Progress in Nucleic Acid Research and Molecular Biology, Volume 6

PROGRESS IN

Nucleic Acid Research and Molecular Biology Volume 6

Contributors to Volume 6

G. P. GEORGIEV AHARON GIBOR S. GRANICK W. GUSCHLBAUER H. HYDEN J. MASSOULIE A. M. MICHELSON KIN-ICHIRO MIURA SEVER0 OCHOA ROBERT P. PERRY CHARLES WE ISSMAN N STEPHEN ZAMENHOF

PROGRESS IN

NucIeic Acid Research and Molecular Biology edited b y

J. N. DAVIDSON

WALDO E. COHN

Department of Biochemistry The University of Glasgow Glasgow, Scotland

Biology Division Oak Ridge National Laboratory Oak Ridge, Tennessee

Volume

6

7967

ACADEMIC PRESS New York and London

COPYRIGHT@ 1967, BY ACADEMICPRESS INC. ALL RIGHTS RESERVED. NO PART O F T H I S B O O B MAY B E REPRODUCED I N ANY FORM, BY PHOTOSTAT, MICROFILM, OR ANY O T H E R MEANS, WITHOUT W R I T T E N PERMISSION FROM T H E PUBLISHERS.

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LIBRARY OF CONGRESS CATALOGCARDNUMBER:63-15847

P R I N T E D I N T H E UNITED STATES OF AMERICA

List of Contributors Numbers in parentheses refer to the pages on which the authors’ contributions begin.

G. 1’. GEORGIEV (259), Institute of Molecular Biology, Academy of SciUSSR ences of USSR,MOSCOW, AHARON GrnoR (143), Rockefeller University, New York, New York

S . GRANICK (143), Rockefeller University, New York, New York W. GUsCHLnAUER (83), Ddpartment de Biologie, Centre d’Etudes N u cldaires de Saclay, Gif-sur- Yvette,France H. HYDI~N (187), Institute of Neurobiology, Medical Faculty, University of Goteborg, Goteborg, Sweden

J. M A S S O U L(83), I ~ Institut de Biologie Physico-Chimique, Paris, France A. M. MICHELSON (83), Institut de Biologie Physico-Chimique, Paris, France KIN-ICHIRO MIURA(39), Institute of Molecular Biology, Faculty of Science, Nagoya University, Chikusa, Nagoya, Japan SEVEROOCHOA(353), Department of Biochemistry, New York University SchoolJof Medicine, New York, New York ROBERTP. PERRY(219), Department of Molecular Biology, The Institute for Cancer Research, Philadelphia, Pennsylvania CHARLESWEISSMANN(353), Department of Biochemistry, New York University School of Medicine, New York, New York STEPHENZAMENHOP(1), Department of Medical Microbiology and Immunology, Department of Biological Chemistry, and Molecular Biology Institute, University of California, Los Angeles, California

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Preface I n this sixth volume of Progress i n Nucleic Acid Research and Molecular Biology we have endeavored as usual to preserve the balance between the chemical and biological aspects and to include topics of wide general interest at the present time. We should once more point out that it is not our intention to sponsor an annual or fixed-date publication in which literature appearing in a given period of time is summarized, as in the more customary type of bibliographic review or literature survey. As we have emphasized on several previous occasions, our aim is to present “essays in circumscribed areas” in which recent developments in particular aspects of the field of nucleic acids and molecular biology are discussed by workers provided with an opportunity for more personal expression than is normally met in review articles. To this end it is our policy to encourage discussion, argument, and speculation, and the expression of points of view that are individualistic and perhaps even controversial. It is, of course, to be expected that different authors will interpret this charge in different ways, some essaying a broad and philosophical vein, some developing or describing new theories or techniques, some taking the opportunity to assemble a number of fragmentary observations into a coherent pattern, and some reviewing a field in a more conventional manner. We have not attempted to define or restrict any author’s approach to his chosen subject, and have confined our editing to ensuring maximum clarity to the reader, whom we envisage to be a person himself active in or concerned with the general field of nucleic acids and molecular biology. Needless to say, we do not necessarily*share all the opinions or concepts of all the authors and accept no responsibility for them. We seek rather to provide a forum for discussion and debate, and we will welcome further suggestions from readers as to how-this end may best be served. Indeed, we should like to encourage .readers to write to us with their comments. Abbreviations used for nucleic acids and their derivatives are now fairly well established by international authority. Those pertinent to our subject are not listed at the beginning of each chapter, but will be found on the following page.

J.N.D. W.E.C. Pebruarfy, 1967

vii

Abbreviations and Symbols Abbreviations used1without definition are those recommended by the IUPAC-IUB Combined Commission onFBiochernica1 Nomenclature, as printed in the J. Biol. Chem. 241,527 (1966), Biochim. Biophys. Acta 108, 1 (1965), Biochemistry 5, 1445 (1966), Arch. Biochem. Biophys. 115, 1 (1966), Vi/5irology 29, 480 (1966), and Biochem. J. 101, 1 (1966).

*,

A, c, G, I, u, T, x N pu, PY AMP, CMP, GMP, IMP, UMP, qMP, TMP, XMP, etc. dAMP, etc. 2’-AMP, 3’-AMP, (5’-AMP), etc.

ribonucleoside residues in polymers (specific) ribonucleoside residues in polymers (general) purine, pyrimidine ribonucleoside (general) 5’-phosphates of the above nucleosides

