eukaryotic ribosomes (cont.) preribosomes, processing, 31-44 pre-rRNA

processing, 31-44 structure, 32-33

ribosomal genes, 2-16 differential replication, 16 transcription, 16-31

in vitro, 29-31 5 S rRNA genes, 12-14

Evolution of L-rRNA sequences, 32 of rRNA terminal nucleotides, 40 of S-rRNA sequences, 32

Exonucleases, action on pre-rRNA, 35 Extrachromosomal rRNA genes, 5

F.-ATPase, in Escherichia coli, 351 F.-F1-ATPase complex, in Escherichia coli,

351 F1-ATPase

biosynthesis, 280 in Escherichia coli

mutants, 351 reconstitution, 350

~ subunit, 342 latency, 349 in Micrococcus Iysodeikticus, 346

cytoplasmic face of plasma membrane, 340

membrane location, 336-352 molecular weight, 347 possible glycosylation, 348 proteolytic degradation, 342 subunit composition, 347

10-nm particles, 319 reconstitution in E. coli, 350 in Salmonella typhimurium, subunit

composition, 349 Fatty seeds, glyoxysomes, 184-185

proliferation, 187 Ferritin-antibody labeling, lipotechoic acids,

358 Flavoprotein core, NADH dehydrogenase,

302 Flavoprotein fragments

in complex I, 273 in NADH dehydrogenase, 302

"Flip-flop," monolayer phospholipids, 189 5-Fiuoroorotate, and mRNA biosynthesis, 53

Index

FMN, in NADH dehydrogenase, 269 Free ribosomes, effect of partial hepatectomy,

126 Freeze-fracture

bacterial membranes, 318 mesosomes, 319

Galactosyltransferase, in thyroid subcellular fractions, 221, 243

GDP-mannose, and lipomannan biosynthesis, 327

Gel filtration, thyroid homogenates, 218 Gene expression, eukaryotic ribosome

biogenesis model, 59 Gene-spreading technique, 26 Gluconeogenesis, and glucose-6-phosphatase,

228 Glucose-6-phosphatase, in thyroid subcellular

fractions, 228 correction by phenyl phosphatase, 229 nuclear activity, 229

~-Glucuronidase, in thyroid subcellular fractions, 218

latency, 217 Glutamate dehydrogenase, synthesis by

membrane-bound polysomes, 135 ~-Glycerophosphatase, in thyroid subcellular

fractions, 217 Glycoprotein G, in vesicular stomatitis virus

(VSV),I92 Glycoproteins

in lysosomes, 149 in mitochondria, 149 oligosaccharide patterns, 150 synthesis in rough endoplasmic reticulum,

193 Glycosylation

at polysome level, 152 of secretory proteins, 128 of vesicular stomatitis virus protein, 192

Glycosyltransferases in Golgi apparatus, 199 membrane-bound, 150

Glyoxylate cycle, in glyoxysomes, 173 Glyoxysomes

biochemical constitution, 173 biogenesis, 171-203 from fatty seeds

biochemistry, 184-185 proliferation, 187

Index

Glyoxysomes (cont.) glyoxylate cycle, 173, 185 isocitrate lyase, 185 malate dehydrogenase biosynthesis, 203 malate synthase, 185 membranes

alkaline lipase, 198 cytochrome bs, 198 N-demethylases, 198 hydroxylases, 198 polypeptide composition, 199 similarity to endoplasmic reticulum, 199

morphology, 172-174 mRNAs, 187 .a-oxidation pathway, 173 proximity to spherosomes, 184 topographic relationships, 172-174

Golgi apparatus, 118 glycosyl transferases, 150 and protein glycosylation, 199 proteins, site of synthesis, 136 from thyroid, 243-244

separation from plasma membrane, 243 Gradient centrifugation, thyroid

homogenates, 221-227 Gram-negative organisms

freeze-fracture studies, 318 outer membrane, 311 plasma membrane, 311

Gram-positive organisms, freeze-fracture studies, 318

Guaiacol, peroxidase substrate, 233 Guaiacol peroxidase

effect of heparin on sedimentation, 233 in thyroid rough endoplasmic reticulum,

233

~+-ATPase, in Micrococcus Iysodeikticus, 352

Heparin and peroxidase sedimentation, 233 and thyroid rough endoplasmic reticulum,

225 Hepatoma, albumin synthesis, 127 Heteroduplex mapping, 10 Heterogeneity, rough endoplasmic reticulum,

151 Histone mRNA, 143 Histones, and transcribed rDNA, 23 HnRNA, biosynthesis, 52

Homogenization, thyroid tissue, 214 get filtration of homogenates, 218

399

Human chromosomes, rRNA gene clusters, 4 Hydrophilic probes, NADH dehydrogenase

labeling, 282-284 Hydrophobic interactions, in NADH

dehydrogenase, 281,285,286 Hypertrophic thyroid, nuclear lipids, 240

Immunoelectron microscopy, ribosomal proteins, 10, 101

Immunoelectrophoresis crossed two-dimensional, 353 of Micrococcus Iysodeikticus membranes,