5‘-phosphate of 2’-deoxyribosyl adenine, etc. 2’-, 3’-, (and 5’-, where needed for contrast) phosphate of adenosine, etc. 5’-(pyro)diphosphate of adenosine, etc. ADP, etc. 5’-(pyro)triphosphate of adenosine, etc. ATP, etc. inorganic orthophosphate and pyrophosphate Pi, PPi 3’45’ polymer of ribonucleotide N poly N, or (Wn,or (rNIn 3’45’polymer of deoxyribonucleotide N poly dN, or (dN)” 3’45’ copolymer of N-N’-N-N’in regular, poly (N-N’), or r(N-N’)alternating, known sequence or (rN-rN’), 3‘-W copolymer of dN-dN‘-dN-dN’- in poly d(N-N’), or d(N-N’), regular, alternating, known sequence or (dN-dN’), 3’45’ copolymer of N and N‘ in random polyI(N, N‘) or (N, N‘), sequence two chains, generally or completely associated P O ~ Y.(A)-PO~Y (B) or (A).(B) two chains, association unspecified or unknown P O ~ Y (A), P O ~ Y(B) or (A),(B) P O ~ Y(A) P O ~ Y(B) or (A) (B) two chains, generally or completely unassociated ribonucleic acid or ribonucleate RNA deoxyribonucleic acid or deoxyribonucleate DNA messenger RNA; ribosomal RNA mRNA; rRNA nRNA nuclear RNA transfer RNA (RNA that accepts and transfers tRNA amino acids; amino acid-accepting RNA) “Charged” tRNA (tltNA carrying aminoacyl Aminoacyl-tRNA residues) the transfer RNA molecule that normally Alanine-tRNA or accepts alanine, etc. tRNAAla,etc. Alanyl-tRNA*la or the same, with akanyl residue covalently linked Ala-tRNAA1a RNase, DNase ribonuclease, deoxyribonuclease

+

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I n naming enzymes, the recommendations of the Commission on Enzymes of the International Union of Biochemistry (1965) are followed as far as possible. viii

Contents LIST OF CONTRIBUTORS. . . . . . . . . PREFACE . . . . . . . . . . . . . . . SYMBOLS AND ABBREVIATIONS. . . . . . . . CONTENTS OF PREVIOUS VOLUMES. . . . . . ARTICLES P L A N N E D FOB FUTURE VOLUMES . . .

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Nucleic Acids and Mutability

STEPHENZAMENHOF I . Introduction . . . . I1. Definitions . . . . I11. Detection of Mutations . I V. Inheritance of Mutability V . Summary and Conclusions References . . . .

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Specificity in the Structure of Transfer RNA

KIN-ICHIRO MIURA I . Introduction . . . . . . . . . . I1. Base Composition of tRNA . . . . . I11. The Arrangement of Nucleotides in tRNA . IV . The Three-Dimensional Structure of Transfer V. Studies on the Functional Sites in tRNA . . References . . . . . . . . . .

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RNA . . . . . . . . . . . . . . . . . . . . . .

39 40 43 59 63 76

Synthetic Polynucleotides

A . M . MICHELSON. J . MASSOULIE. AND W . GUSCHLBAUER

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I Introduction . . . . . . . . . . . . I1. Preparation of Polynucleotides . . . . . . I11 Techniques for Investigating the Physical Chemistry I V Homopolynucleotides . . . . . . . . . V Polynucleotide Complexes . . . . . . . . VI . Role of Sugar Phosphate Backbone . . . . . VII . Reversibility . . . . . . . . . . . . VIII Displacement Reactions . . . . . . . . I X . Polynucleotide Analogs . . . . . . . . . X Theory and Practice of Helix-Coil Transitions . . XI Factors Governing Structure . . . . . . . References . . . . . . . . . . . . ix

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84 84 85 . . . . . . 98 . . . . . . 104 . . . . . . 119 . . . . . . 122 . . . . . . 123

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CONTENTS

The DNA of Chloroplasts. Mitochondria. and Centrioles S. GRANICKA N D AHARONGIBOR I . Introduction . . . . I1. Chloroplast DNA . . I11 Mitochondria1 DNA . IV . Centriole DNA . . . V The Role of Cytoplasmic VI . Summary . . . . . References . . . .

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. . . Nucleoids . . .

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. 143 . 146 . . . . . . . . . . . 161 . . . . . . . . . . . 172 in Inheritance . . . . . . . 179 . . . . . . . . . . . 182 . . . . . . . . . . . 183 . .

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Behavior. Neural Function. and RNA H. H Y D ~ N I . Introduction . . . . . . . . . . . . . . . . . . I1. Problems Discussed . . . . . . . . . . . . . . . . I11. Methods of Analysis . . . . . . . . . . . . . . . I V. Biosynthesis of Rapidly Labeled RNA in Brain Cells . . . . . . V . Acidic Proteins Specific for the Brain . . . . . . . . . . . VI . Base Ratios of RNA during Physiological Stimulation . . . . . . VII . Base Ratios of RNA during Chemical Induction of RNA Synthesis . . VIII . The Emergence of RNA Rich in Adenine and Uracil during Learning Experiments . . . . . . . . . . . . . . . . . I X . The Possible Transfer of RNA between Glia and the Associated Neuron X . The Synthesis of an Asymmetric. Adenine-Rich RNA in Parkinson’s Disease . . . . . . . . . . . . . . . . . . . X I . A Working Hypothesis . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . .

187 190 191 194 197 199 199 202 208 210 213 217

The Nucleolus and the Synthesis of Ribosomes

ROBERTP . PERRY I . Characterization of the Genes Coding for Ribosomal I1. Synthesis of the Precursor of Ribosoma.1RNA . . I11. Subsequent Events in the Formation of RNA . .

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I . Introduction . . . . . . . . . . . . . . . . . I1. Phenol Fractionation of Nuclear RNA’s and Their Characteristics .

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IV Appearance V. Synopsis . Addendum References

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The Nature and Biosynthesis of Nuclear Ribonucleic Acids G. P. GEORGIEV

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CONTENTS

111. Fractionation of Subnuclear Structures and Nuclear Ribonucleoproteins IV. The Biosvnthesis of RNA in Nuclear Structures and Its TransDort to the Cytoplasm . . . . . . . . . . . . . . . . . . V . Conclusion . . . . . . . . . . . . . . . . . . Note Added in Proof . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . .

288 304 340 341 342

Replication of Phage RNA CHARLESWEISSMANNAND SEVERO OCIIOA I . Introduction . . . . . . . . . . . . . I1. General Properties and Biology of I’LNA Phages . . I11 Mutants of RNA Phages . . . . . . . . . . . I V . Messenger Function of Phage RNA . . . V Replication of Phage RNA . . . . . . . . . V I . Conclusions and Summary . . . . . . . . . VII . Appendix: Identification and Analysis of Viral RNA . References . . . . . . . . . . . . .

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353 354 355 357 358 . . . . . 386 . . . . . 387 . . . . . 395

AUTHORINDEX . .

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SUBJECTINDEX . .

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Contents of Previous Volumes Volume 1 "Primer" in DNA Polymerase Reactions

F. J. BOLLUM The Biosynthesis of Ribonucleic Acid i n Animal Systems

R. M. S. SMELLIE The Role of DNA i n RNA Synthesis

JERARDHURWITZ AND J. T. AUGUST Polynucleotide Phosphorylase

M. GRUNBERG-MANAGO Messenger Ribonucleic Acid

FRITZLIPMANN The Recent Excitement in the Coding Problem

F. H. C. CRICK Some Thoughts on the Double-Stranded Model of Deoxyribonucleic Acid

AARONBENDICH AND HERBERT S. ROSENKRANZ Denaturation and Renaturation of Deoxyribonucleic Acid

J. MARMUR, R. ROWND,AND C. L. SCHILDKRAUT Some Problems Concerning the Macromolecular Structure of Ribonucleic Acids A. S. SPIRIN The Structure of DNA as Determined b y X-Ray Scattering Techniques

VITTORIO LUZZATI Molecular Mechanisms of Radiation Effects

A. WACKER

AUTHORINDEX-SUBJECTINDEX Volume 2 Nucleic Acids and Information Transfer

LIEBEF. CAVALIERI AND BARBARA H. ROSENBERG Nuclear Ribonucleic Acid

HENRYHARRIS

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xiv

CONTENTS OF PREVIOUS VOLUMES

Plant Virus Nucleic Acids

ROYMARKHAM The Nucleases of Escherichia coli

I. R. LEHMAN Specificity of Chemical Mutagenesis

DAVIDR. KRIEG Column Chromatography of Oligonucleotides and Polynucleotides

MATTHYS STAEHELIN Mechanism of Action and Application of Azapyrimidines

J. SKODA The Function of the Pyrimidine Base i n the Ribonuclease Reaction

HERBFJRT WITZEL Preparation, Fractionation, and Properties of sRNA

G. L. BROWN

AUTHORINDEX-SUBJECT INDEX Volume 3 Isolation and Fractionation of Nucleic Acids

K. S. KIRBY Cellular Sites of RNA Synthesis

DAVIDM. PRESCOTT Ribonucleases in Taka-Diastase: Properties, Chemical Nature, and Applications

FUJIOEGAMI,KI':NJI TAKAHASHI, AND TSUNEKO UCHIDA Chemical Effects of Ionizing Radiations on Nucleic Acids and Related Compounds

JOSEPHJ. WEISS The Regulation of RNA Synthesis in Bacteria

FREDERICK C. NEIDHARDT Actinomycin and Nucleic Acid Function

E. REICHAND I. H. GOLDBERG De Novo Protein Synthesis in Vitro

B. NIRMAN AND J. PELMONT

CONTENTS OF PREVIOUS VOLUMES

Free Nucleotides in Animal Tissues

P. MANDEL

AUTHORINDEX-SUBJECT INDEX Volume 4

Fluorinated Pyrimidines

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CHARLES HEIDELBERGER Genetic Recombination in Bacteriophage

E. VOLKIN DNA Polymerases from Mammalian Cells

H. M. KEIR The Evolution of Base Sequences in Polynucleotides

B. J. MCCARTHY Biosynthesis of Ribosomes in Bacterial Cells

SYOZOOSAWA 5-Hydroxymethylpyrimidines and Their Derivatives

T.L. V. ULRRICHT Amino Acid Esters of RNA, Nucleosides, and Related Compounds

H. G. ZACHAU AND H. FELDMANN Uptake of DNA by Living Cells

L. LEDOUX AUTHORINDEX-SUBJECT INDEX Volume 5

Introduction to the Biochemistry of D-Arabinosyl Nucleosides

SEYMOUR S. COHEN Effects of Some Chemical Mutagens and Carcinogens on Nucleic Acids

P. D. LAWLEY Nucleic Acids in Chloroplasts and Metabolic DNA

TATSUICHI I WAMURA Enzymatic Alteration of Macromolecular Structure

P. R. SRINIVASAN AND ERNESTBOREK Hormones and the Synthesis and Utilization of Ribonucleic Acids

J. R._TATA

xvi

CONTENTS OF PREVIOUS VOLUMES

Nucleoside Antibiotics

JACKJ. F o x , KYOICHI A. WATANABE,

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ALEXANDER BLOCH

Recombination of DNA Molecules

CHARLESA. THOMAS, JR. Appendix I. Recombination of a Pool of DNA Fragments with Complementary Single-Chain Ends

G. S. WATSON, W. K. SMITH,

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CHARLESA. THOMAS, JR.

Appendix II. Proof That Sequences of A, C, G, and T Can Be Assembled to Produce Chains of Ultimate length Avoiding Repetitions Everywhere

A. S. FRAENKEL AND J. GILLIS The Chemistry of Pseudouridine

ROBERTWAQNERCHAMBERS The Biochemistry of Pseudouridine

EUGENE GOLDWASSER AND ROBERT L. HEINRIKSON

AUTHORINDEX-SUBJECT

INDEX

Articles Planned for Future Volumes The Mutagenic Action of Hydroxylamine

D. M. BROWNAND J. H. PHILLIPS The Search for Messanger RNA of Hemoglobin

H. CHANTRENNE, A. BURNY,AND G. MARBAIX Autoradiographic Studies on DNA Replication in Normal and leukemic Human Chromosomes

F. GAVOSTO Ribonucleic Acids and Information Transfer in Animal Cells

A. A. HADJIOLOV Proteins of the Cell Nucleus

L. S. HNILICA RNA and Protein Synthesis in the Early Embryo

M. NEMER Enzymatic Reduction of Ribonucleotides

P. REICHARD AND A. LARSSON The Photochemistry, Photobiology, and Repair of DNA

R. B. SETLOW Cytochemical localization of Nucleolytic Enzymes

D. SHUGAR AND H. SIERAKOWSKA Oligonucleotide Separation

H. A. SOBERAND G.

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RusrnzKY

The Genetic Code

C. R. WOESE

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Nucleic Acids and Mutability'

STEPHENZAMENHOF Department of Medical Microbiology and Immunology, Department of Biological Chembtw, and Mokcular Biology Institute, University of California, Los Angeles, California

I. Introduction . . . . . . . . . 11. Definitions . . . . . . . . . . A. Spontaneous and Induced Mutations . B. Definition of Mutation . . . . . 111. Detection of Mutations . . . . . . IV. Inheritance of Mutability. . . . . . A. Modes of Inheritance of Mutability . . B. Mutability end Development; Protection C. Regulation of Changes in Mutability . V. Summary and Conclusions . . . . . References . . . . . . . . . .