345 of succinylated lipomannan, 346

Immunoglobin secretion, 119 Immunoglobulin, light chain biosynthesis,

152 Initiation complex, 96 Initiation factor IF-3

effect on ribosomal proteins, 99 interaction with 16 S rRNA, 98

Initiation of protein synthesis, effect of Verrucarin A, 147

Insects, rRNA gene amplification, 5 Insulin secretion, 119 Intermediate pre-rRNA, 37 Introns, in rRNA genes, 8 Iodonaphylazide

labeling of NADH dehydrogenase, 285 [ 12SIl peroxidase, labeling of membranes, 366 Iron-protein fragment

in complex I, 273 in NADH dehydrogenase, transmembrane

organization, 299 Iron-sulfur groups, in NADH

dehydrogenase, 271 Isocitrate dehydrogenase, in Micrococcus

Iysodeikticus cytosol, 365 Isoelectric points, NADH dehydrogenase

subunit polypeptides, 281, 282

Kethoxal, action on 16 S rRNA, 16 Kidney glycoproteins, synthesis by bound

polysomes, 127

Lactate dehydrogenase, in thyroid cytosol, 221

Lactobacilli, lysoteichoic acids, 358

400

Lactoperoxidase iodination, NADH dehydrogenase, 283

Laser light scattering, rRNA studies, 85 Latency, F1-ATPase, 342 Leaf peroxisomes, 181-183

proliferation, 186-187 Light chain immunoglobulin

biosynthesis in plasmacytoma, 152 in cell cycle, 154

Lipases, in spherosomes, 187 Lipid biosynthesis, in Micrococcus

lysodeikticus membranes, 328-331 Lipid depletion, NADH dehydrogenase

studies, 294 Lipid-protein interactions, in NADH

dehydrogenase, 292-298 Lipids

in hypertrophic thyroid nuclei, 240 in succinate dehydrogenase, 332 in thyroid mitochondria, 241 in thyroid nuclei, 240

Lipid transport in membranes, 191 Lipolytic enzymes, in thyroid subcellular

fractions, 234-238 Lipomannan biosynthesis

in Micrococcus lysodeikticus, 325-327 mesosomes,322

role of GDP-mannose, 327 Lipopolysaccharides

immunology, 353 localization in Salmonella typhimurium,

358 Lipoprotein globules, in endoplasmic

reticulum, 120 Lipoteichoic acids, ferritin-antibody labeling,

358 Liver peroxisomes

biochemical properties, 181-183 proliferation, 185-186

Long-Acting Thyroid Stimulator (LATS) Inhibitor, 247

L-rRNA evolution, 32 genes, 2 intermediate pre-rRNA, 37 in nucleolus, 58 segment in primary pre-rRNA, 29

Lymphocytes, effect of phytohaemagglutinins, 55

Lysophospholipase, in thyroid, 231

Lysosomes gl ycoproteins, 149 thyroglobulin hydrolysis, 214 from thyroid, 241, 242

heterogeneity, 224 hydrolases, 217 mitochondrial contamination, 219

Lysostaphin, effect on Staphylococcus aureus, 313

Lysozyme

Index

effect on Bacillus megaterium strain KM, 313 effect on Micrococcus lysodeikticus, 309

Malate dehydrogenase, biosynthesis glyoxysomal enzyme, 203 role of membrane-bound polysomes, 135

Mammary gland milk proteins, synthesis by membrane-bound polysomes, 127

Mannan biosynthesis, 326 in Micrococcus lysodeikticus, 325

Mannobiosyldiglyceride, in Micrococcus lysodeikticus, 327

Mannosaminuronic acid polysaccharide, 322 Mannosyl-I-phosphoryl undecaprenol, role in

mannan biosynthesis, 326 Mannosyl transferase, in thyroid subcellular

fractions, 221 Marker enzymes, in thyroid, 216-218 Matrix proteins, in microbodies, 197

site of biosynthesis, 194 Matrix units, in rDNA, 21 Maturation

of pre-rRNA, 34-42 of primary preribosomes, 59

Meiotic recombination, and nontranscribed sequences, 15

Membrane-bound ribosomes in brain cortex cells, 136 in choroidal epithelial cells, 136 cytochrome b, biosynthesis, 193 in differentiating muscle, 136 effect of alcohol administration, 126 nascent collagen polypeptides, 146 synthesis of

catalase, 186 cytosolic enzymes, 128 glutamate dehydrogenase, 135 malate dehydrogenase, 135 mitochondrial proteins, 135

Index

Membrane flow hypothesis, 190 and microbody membrane biogenesis, 201

Membrane lipids, in microbodies, 188-191 Membrane potential, in Micrococcus

lysodeikticus membranes, 334 Membranes

asymmetry, 190,356 bacterial,312-320 from endoplasmic reticulum, 118-125

composition and structure, 119-121 from erythrocytes, 132 from Micrococcus lysodeikticus, 320-352 interaction with

mRNA, 147-148 nascent polypeptides, 143-147 polysomes, 137-148 60 S ribosome subunit, 137-143

labeling by [12SIjperoxidase, 366 lipid transport, 191 peroxisoinal, lipid composition, 198 proteases, 13 2 proteins

in microbodies, 191-193 in reticulocytes, 133 transfer and "signal" hypothesis, 144

symmetry, 120 thyroid,238-251 vesicular stomatitis virus (VSV),

biogenesis, 191 Membrane transport, diphtheria toxin, 197 Membrane vesicles, 214, 362 Menaquinones

in Micrococcus lysodeikticus membranes, 320

in Mycobacterium ph lei, 333 Mesosomes

artifactual nature, 319 fatty-acid exchange with plasma

membrane, 331 freeze-fracture studies, 319 membranes, 313

Methylation, of pre-rRNA, 32 Microbodies

dimensions, 172 matrix, 172 matrix proteins

biosynthesis, 201-203 segregation, 197 site of synthesis, 194

Microbodies (cont.) membranes, 172

membrane flow, 201 models for biogenesis, 193-194 synthesis of

lipids, 188-191 proteins, 198-201

401

models of biosynthetic mechanisms, 187-197

proliferation, 185-187 proximity to endoplasmic reticulum, 174-

185 review literature, 171

Micrococcus flavius, lipomannan, 326 Micrococcus lysodeikticus

absence of lipoteichoic acids, 326 ATPase antigens, 362 ATPase staining, 337 cytoplasmic compartment

catalase, 365 isocitrate dehydrogenase, 365 polynucleotide phosphorylase, 365

Do-carboxypeptidase, 320 F,-ATPase

CaH stimulation, 346 localization in plasma membrane, 340 molecular weight estimates, 347 possible glycosylation, 348 proteolytic degradation, 342 subunit composition, 347

H+-ATPase, 352 history of research, 309 lipomannan, 326 mannan, 325 mannobiosyl diglyceride, 327 mannosaminuronic acid polysaccharide, 322 membranes

absence of phosphatidyl ethanolamine, 328

alcohol dehydrogenase, 332 antigenic architecture, 353-365 biochemical characterization, 320-352 cardiolipin localization, 363 coupling factor, 349-352 cytochromes a2, b, b" and c, 331 D-Iactate dehydrogenase, 332 EDT A shock washes, 318 electron transport chain, 331-334 enzymic distribution, 320-322 F,-ATPase, 336-358

402

Micrococcus lysodeikticus (cont.) fatty acid composition, 328 immunoelectrophoresis, 345 lipid biosynthesis, 328-331 lipomannan biosynthesis, 325-327 malate dehydrogenase, 332 NADH dehydrogenase, 332 10-nm particles, 319 phosphatidyl ethanolamine localization,

363 phosphatidyl glycerol localization, 363 release of cardiolipin synthetase, 318 release of NADH dehydrogenase, 318,

343 solubilization by Triton X-100, 355 succinate dehydrogenase, 332

mesosomes absence of respiratory enzymes, 356 distinction from plasma membranes,

313 inhibition of cardiolipin synthesis, 330 role in lipomannan biosynthesis, 322 succinylated lipomannan, 356

mucopeptide, 310 murein, 310 NADH dehydrogenase

crossed immunoelectrophoresis, 365 release from membranes, 343

nomenclature, 309 peptidoglycan, 310

metabolism, 323-325 plasma membrane

antigenic complexity, 353, 359 asymmetry of antigens, 359 carotenoids, 320 cytochromes, 320 D-alanine carboxypeptidase, 324 dehydrogenases, 320 distinction from mesosomal membranes,

313 general properties, 366 menaquinones, 320 phospholipids, 320

polymannose, 325 protoplasts, surface antigens, 359 respiratory chain

inhi bi tors, 334 membrane potential generation, 334

subfractionation, 320 release of Do-carboxypeptidase, 320

Micrococcus lysodeikticus (cont.) surface antigens, reactions with

concanavilin A, 359 surface profiles, 312 teichuronic acid, biosynthesis, 324 wall polymer biosynthesis, 323-325

Index

Micrococcus sodonensis. lipomannan, 326 Micronucleus, Tetrahymena pyriformis

rRNA genes, 6 Microsomal fraction

carcinogen binding, 124 thyroid, 229

Mitochondria carcinogen binding, 124 F,-A TPase biosynthesis, 280 glycoproteins, 149 inner membrane

ATPase, 336 NADH dehydrogenase organization, 268 stalked bodies, 336

protein synthesis, membrane-bound polysomes, 135

succinate dehydrogenase, 332 from thyroid, 241,242

cardiolipin content, 227 lipid composition, 241 lysosomal contamination, 219

from yeast, attached 80 S ribosomes, 195

Mitotic recombination, 15 Monolayers

phospholipid "flip-flop," 189 in thyroid nuclei, 241

Monooxygenase, in endoplasmic reticulum, 123

mRNA binding to endoplasmic reticulum, 148 for cytochrome b" 193 for glyoxysomal proteins, 187 for histone, 143 interaction with membranes, 147-148 recognition by ribosomes, 95 in rough endoplasmic reticulum, 151 segregation in rough endoplasmic

reticulum, 149 synthesis, effect of a-amanitin, 50 for thyroglobulin, 245 transfer from nucleus to cytoplasm, 148

Multisubunit enzymes, 281, 282 Muramidase, effect on streptococci, 313

Index

Mycobacterium phlei menaquinone, 333 NADH oxidation, 333

Myeloma cells histone mRNA, 143 60 S ribosomal subunits, 146

N-Acetylglucosaminyltransferases, in thyroid subcellular fractions, 243

N-Acetylhexosaminidase, in thyroid subcellular fractions, 242

NADH binding site in NADH dehydrogenase, 289 oxidation via peroxisomes, 177

NADH-cytochrome P-450 reductase, in endoplasmic reticulum, 122

NADH dehydrogenase alternative names, 268 binding site for NADH, 289 chromophores, 281, 290 discontinuous gel electrophoresis, 275 effect of chaotropic agents, 271 effect of cholate, 294 electron-carrying arm, 298 flavoprotein core, 302 flavoprotein fragment

NADH-ferricyanide oxidoreductase, 287 polypeptide composition, 273 surface covering, 302