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against Mutations

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1. Introduction Mutations, the discontinuous events that lead to (or are) changes of genetic information, are among the most important biological phenomena. Mutations furnish the raw material of evolution ( I ) ; mutations may be a t the root of changes (in the host cell and/or the microorganism) that lead to the onset of malignant growth, the outbreaks of epidemics, and hereditary diseases. Mainly because of their visible manifestations, mutations were considered interesting, on and off, since 1650 when the phenomenon was first described ( 2 ) . More modern is the recognition of the importance of the quantitative aspect of this phenomenon, mutability. Obviously, it is this aspect that decides whether the phenomenon is a mild curiosity or a curse. Despite this, the quantitative aspect still This study was aided by grants CA-08128 and HD-01331 from the National Institutes of Health, US. Public Health Service, AT(30-1)-3103 from the U.S.Atomic Energy Commission, and E 5 2 from the American Cancer Society. 1

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STEPHEN ZAMENHOF

remains largely ill-defined and its ramifications are often underestimated. An almost void area that especially deserves study is the problem of inheritance of mutability, since there is no doubt that mutability is heritable (examples in references 3 and 4 ) . In particular, one should ask what is the role of nucleic acids in this special inheritance, and what are the mechanisms involved. It is the purpose of this essay to summarize present2 knowledge on some aspects of this subject; the mutations themselves are not discussed in detail as they have been a subject of recent reviews (6-21). This essay is limited to mutabilities of chromosomal, episomal, and viral nucleic acids; the subject of inheritance in plastids and related entities is considered beyond our scope.2a Similarly omitted is discussion of the role of mutations in evolution, especially since this subject has been dealt with extensively in recent reviews (1, 7-19). Many studies have made it clear that the presently known structures of nucleic acids (1U14) and proteins (16-20) give little or no information about mutability itself, for they are the end result of mutability and of complex phenomena of recombination and natural selection. Ingenious studies a t the molecular level (15, 16) reveal that some mutations are not acceptable to the organism if only because of the requirements of primary, secondary, and tertiary structure of proteins (especially those with enzymatic activities). Thus, most mutations are not registered a t all, a few others are overemphasized, and the mutability spectrum as such is not depicted correctly.

II. Definitions A. Spontaneous and Induced Mutations Although the division of mutations into “spontaneous” and “induced” is a convenient one, it does not seem to have a basic justification. “Spontaneous” mutations must also be induced by mutagens; these mutagens are merely ubiquitous and unknown. The most likely mutagens of this sort are: (1) thermal oscillations (22) leading to mutations by the mechanisms of tautomeric shifts (23, 94) and/or depurination (26, sort are: (1) thermal oscillations (22) leading to mutations by the normal environment of nucleic acids (review in reference 6). The relative importance (degree of participation) of any of these agents may vary from case to case. In this essay, the term “spontaneous” is retained for historical reasons, but whenever possible the two types of mutations are discussed jointly. ’Main references as of March, 1966; references marked “a” added in proof. a* See articles by Iwamura in Volume 5 and by Granick and Gibor in this volume.

NUCLEIC ACIDS AND MUTABILITY

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B. Definition of Mutation Despite voluminous literature, the definition of mutation itself is not generally agreed upon. Since a good definition would probe deeply into the nature of the phenomenon, it cannot be dismissed simply as a matter of semantics. It is to be noted that avoidance of a definition of mutation and related terms has led in the past to considerable misunderstanding among investigators discussing different phenomena under the same name (6, pp. 63-66). The observation of phenotypic changes is still the most practical method of recognizing mutation; in the past, such changes were indeed inherent in definitions of mutation (27). It is clear now that such a definition is basically wrong. As is discussed below, many mutations will never lead to changes in phenotype, due to the degeneracy of the code [recent reviews in (28, M a ) 1. Mutations may occur in genes that are totally inactive, such as injured or “silent sections” of DNA (29). Mutations may occur in cells that die for other reasons (such as “too strong” a mutagen) but still contain replicable genetic material? Other examples are given in Section 111. It is felt that the changes in genetic material itself (nucleic acids) constitute the mutations, regardless of how they manifest themselves. However, even accepting this, the definition of mutations remains difficult. The often used simple definition, “mutation is a change in a gene” (6),is inadequate (6) because, for instance, changes in p H and temperature within certain limits (30-32) may produce changes in nucleic acids (such as a change in ionization) that are transient and nonheritable and clearly are not mutations. The realization of the existence of these transient (phenotypic) changes in nucleic acids demands addition of a limiting statement: “a change in a gene that persists after the mutagen has been removed from the system.” However, such a definition is again meaningless in the case of spontaneous mutations; there, the mutagens that are unknown but not necessarily qualitatively different from “inducing mutagens” cannot be removed from the system; the phenotypic responses of the nucleic acids blend imperceptibly with the genotypic response. A definition of mutation necessitates a definition of the gene, and this, in itself, is still under d i ~ p u t e .The ~ definition “mutation is a *It stands to reason that genetic material cannot be considered dead as long as it is able to replicate under some conditions, either as such or after rccoinbination with other genetic material. ‘For instance, the popular definition of a gene as a ‘(segment of nucleic acid that determines the structure of a protein or a polypeptide” is acceptable only for structural genes; such a definition excludes many regulatory genes.