FMN content, 269 fragmentation of enzyme, 271-272 functional unit, 269-271 gel electrophoresis, 272 hydrogen-carrying arm, 298 hydrophobic interactions, 281 iron-protein fragment, 273

redox potential, 290 transmembrane organization, 299

iron-sulfur groups, 271 isolation of pure enzyme, review literature,

267 labeling by diazobenzene sulfonate, 282 labeling by iodonaphthylazide, 285

effect of lipid depletion, 297 lactoperoxidase iodination, 283 lateral organization, 300-301 lipid depletion studies, 294

effect on iodonaphthylazide labeling, 297

lipid-protein interactions, 292-298

NADH dehydrogenase (cont.) membrane-bound enzyme, review

literature, 267

403

from Micrococcus lysodeikticus. crossed immunoelectrophoresis, 365

model of structure, 301-303 molecular organization, 267-303 NADH binding site, 289 paramagnetic species, 269 phospholipids, 292-298 polypeptide composition, 272-279 polypeptide isoelectric points, 281-282 protein components, 271-279 protein structure, 279-292 proteolytic digestion, 286-289 organization in membrane, 298-301 reaction with azidodibenzofuran, 292 reaction with diphenyliodonium, 290 reaction with ubiquinol-cytochrome c

oxidoreductase, 300 reduction by dithionite, 281 release from Micrococcus lysodeikticus

membranes, 318 review literature, 267 role of cardiolipin, 295 steady state kinetics, 270 structure/function relationships, 289-292 subunit components

Coomassie Blue staining, 276 coordinated antiserum precipitation, 278 labeling by diazobenzene sulfonate, 282 labeling by iodonaphthylazide, 285 polypeptides, 272-279 proteolytic digestion, 286

terminology, 268-269 transmembrane organization, 298-300 treatment with chaotropic agents, 272 ubiquinone reductase, effect of

phospholipid removal, 292 NADH-ferricyanide oxidoreductase, 287 NADH-ubiquinone oxidoreductase, 268 NADPH-cytochrome c reductase

in plasma membrane cytoplasmic face, 133 in thyroid nuclear fractions, 229

Nascent polypeptides, interaction with membranes, 143-147

Nectin, and ATPase attachment to membranes, 350

Neutron scattering, and rRNA packing, 99 Nitrogen cavitation, 248

404

10-nm particles, in Micrococcus Iysodeikticus membranes, 319

N', N'-dimethyladenosine, in 16 S rRNA, 103

Nonhistone proteins, and transcribed rDNA, 23

Nonribosomal proteins, and preribosomes, 44 Nontranscribed sequences

and recombination, 15 in rDNA, 21 in rRNA genes, 10

Nuclei carcinogen binding, 124 from Chinese hamster kidney cells, RNA

polymerase, 236 5 S rRNA synthesis, 31 from hypertrophic thyroid

chemical composition, 239 isopycnic density, 238

from thyroid, 238-241 chemical composition, 239 exogenous DNA and RNA polymerase,

236 glucose-6-phosphatase, 229 isolation techniques, 216 isopycnic density, 238 lipid composition, 240 NADPH-cytochrome c reductase, 229 peroxidase, 229

Nucleolus from L cells, endonucleases, 42 L-rRNA particles, 58 maturation of primary preribosome, 59 from Novikoff hepatoma, endonucleases, 42 and ribosome biogenesis, 60 ribosome migration, 57 RNA polymerase turnover, 49 45 S transcription products, 30

Nucleoplasm, ribosome migration, 57 Nucleosomes, unfolding, 24 5'-Nucleotidase, in thyroid subcellular

fractions, 222 Nucleotide precursors, in peptidoglycan

biosynthesis, 323 Nucleotides, effect on rRNA gene

transcription, 51 Nucleotide sequence, E. coli 23 S rRNA, 83 Nucleus

degradation of ribosomes, 55 rRNA wastage, 55

Index

Obligate aerobes, F1-ATPase latency, 349 Oligonucleotides, release from rRNA, 101 Organelles, thyroid, 238-251 tJ-Oxidation pathway, in glyoxysomes, 173

Palmitoyl coenzyme A, oxidation by peroxisomes, 179

Partial hepatectomy effect on free ribosomes, 126 effect on peroxisomes, 185

Penicillin, and bacterial cell-wall biosynthesis, 323

Peptidoglycans biosynthesis, 323 in Micrococcus Iysodeikticus, 323-325

Peptidyl transferase center, aminoacyl tRNA binding, 16

"Peri plasmic" protein, 331 Peroxidase

substrates, 233 in thyroid subcellular fractions, 232-234

nuclear activity, 229 Peroxisomes

biochemical properties, 174-185 biogenesis, 171-203 catalase

contribution to total peroxisomal protein, 174

site of synthesis, 186 effect of clofibrate, 178 effect of partial hepatectomy, 185 effect of tibric acid, 180 effect of Wy-14643, 180 fatty acid oxidation pathway, 118 from leaves

biochemical properties, 181-183 malate dehydrogenase, 182 ontogeny, 186 photorespiratory glycolate pathway, 182 proliferation, 186-187 urate oxidase, 182

from liver biochemical properties, 181-183 proliferation, 185-186

matrix proteins biosynthesis, 201-203 segregation, 197

membrane biogenesis models, 193-194 membrane lipid biosynthesis, 188-191,

197,198

Index

Peroxisomes (cont.) membranes, continuity with endoplasmic

reticulum, 189 lipid composition, 198 protein biosynthesis, 188-191, 198-201

morphology, 172-174 palmitoyl-CoA oxidation, 179 proliferation, 185-187 substrate-mediated electron shuttles, 177 topographical relationships, 172-174