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change in nucleotide sequence” is improper, because it would define as mutations phenomena that, I feel, are not mutations, such as recombinations, the insertion of temperate phage DNA into host DNA during lysogenization, etc. On the other hand, such a definition excludes phenomena that may be considered mutations, such as the loss of an entire chromosome (in higher organisms). These few examples illustrate the difficulties in devising a simple definition of mutation, and a complex one appears to be the only solution. Thus, one could say that a mutation is a change in the substances (nucleic acids) carrying genetic information, in which ( a ) the change remains upon replication of this substance in the absence of any known mutagens; ( b ) the change is not brought about by simple addition of a substance already carrying genetic information (an episome in general, or a virus nucleic acid in particular) ; (c) the change is not caused by a simple exchange with a substance already carrying genetic information (another chromosome or its part). It is realized that such a definition is, of necessity, clumsy, arbitrary, and only temporary. In particular, others may feel that inheritable addition of unchanged genetic material [addition of an extra chromosome in higher organisms (mongolism) or duplication of a gene, probably due to unequal crossing over] may deserve the name mutation: the sequence of bases in the genes themselves may not have changed but the total amount of genetic information has. An even stronger argument may be that the incorporation of an episome or an exchange (recombination) is a mutation, for here even the sequence of bases has changed. “Selfing” (33) and unequal crossing over as an explanation for the appearance of frequent prototrophs in an auxotrophic population has been suggested (reviewed in reference 34). However, in the case of trsnsduction, the proximity of the transducing fragment rather than “selfing” itself appears to be the cause of higher mutability (35).A special borderline case is crossing over within a triplet (codon) (36) [which may be even more frequent than classical mutations (28)] ; this crossing over may occur in heterozygotes bearing a t the same site codons for two different amino acids (36) or different codons for the same amino acid (28). I n either case, the result is a codon for a third, different amino acid; this phenomenon, which may be termed “allogenic recombination,’ by analogy to allogenic transformation (37‘), cIosely resembles mutation in its effects. These findings add a new dimension to the study of evolution, for if such an intracodon recombination is not a mutation, then evolution, in the sense of addition of new information, can theoretically progress even in the absence of “mutations” and certainly in the complete absence of mutagens.

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NUCLEIC ACIDS AND MUTABILITY

5

The quantitation of mutations is the next problcm that encounters difficulties in definition. For simplicity, let us assuine for the moment that all mutations are detectable. The popular term “mutation ratc” denotes the number of mutational events per biological entity (cell or virus) per generation; the latter has been equated with the replication of nucleic acid. The notion that such a term really describes the outcome of the entire mutational process is open to serious objections (6),as is seen below. Mutation is seldom a single phenomenon. In general, the changes (injuries) brought about by most mutagens (deamination by nitrous acid, depurination by heat, alkylation, pyrimidine dimer formation by UV, etc.) are not those that are retained on replication of nucleic acids5; these injuries (first step) must be translated into changes in base sequence that are acceptable by the mechanisms of replication of nucleic acids (second step). The two steps usually occur a t different times and under different conditions; which of these steps should be considered more important for the final outcome will depend on which step is the rate-limiting reaction. The cases in which the number of mutations increases with the dose of mutagen or the duration of treatment (38, 25), but is independent of the generation time, i.e., frequency of DNA replication (39-41), are examples of the situation in which the first step is most likely to decide the outcome. In other cases, the ratelimiting step appears to be the replication of nucleic acid, as the number of mutations depends on generation time and the number of generations (42, 43). Often both cases (independence or dependence on generation time) can be demonstrated for the same organism by simply changing growth-limiting factors (44,45). A special case is the mutagenic effect of incorporation of base analogs (such as 5-bromouracil) (4660); both steps (first step, incorporation of analog; second step, error on subsequent replication) obviously involve the replication of nucleic acid. The two replication processes may take place under different conditions resulting in different numbers of generations in each. Finally, the “mutation rate” often may not give information about the phenomenon simply because the mutation rate is p e r generation, and the step determining the quantity of mutation may be the one in which a “generation” (replication of nucleic acid) does not occur at all 6The deamination of cytosine to uracil in viral RNA is one notable exception, as uracil is directly acceptable by the mechanism of RNA replication. ‘It is conceivable that in some cases the two steps coalesce into one; the error is committed by the incorporation of analog (e.g., incorporation of Sbromouracil at a site not complementary to adenine). In vitro studies suggest that the process of mutagenesis by analogs is far from understood (61,62).

G

STEPHEN ZAMENHOF

[for instance, when dry spores are heated to high temperatures (2511 ; or, if it does occur, it does not determine the final outcome. Another practical difficulty stems from the finding that analogs can be incorporated and removed from DNA without net D N A synthesis (55-56). This phenomenon, which may be termed “DNA turnover,” is probably due to cryptic breakdown and resynthesis of portions of DNA (67, 68). Thus, even those mutations that depend on DNA synthesis may appear to occur “without a generation” [stationary cultures (69, 60)1. Another complication is caused by the fact that mutations initiated in the first step may be subject to repair in a step intermediary between the first and second. This repair, first demonstrated for W [photoreactivation (Sl)] and for nitrogen mustard ( 6 2 ) , has recently been the subject of extensive study. Some of the mechanisms involved [dark reactivation; excision of thymine dimers followed by resynthesis (63, 6 4 ) ] appear to be applicable to other mutagens as well (65, 66). These phenomena (excision and resynthesis) may also be the basis of the turnover of DNA described previously. Thus, the outcome is determined also by a step whose efficiency has been shown to depend on factors [enzymes (67), intensity of reactivating light, pressure ( 6 2 ) , duration of this step, presence of amino acids (68),chloramphenicol (68), basic dyes (SO)] differing entirely from those determining the first step (intensity of mutagen) or second step (errors during replication). Another term used to quantitate mutations is “mutant frequency.” This term simply denotes the final proportion of mutants detectable in a population, regardless of how and when they arose. This term is useful, provided that one clearly bears in mind the above complexity of the phenomena. I n this article, the term “mutability” denotes the over-all intrinsic ability of sites of informational substance (nucleic acids) to undergo mutation, whether the latter is in practice detectable or not. The limitation in detection is the subject of the next section. From the foregoing, it is clear that a universally acceptable quantitation of “mutability” covering all the steps (including repair) and all the conditions is not feasible. All one can do is compile, from the existing data on mutation rate, mutant frequency, etc., a composite estimate that is valid only in conjunction with the clear realization of all the complexities and all the pitfalls involved.

111. Detection of Mutations Clearly, the above temporary assumption, made for the sake of definition, that all mutations eventually occurring are detectable, is false on several accounts.