Phosphatidylcholine in bovine heart mitochondria, 293 in peroxisomal membranes, 198

Phosphatidylethanolamine absence from Micrococcus lysodeikticus

membranes, 328 in bovine heart mitochondria, 293 in peroxisomal membranes, 198

Phosphatidylinositol, in peroxisomal membranes, 198

Phosphodiesterase, in thyroid, 217 Phospholipase, in thyroid, 231, 232 Phospholipid exchange proteins, 198 Phospholipids

biosynthesis, 188 "flip-flop," 189 in Micrococcus lysodeikticus membranes,

320 in NADH dehydrogenase, 267

role in ubiquinone reductase, 292 Photorespiration

effect of oxygen, 181 and photosynthesis, 181

Physarum polycephalum rONA matrix units, 21 r-chromatin, 18 5 S rRNA genes, 12

Plant leetins, membrane transport, 197 Plasmacytoma, immunoglobulin biosynthesis,

152 Plasma membrane

asymmetry, 120 bacterial, 310 cytoplasmic face, 133 in gram-negative bacteria, 311 in Micrococcus lysodeikticus

antigenic complexity, 353, 355 general properties, 366

NADPH-cytochrome c reductase, cytoplasmic face localization, 133

Plasma membrane (cont.) prokaryotic, 310

compared to eukaryotic, 367 proteins, site of biosynthesis, 136 reaction with agglutinins, 363 in thyroid, 245-251

adenylate cyclase, 246 contamination of isolated fractions,

246

405

heterogeneity of isolated fractions, 250 lipid content, 247 method of isolation, 245

Plasmids, with Drosophila rRNA genes, 30 Pleurodeles waltlii. rRNA genes, 19 Polyaldehydes, cross-linking of ribosomal

proteins, 11 Polymannose, in Micrococcus lysodeikticus.

325 Polynucleotide phosphorylase, in

Micrococcus lysodeikticus cytosol, 365

Polysome run off, 147 Polysomes

attachment via dangling messenger, 141 effect of alcohol administration, 126 effect of ethionine, 141 free, protein synthesis activity, 125-127 in Golgi fractions, 136 interactions with membranes, 137-148 membrane-bound

protein synthesis activity, 320-323 synthesis of secreted proteins, 323-

325 protein glycosylation, 152 from thyroid, 244, 245

Polytron homogenizer, 248 Posttranscriptional control, of ribosome

biogenesis, 57-58 Posttranslational modification, of proteins in

endoplasmic reticulum, 155 pppNp, in primary pre-rRNA, 28 pre-mRNA

effect of 5-fluoroorotate, 53 effeet of toyocamycin, 53

Preribosome intranuclear degradation, 55 nonribosomal proteins, 44 processing, 31-44 and structural ribosomal proteins, 52 structure, 293-296

406

Pre-rRNA action of nucleases, 35 denaturation spectra, 33 gel electrophoresis, 34 maturation pathways, 34-42 methylation, 342 processing, 31-44

intermediates, 37 and rRNA pools, 39 secondary structure, 32, 33 uridine to pseudouridine conversion, 32

Primary preribosome, maturation, 59 Primary pre-rRNA

heterogeneity, 27 pool composition, 27 pppNp content, 28 size variations, 18 termination signal, 29 use of gene-spreading technique, 26

Primary rONA transcript, 25 Proalbumin, 144 Proinsulin, 144 Prokaryote plasma membrane, 310 Prolyl hydroxylase, action on collagen, 146 Promoter signals, in rRNA genes, 13 Proteases, in membranes, 132 Protein binding sites, in E. coli rRNA, 94-

99 Protein glycosylation, and Golgi apparatus,

199 Protein-mediated lipid transport, 191 Protein synthesis

by free polysomes, 125-127 inihibition by colicin E3, 95 initiation, inhibition by Verrucarin A, 147 by membrane-bound polysomes, 125-127 posttranslational modifications, 149-151 role of endoplasmic reticulum, 117-155 by rough endoplasmic reticulum, 125-137,

148-154 in thyroid, 244, 245

Proteolysis of NADH dehydrogenase, 286 Proton translocation, in Micrococcus

lysodeikticus F1-A TPase, 352 Protoplasts, labeling by [12SIJperoxidase, 366 Pyrophosphate, effect on guaiacol peroxidase

sedimentation, 233

Qp RNA, initiation complex with 70 S ribosomes, 96

Radius of gyration of rRNA, 86 Rat liver

Index

peroxisomal fatty acyl-CoA oxidation, 181 phosphatidyl exchange proteins, 198 ribosomal turnover, 54

r-Chromatin and active transcription units, 20 from extrachromosomal rRNA genes, 18

rONA his tones, 23 nonhistone proteins, 23 non transcribed segments, 21 nucleosome chain compaction, 23 oocyte, 13 Physarum polycephalum, 21 primary transcript, 25 somatic, 13 Tetrahymena pyriformis, 11 transcription units, 25

heterogeneity, 27 Recombination, and nontranscribed

sequences, 15 Regenerating liver, peroxisomes, 185 Respiratory chain, in Micrococcus

lysodeikticus, 334 membrane potential gradient, 334

Reticulocytes, membrane protein synthesis, 133

Rhodopseudomonas sphaeroides chromatophores, 361 spheroplast antibody absorption, 312

Ribonuclease effect on rRNA, 101 from thyroid, 217

Ribonucleoprotein domains cross-linking studies, 11 in ribosomal subunits, 86-90 S4, S8, SIS, and S7 proteins, 12

Ribophorins lactoperoxidase iodination, 139 and 60 S ribosomal subunit, 139

Ribosomal genes, for eukaryotic ribosomes, 2-16

Ribosomal L-proteins, recycling, 44 Ribosomal protein L24, binding site on

rRNA,84 Ribosomal protein S4, binding site on rRNA,

84 Ribosomal proteins

asymmetry, 10

Index

Ribosomal proteins (cant) cross-linking by polyaldehydes, 11 effect of initiation factor IF-3, 99 effect on RNA-RNA interactions, 14 effect on rRNA conformation, high

temperature interactions, 15 localization in ribosomal subparticles, 81 macromolecular structure, 81 and RNA tertiary structure, 91-94 role in r RN A tertiary structure, 14 spatial organization in ribosomes, 104 in "80 S" preribosomes, 43 topography, 10

Ribosomal RNA, see rRNA Ribosomal S proteins, recycling, 44 Ribosomal subunits

reconstitution, 13 ribonucleoprotein domains, 86-90 50 S particle, 13

Ribosomes binding to membranes

effect of EDTA, 137 role of integral membrane proteins, 139

biogenesis major stages, 45 model of gene expression, 59 and nucleolar structure and function,

60 review literature, I

detachment from rough endoplasmic reticulum, 125

effect of colicin E3, 95 from E. coli, 81-104

ribosomal RNA secondary structure, 83-85

function review literature, 86-90 role of ribosomal RNA, 94-99

initiation complex with Q~ RNA, 96 intranuclear degradation. 55 from liver, effect of partial hepatectomy,

126 maturation, 42 migration, 57 mobility in rough endoplasmic reticulum,

140 protein pool, 52 proteins, effect of actinomycin D on

biosynthesis, 52 recognition of mRNA, 95

407

Ribosomes (cont.) reconstitution

cooperative effects, 98 reconstitution in vitro, 98

ribonucleoprotein segments ("domains"), 86

ribosomal protein organization, 81 spatial organization, immunoelectron

microscopy, 101 23 S rRNA, nucleotide sequence, 91-94 structural proteins, 52 structure, review literature, 82 subunits, "domains," 86 topography of rRNA, 94-99 turnover, 54 X-ray scattering studies, 100 on yeast mitochondrial surface, 195

Ribulose biphosphate carboxylase, transport of small subunit, 196

Ricin agglutinin, 363 Ricinus communis, triglyceride store, 184 RNA polymerase, adenylation, 50

effect of exotoxins, 50 in eukaryotes, 16 multiple forms, 234 in nuclei

from Chinese hamster kidney cells, 236 nucleolar turnover, 49

phosphorylation, 50 in thyroid

bimodal localization, 238 subcellular distribution, 234-238

RNA polymerase A, 16 diploid cell content, 48 effect on rRNA gene transcription, 51 metabolic stability, 49 and rRNA gene transcription, 48

RNA polymerase B, 16 RNA polymerase C, 16 RNA-protein interactions, 87 RNA-RNA interactions, 87

in ribosomal domains, 87 in terminal fraction of 23 S rRNA, 88

RNP fragments, in 30 S ribosomal subunits, 87

RNP particles, shadow casting, 103 Rough endoplasmic reticulum

continuity with smooth, 123 functional heterogeneity, 148-154 guaiacol peroxidase, 233

408

Rough endoplasmic reticulum (cont.) heterogeneity, 151 integral membrane proteins, 139 protein synthesis, 148-154 proximity to microbodies, 173 relative amount of smooth reticulum, 121 ribosome detachment, 125

effect of EDTA, 137 ribosome mobility, 148 segregation of mRNA, 149 synthesis of specific proteins

compartmentalization, 151-154 sites of synthesis, 151-154

thyroid, effect of heparin, 225 vectorial discharge of proteins, 143

rRNA conformation, 13, 15,82 effect of ionic strength, 82 effect of ribonuclease, 101 from E. coli, secondary structure, 83-85 flexibility, 15 in free and bound state, 84 gel electrophoresis, 13 genes

in Acheta domesticus, 19 active, 20 amplification

in amphibia, 5 in insects, 5

clustering, 3 differential replication, 16 in Drosophila melanogaster, 8 in Dytiscus marginalis, 19 in eukaryotes, 2-12 extrachromosomal, 5 factors affecting

elongation, 47 initiation, 47 termination, 47

introns,8 mechanism of activation, 24 in micronucleus of Tetrahymena

pyriformis, 6 nontranscribed segments, 10 number, 46 in plasmids, 30 prokaryotic, 3 promoter signals, 13 rate of transcription, 51 and RNA polymerase A, 48

rRNA (cont.) in Saccharomyces carlsbergensis, 8 in Saccharomyces cerevisiae, 8 switching on and off, 47 terminator signals, 13 transcription, 16-31,45

complex, 16-18 process, 18-29 units, 6

in Triturus, 19 in Xenopus laevis, 9, 19

hypochromic effects, 84 intranuclear wastage, 55 laser light scattering, 85 melting curve, 84 packing in subunits

neutron scattering studies, 99 X-ray studies, 99

radius of gyration, 86 in ribosomal subunits, 85-86 role in ribosome function, 94-99 secondary structure

in free state, 83-85 in ribosomes, 83-85

studies with antibodies, 103 terminal nucleotide evolution, 40 tertiary structure, 91-94

and ribosomal proteins, 14 topography in ribosomes, 94-99 translational diffusion constant, 85 tRNA binding to A site, 16 volume in ribosomal subunits, 86

rRNA pools, and pre-rRNA, 39

Index

Saccharomyces, primary pre-rRNA, 18 Saccharomyces carlsbergensis, rRNA genes,

8 Saccharomyces cerevisiae, rRNA genes, 8 Salmonella typhimurium

F,-ATPase subunit composition, 349 Iipopolysaccharides, 358

SDS-polyacrylamide gel electrophoresis of complex I, 272 and crossed immunoeiectrophoresis, 354