NUCLEIC ACIDS AND MUTABILITY

7

The known mutational changes that affect systems carrying genetic information and are still not lethal range from gross aberrations, such as the loss of an entire chromosome (in higher organisms), to point mutations affecting one base or base pair only. The former are often detectable by simple microscopic examination ( I ) , and would be so even if there were no phenotypic expression of mutation: the change (loss) of the genetic component (DNA) is presumably sufficient to establish the fact that a mutation has occurred. A loss of an entire chromosome that is not lethal is a rare event; however, other chromosomal aberrations that are often not lethal (loss of a part of a chromosome, duplication, translocation, inversion) are often also detectable ( 1 ) . The demonstration of the heritable change in appearance of chromomeres might also be considered proof of mutation without reference to phenotypic expression, for it is possibly the change in conformation or loss of the DNA molecules themselves that can be demonstrated on suitably strained preparations. However, the present knowledge of this subject is rather primitive and still cannot compete well with the type of direct evidence one obtains from the study of microorganisms. I n these, large deletions or duplications ought to be directly demonstrable without reference to phenotypic expression. It has been reported (70) that a deletion in phage DNA can be demonstrated (by genetic methods) as a shortening of the distance between the markers on each side of the deletion; this genetic proof is valid regardless of whether or not the gene with the deletion codes for any protein a t all. The most numerous (71-73) and probably the most important mutations may be point mutations in which presumably a single base or base pair is deleted, changed, or inserted. The detection of such a small change without reference to phenotypic expression would require the analysis of the nucleic acid base sequence. The existing techniques, however, are far from enabling us to do so, except for RNA molecules of the size of transfer RNA (74). A difference found in the sequence of nucleotides in transfer RNA would reveal that a mutation has occurred in the DNA segment coding for this RNA. Such a demonstration would be free of some complications inherent in the proof of mutation by analysis of amino acid sequence, in that the degeneracy of the code would not be involved. Attempts to separate a single chemical species of messenger RNA have been reported recently (75-79) ; preliminary evidence indicates that RNA molecules of even smaller sizes can be fractionated almost equally well (79). If and when the analysis of base sequences of such fragments becomes feasible, an even better tool will be available for the direct demonstration of changes in the base sequence of messenger RNA and therefore of mutations in DNA.

8

STEPHEN ZAMENHOF

A special case in which the isolation of a single chemical species of messenger RNA is already feasible is the RNA of the Satellite Tobacco Necrosis Virus (80).This RNA is also the viral genome; i t does not seem to be anything more than a single structural gene for the coat protein (81,82). A mutation in such a virus should indeed be demonstrable directly, as a change in base sequence. Another example of such a demonstration is the evidence for “mutational” in vitro incorporation of guanine into a dA-dT polymer in which some of the thymine molecules are replaced by 5-bromouracil ( 5 1 ) . Such a demonstration is subject a t present to objections against equating the synthesis of DNA in vitro with that in vivo. In general, demonstration of a mutation on the basis of nucleic acid analysis encounters an additional difficulty (applicable also to other cases), namely that the singling out of a suspected mutant among many genetically unchanged cells requires guidance that a t present can only be furnished by phenotypic expression of the mutation. Thus, for the time being, the study of phenotypic expression is still the only practical method for general detection of mutations. However, as is seen below, this method is beset with errors. Two of the reasons that, even on theoretical grounds, preclude the phenotypic expression of all mutations were mentioned in Section I1,B. If, by reason of a “point mutational change,” the sequence of bases in a triplet is changed, this altered triplet may still code for the same amino acid due to the degeneracy of the code; the proportion of mutations that are undetectable for this reason has recently been estimated to be a t least 20% of the total (28,28a). If a mutation occurs in a segment of nucleic acid that has mutated previously to the extent of not producing a protein, or is “silent”‘ (71)for other reasons such as duplication, but still replicates, then further mutations in this segment will be undetectable. A mutation in a genome that already carries an active suppressor of this mutation may also be undetectable. A more common reason for the lack of detection of mutation is that the change produced in an active protein (enzyme) is too small to affect its activity [examples in (36, 83, 28a)l. This may be true in the case of replacement of one amino acid by another of the same general type [“conservative mutation” (83a)] ; it may also be true if the change occur’pIoutside the active center and in general in the region of the protein molecule that is not critical for enzymatic activity or conforination. Less is known about mutations in regulatory genes. Those resulting in a change from inducible to constitutive enzyme production can often he demonstrated; sometimes more difficult is the demonstration of muta-

‘Heterochromatin in higher organisms.

NUCLEIC ACIDS AND MUTABILITY

9

tion in the “regulatory” (allosteric) part of a structural gene. For the reasons given above, and probably many others, a considerable proportion of mutations in regulatory genes must also pass unnoticed. Another class of mutations largely undetectable in haploid organisins is that inactivating “csscntial gencs.”x Onc cxaiiiple is genes that produce enzymes nianufacturing essential high polymers (such as nucleic acids) or genes that produce ribosomal RNA’s or transfer RNA’s themselves. All these products cannot be supplied to the mutant cell from without: the consequence of the inactivation of such genes under all conditions is the death of the organism and failure to score it as a mutant. One should add that some of these mutations may be amenable to study, namely, if special conditions are found in which death does not occur (“conditional lethal mutations”). Other lethal mutations that cannot be scored in haploid organisms are those that produce an intracellular poison that cannot be neutralized. Still others may be lethal by virtue of hindering DNA replication. The deamination of guanine to xanthine has been cited as lethal for the DNA molecule (84). Cross linking of two strands of DNA may occur with many mutagenic agents ( W, nitrous acid, mitomycin, bifunctional alkylating agents) as the first mutational step (see Section I1,B) ; if this step is not translated into a second step acceptable by the mechanism of replication of nucleic acids but remains as is, lethality may or must follow. To overcome the difficulty that mutation cannot be demonstrated in a dead cell, an approach has been devised in which the molecules of DNA (or their fragments) are mutated in vitro (85-88). These molecules can subsequently be used in bacterial transformation for demonstrating, by means of phenotypic expression in bacterial cells, that the mutation in vitro has indeed occurred. In such experiments, the cell itself is never in contact with the mutagen; thus, the procedure is free from many of the objections listed above. Although these techniques are difficult and not universally applicable, the method is certain to find wider use in the future; conceivably, it could also be adapted to recombinations other than transformation. On the basis of all the foregoing difficulties and objections, one might conclude that, since one can account for only an unknown and probably variable fraction of the total mutations occurring, the entire problem of quantitation of mutations, and consequently also the problem of the mechanism of inheritance of mutability, are not yet ready for study. The author feels, however, that the problem is too important to be set aside for an indefinite period, and that, bearing in mind the above Recessive lethal mutations in diploid organisms can sometimes be conveniently scored ; they have been used extensively in Drosophikz genetics.

10

STEPHEN ZAMENHOF

limitations, one can still hope to draw some valid conclusions concerning this latter mechanism and the role of nucleic acid in it.

IV. lnheritunce of Mutability A. Modes of Inheritance of Mutability 1. INTRODUCTION

That the degree of the ability of a gene to undergo mutations, either spontaneous or induced, is transferable from one generation to another has been known for a long time [for bacteria, see Table I (S)]. With modern recognition of the multitude of mutational sites, ultimately as small as a base or base pair, i t was demonstrated in viruses (8S9S) and in bacteria (94, 4, 8S, 7s) that each site has its own characteristic ‘(spontaneous” or induced mutability, which is retained upon reproduction (replication of nucleic acids). The carriers of this information about mutability and possible mechanisms of its transfer from one generation to another will now be discussed. A casual observer may dismiss the problem by saying that, since nucleic acids are the only substances known to carry genetic information, they must, somehow, also carry this information. Moreover, since molecules of DNA are known (at present) to do only two things, to direct the synthesis of their own replicas and to direct the synthesis of RNA, one or both of these processes must also be the mechanism of transfer of information about mutability. As is seen below, the actual situation is much more complex than this incomplete picture. For instance, how can the accepted mechanism of replication without a change (23) transfer the information about replication with a change (mutation)? How can direction of the synthesis of RNA, a substance which is not implicated in a change of base sequence of DNA, be involved in the determination of mutability? To be able to cope with these problems it is first necessary to discuss the modes of inheritance of mutability. 2. MUTATOR GENES

That the mutability of some genes can be influenced (greatly increased) by other genes was discovered rather early in Drosophila (95) and in maize (96, 97). Four different cases of such genes increasing mutability, termed “mutator genes,” were found subsequently in bacteria (98-105): three in E . coli and one in S. typhimurium. It appears that, in bacteria, mutator genes may influence the mutability of essentially all other genes although preference in the direction of specificity

11

NUCLEIC ACIDS AND MUTABILITY

TABLE I FREQUENCIES OF SPONTANEOUS A N D INDUCED REVERSIONS I N E. coli IN THIRTY-FIVE INVOLVING ELEVEN AMINO ACIDS (3) NUTRITIONAL DEFICIENCIES

Strain

Deficiency

12-72 12-22 WP-12 12-29 12-56 M-1 Sd-4-73

Phenylalanine Tryptophan Arginine Leucine Threonine Leucine Methionine or cystine Tryptophan Histidine Cystine Histidine Arginine Methionine+ threonine Leucine Leucine Arginine Tryptophan Leucine Leucine Methionine Methionine Methionine or threonine Threonine Tryptophan Histidine Tryptophan Lysine Proline Histidine Tryptophan Tryptophan Tyrosine Methionine Proline Histidine

M-4 12-91 Sd-4-77 12-23 12-16 WP-7 M-3 12-57 D-84 Sd-4-55 12-72 WP-3 Sd-4-49 12-33 12-66 12-36 12-32 WP-6 WP-12 WP-8 WP-14 R-4-88 WP-2 12-61 WP-10 12-11 12-100 w-74

8-Propiolactone.

Spontaneous ( x 10-9)

MnC12 ( x 10-8)

uv ( X 10-8)

p-pl. ( X 10-8) Q

0.01 0.06 0.06 0.07 0.23 0.25 0.27

11 0 52 25 11 24 802

100 0 510 222 72 1,200 125

3.3 0 4.4 8 2.5 2.4 16

0.31 0.34 0.34 0.39 0.39 0.40

448 121 240 0 1,720 0

10,700 22 148 0 440 0

217 4.3 9 0 96 0

0.89 0.93 1.08 1.20 1.33 1.42 1.45 1.56 1.93

1,050 224 63 10,200 594 4,000 594 940 2,400

6,300 940 4,600 1,800 57 1,070 186 1,020 1,570

2.8 6 16 60 28 65 37 8 380

2,185 1,870 3,560 14,700 2,390 7,240 0 2,310 6,320 1,740 1,333 0 8,000+

435 1,450 2,220 3,110 2,190 9,400 0 3,160 4,650 2,320 1,020 0 5,030+

2.17 2.80 2.88 4.88 5.02 5.16 5.54 5.93 7.27 12.3 12.5 24.8 37.4

55 147 65 291 232 2,260 0 129 45 100 8,130 0 4,150+

12

STEPHEN ZAMENHOF

of mutations has been noted (100,106, 107). It was suspected that the mutator genes somehow affect DNA (loo),either by lowering its stability or by producing a mutagenic s u b ~ t a n c eHowever, .~ evidence for such mechanisms has not been obtained. The suggestion has also been made (104)that a mutator gene may produce a mutagenic base analog, but evidence for this has not been presented. Recently, more light has been shed on this subject. A genetic determinant for generalized high mutability in a wild strain of E . coli (99) has been transferred by sexual recombination to E. coli K-12, and its genetic mapping and expression in this system have been studied (108). This mutator gene, designated ast, has essentially the same effect on E . coli K-12 as on its original host and serves to elevate bacterial mutation rates up to about 1000-fold (108).I n addition, the presence of ast in a host serves to elevate the mutation rates of virulent bacteriophage and episomic elements (109). One could speculate that increased mutability is a result of a mutation in a structural gene controlling the production of DNA-polymerase. Such a defective enzyme might produce more errors (mutations) during DNA replication. A gene controlling the production of T 4 phage DNA-polymerase has been discovered recently (110, I l l ) , and preliminary evidence (112) suggests that phage carrying some mutations of this gene indeed have a mutation (reversion) frequency elevated up to 2000-fold. Thus, this mutant gene appears to be a phage “mutator gene.” Different alleles of this gene seem to affect different sites of the rII gene and to produce . the other hand, two different different “hot spot” patterns ( 1 1 2 ~ )On bacterid mutator genes also produce a small but significant increase in the mutation rate of phage T4 that has its own DNA-polymerase (109,113). Hence, these results are a t present difficult to interpret. A different class of mutational influences are the cases in which an episome (prophage) increases mutation rates of its host (114). The increase is considerable (total auxotroph frequency 1 to 3%), and the process seemingly resembles lysogenic conversion to a mutator phenotype. However, the phenomenon seems to be basically different from the typical action of a mutator gene in that high mutability is induced only during the infection and lysogenization ; after establishment of the prophage, mutability reverts to normal. A case in which higher mutability appears to be caused by the proximity of a transducing fragment has bcen described (35). Other cases of episome-induced high mutability have also been reported (115-117); in some cases the site of action of the episome is a suppressor gene (115,117) that affects one or a few ‘This substance could be classified as a special intracellular mutagen.