Secretory proteins intracellular storage, 118 role of glycosylation, 128 synthesis on membrane-bound polysomes,

127, 128 vectorial transfer, 117

Index

Serine dehydratase, synthesis on membranebound polysomes, 128

Shadow casting, RNP particles, 103 Sialic acid, in thyroid plasma membranes,

219 Sialyltransferase, in thyroid subcellular

fractions 221, 243 Signal hypothesis, 144 Signal peptides, 144 Smooth endoplasmic reticulum

in adrenal gland cells, 119 continuity with rough endoplasmic

reticulum, 123 proximity to microbodies, 173 ratio to rough endoplasmic reticulum, 121 in testis interstitial cells, 119

Sonication, thyroid microsomal fraction, 229

Soybean agglutinin, 363 Spheroplasts, from Rhodopseudomonas

sphaeroides, 312 Spherosomes

Jipases, 184 proximity to glyoxysomes, 184

"55 S" preribosomes, and "80 S" preribosome, 42

"80 S" preribosome nonribosomal proteins, 44 precursor of "55 S" preribosome, 42 protein complement, 43

21 S pre-rRNA, 38 32 S pre-rRNA, 38 36 S pre-rRNA, 38 5 S rONA, injection into Xenopus laevis

oocytes,31 30 S ribosomal subunit, digestion with

RNase, 87 50 S ribosomal subunit, proteins involved in

packing, 13 60 S ribosomal subunit

association with ribophorins, 139 interaction with membranes, 137-143 in myeloma tissue culture, 146

70 S ribosomes, chloroplastal, 196 ribulose biphosphate carboxylase synthesis,

196 S-rRNA

evolution, 32 genes, 2 intermediate pre-rRNA, 37

5 S rRNA in Drosophila melanogaster, 14 genes

in eukaryotes, 12-14 in lower eukaryotes, 12

in vitro synthesis, 31 5.8 S rRNA

genes, 2 7 S precursor, 37 terminal nucleotides, 41

16 S rRNA binding sites for IF-3, 98 N", N"-dimethyladenosine, 103 protein binding sites, 84 secondary structure, 85 3'-terminal segment and mRNA

recognition, 95 23 S rRNA, E. coli

nucleotide sequence, 83 500-nucleotide terminal fragment, 88 protein binding sites, 84

Stalked bodies in Bacillus stearothermophilus, 337 on mitochondrial inner membrane, 336

Staphylococci, surface profiles, 312 Staphylococcus aureus, cardiolipin

biosynthesis, 330

409

Steady state kinetics, NADH dehydrogenase, 270

Steroid hormone biosynthesis, in smooth endoplasmic reticulum, 119

Streptococci effect of phage muramidase, 313 surface profiles, 312

Streptococcus faecalis, membrane ATPase, 336

role of nectin, 354 Subcellular biochemistry, of thyroid, 213-251 Succinate dehydrogenase, removal of lipids, 332 Succinylated lipomannan

immunoelectrophoresis, 346 in mesosomes, 356

Supranucleosomal globules, 23

Teichuronic acid, biosynthesis in Micrococcus lysodeikticus, 324

Terminal fragment, 23 S rRNA, 88 Terminal nucleotides, in rRNA

evolution, 40 of 5.8 S rRNA, 41

410

Termination signals in primary pre-rRNA, 29 in rRNA genes, 13

Tetrahymena pyriformis r-chromatin, 18 rDNA,11 rRNA gene in micronucleus, 6 5 S rRNA genes, 12

Tetraiodothyronine (T4), formation, 214 Thyroglobulin

biosynthesis, 244 lysosomal hydrolysis, 214 mRNA,245 subunit exchange, 244

Thyroid A and B granules, 150 acid phenylphosphatase

in "mitochondrial" pellet, 242 subcellular distribution, 224

acid phospholipase, 231 adenylate cyclase, in plasma membrane

preparations, 241 alkaline phospholipase, 231 antiluminal membrane, 213 catalase, subcellular distribution, 225 cell fractionation studies, 213-231 cholesterol, in endoplasmic reticulum,

225 CMP-N-acetylneuraminic acid: GM3-

sialytransferase, subcellular distribution, 244

colloid, 213 cytochrome oxidase latency, 217 cytosolic lactate dehydrogenase, 221 deoxyribonuclease, 217 differential pelleting, 218-221 di-Na-phenylphosphatase, 217 endogenous nuclear DNA, 236 endoplasmic reticulum, cholesterol content,

225 enzyme localization, 231-238 follicles, 213 galactosyl transferase, subcellular

distribution, 221, 243 glucose-6-phosphatase, correction in assay

by phenylphosphatase, 229 ,8-glucuronidase latency, 217 ,8-glycerophosphatase, 217 Golgi apparatus, separation from plasma

membranes, 243

Thyroid (cont.) gradient centrifugation, 221-227 guaiacol peroxidase, 233 homogenization, 213-216

gel filtration of homogenates, 218 by microcavitation, 248 by Polytron homogenizer, 248 problems, 214

Index

lipolytic enzymes, subcellular localization, 231-232

liposomes, 241, 242 Long-Acting Thyroid Stimulator (LATS)

Inhibitor, 247 Iysophospholipase, 231 Iysosomes, 241, 242

acid ribonuclease activity, 228 contamination with mitochondria, 218 enzyme markers, 228 heterogeneity, 224 hydrolases, 217 phosphodiesterase activity, 228

mannosyltransferase, subcellular distribution, 221

marker enzymes, 216-218 membranes, damage by homogenization,

214 microsomal fraction, effect of sonication,

229 mitochondria

cardiolipin content, 227 contamination with Iysosomes, 218

mitochondrial/lysosomal fraction, 222 neutral phospholipase A2, 232 N-acetylglucosaminyltransferase,

subcellular distribution, 243 N-acetylhexosaminidase, in

"mitochondrial" pellet, 242 nitrogen cavitation, 248 nuclei,238-241

glucose-6-phosphatase, 229 lipids, 246 methods of isolation, 216 monolayers, 241 NADPH-cytochrome c reductase, 229 peroxidase, 229 RNA polymerase, 236

organelles, 238-251 peroxidase, subcellular distribution, 232-

234 peroxisomes, catalase activity, 225

Index

Thyroid (cant.) phenyl phosphatase, and glucose-6-

phosphatase, 229 phosphodiesterase, 217 phospholipase A, Ca2+ stimulation, 231 phospholipase A" and lysosomal markers,

232 phospholipases, 231 plasma membranes, 245-251

adenylate cyclase, 219, 249 alkaline phosphatase, 222 ATPase, 222 cholesterol, 219, 247 contamination, 246 heterogeneity, 250 lysosomal contamination, 250 method of isolation, 245 mitochondrial contamination, 250 (Na+, K+)ATPase, 249 5'-nucleotidase, 185,249 phospholipid content, 247 separation from Golgi apparatus, 243 sialic acid, 219, 247

polyribosomes, 244, 245 and thyroglobulin synthesis, 245

protein synthesis, 244, 245 ribonuclease, 217 RNA polymerase

extraction from nuclei, 236 forms lA, I., IlIA, and III., 234 possible bimodal localization, 238 subcellular localization, 234-238

rough endoplasmic reticulum, effect of heparin, 225

sialyltransferase, subcellular distribution, 221,243

subcellular biochemistry, 213-251 subcellular fractionation, technical

problems, 251 subcellular localization of

lipolytic enzymes, 231, 232 peroxidase, 232-234 RNA polymerase, 234-238

thyroglobulin biosynthesis, 244 TSH-sensitive adenylate cyclase, 249 zonal centrifugation, 222

Tibric acid, effect on peroxisomes, 180

Toyocamycin, effect on pre-mRNA biosynthesis, 53

Transcription, of rRNA genes, 16-31 in vitro studies, 29-31 transcription complex, 16-18

Transcriptional control, of ribosome biogenesis, 46-51

Transcription products, 45 S nucleolar, 30 Transcription units

definition, 6 and r-chromatin, 20 rDNA heterogeneity, 27 for rRNA genes, 2 varia tion in length, 27

"Transcripton," 6 Transglycosylation, bacterial cell-wall

biosynthesis, 323

411

Translational diffusion constant, rRNA, 85 Transpeptidation, in bacterial cell-wall

biosynthesis, 323 Triglycerides, in Ricinus communis, 184 Triiodothyronine (TJ), formation, 214 Triton X-lOO, solubilization of M.

/ysodeikticus membranes, 355 Triturus, rRNA genes, 19 Triturus a/pestris, extrachromosomal rRNA

genes, 5 tRNA

binding to A site of rRN A, 16 T,yCG sequence, 16

TSH-sensitive adenylate cyclase, 249 Turnover of ribosomes, 54 Tyrosine aminotransferase, synthesis by

membrane-bound polysomes, 128

Ubiquinol-cytochrome c oxidoreductase, 300

UDP-muramylpentapeptide, 324 UDP-N-acetyl-D-glucosamine, in

peptidoglycan biosynthesis, 323 Uricase biosynthesis, 202 Uridine, pre-rRNA conversion to

pseudouridine,32

Vectorial discharge of rough endoplasmic reticulum proteins,

143 of secretory proteins, 117

Verrucarin A effect on polysome spirals, 141 initiation inhibition, 147

412

Vesicular stomatitis virus (VSV) glycoprotein G, 192 membrane biogenesis, 191 viral envelope, 132

Vesiculation, and peroxisome biogenesis, 189

Wy-14643, effect on peroxisomes, 180

Xenopus laevis anucleolate mutants, 9 oocytes

injection of cloned 5 S rONA, 31 oocyte 5 S rONA, 13

Xenopus laevis (cont.) r-chromatin, 18 rRNA genes, 19 somatic 5 S rONA, 13

X-ray scattering of ribosomes, 100

Yeast mitochondrial F,-ATPase, 280 mitochondrial surface ribosomes,

195

Index

Zonal centrifugation, thyroid mitochondrial/ lysosomal fraction, 222

subcellular biochemistry: volume 7

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