NUCLEIC ACIDS AND MUTABILITY

13

loci only. I n a t lcast one case high mutahility appears to be permanent (115).The mechanism of episomally mediated increased mutability is largely unknown but some of the phenomena may involve a change in base sequence of host DNA by insert)ion of a base sequence of the cpisoiiie.

3. DNA SEGMENTS AS CARRIERSOF INFORMATION CONTROLLING THEIROWN MUTABILITY As can be seen from the preceeding section, the existence of mutator genes is well-documented. One may ask whether these mutator genes are ubiquitous or exceptional. In other words, is the mutability of all genes determined, as a rule, by mutator genes, or by some other factors? Logically, these other factors could be the DNA of these genes themselves (i.e., DNA other than that of mutator genes). That this could indeed be so was suggested by the pioneering work of Benzer (89, 90, 92, 93), who mapped mutabilities along the DNA molecule of a phage and discovered a lack of uniformity of distribution (spectrum of mutabilities; Fig. 1 ) . I n particular, 40% of the total spontaneous mutations in the rII region of the phage chromosome occurred in only two spots (socalled “hot spots”). The spectra may be different for “spontaneous” and for induced mutations (91, 92) and depend on the choice of mutagen (118-122). Obviously, the spectra do not follow Poisson’s distribution (Table 11). With reference to “spontaneous” mutations, these results, although impressive, still do not unequivocally rule out the presence of a mutator: in each case, the work was performed using one host bacterium only (72, 119), and it was still possible that the characteristic appearance of mutants after growth in this host, and the presence of two “hot spots,” in particular, was determined by some special “mutator gene’’ in the host. To overcome this objection and to explore the situation in bacterial DNA, the author and his collaborators prepared DNA from several donor strains with different mutation rates and of different origin, and used it in the transformation (from prototrophy to auxotrophy) of the same recipient strain ( 4 ) . Each of the resulting transformants was found to exhibit the same mutation rate (back to prototrophy) for a particular marker as the donor from which this DNA was derived (Table 111).Thus, the information determining mutability was present not in the recipient strain but in the segment of DNA transferred by transformation.10 It appears, therefore, that as a rule a DNA segment ‘OIt could be argued that even such a small segment contains a “mutator gene” for sites (loci) within this segment. If, however, this is true of every segment of DNA in every strain, then the concept of a “mutator gene” becomes indistinguishable from the concept that the DNA segment determines the mutability of its loci.

14

STEPHEN ZAMENHOF

AIa

A 46

Atbi

A4c

~ 4 b

Albz

~ 4 0

A31

P3h

63p

A31

b

A50

A5b

A5ci

P5d

A5C2

88

890

B9b

BlO

FIG.1. See opposite page for legend.

determines the mutability of the loci it contains. The determination of mutability by DNA of another special segment (mutator gene) therefore appears to be an exception. These studies raise several questions. How long must the DNA segment be to carry such information about mutability? To obtain some evidence pertaining to this subject, in the next study (164) transformation was performed under conditions in which structural genes immediately adjacent (on both sides) to the one being transformed were not

15

NUCLEIC ACIDS AND MUTABILITY

A2b

A20

A2c A 2 d A2c

A2f

1 83

Q

82

I

A6d

T .0

0

m

la

I FIQ.1. Topographic map of the TII region of T4 phage DNA, showing the frequency of spontaneous mutations observed at the various sites. Each square represents one occurrence. From (9.9).

transferred; thus, the fragment of DNA transferred is probably no larger than one structural gene. Such a study is subject to the objection that the base sequences in the donor and recipient may be essentially the same (except for one improper base that differentiates the auxotroph from the prototroph) ; therefore, no new information (except this one base) is transferred from the donor to the recipient. To overcome this objection, donor strains were used that were sufficiently unrelated to the recipient strain so that transformation proceeded with great dif-

TABLE I1 NUMBERS O F GENETICSITES AT WHICH n MUTATIONS HAVE BEEN OBSERVED" MUTANTS OF PHAGE T4 (181)

FOR I N D E P E N D E N T L Y ISOLATED

rII-TYPE

n 1

2

3

4

5

6

7

8

9

10 11

12 13 14 15 16 17 18 19 20 24 27

The numbers in parentheses are sites at which several spontaneous mutants were observed in the original standard type stock; these sites are included in the main figures but they probably belong to the background of spontaneous mutants. The example of a Poisson distribution has been calculated for an average number of 1.2 mutants per site. The number of sites with no mutant would then be 27.

UJ

e

z3 N

17

NUCLEIC ACIDS AND MUTABILITY

ficulty (1% of the normal transformation rate). It has been repeatedly suggested that difficulties in transformation between unrelated strains (“interspecies” transformation) ,I1 or even sometimes between strains that are related (125), are due to different base sequences in the donor and the recipient DNA so that the adequate homology essential for efficient transformation does not exist (126-128). A parallel case is known for transduction between Salmonella typhimurium and E. coli. Such a transduction occurs a t a very low frequency; it has been suggested that this is also due to the lack of homology between the two bacterial genomes a t the molecular level (129,1300). TABLE I11 TRANSFER, BY TRANSFORMATION, OF INFORMATION DETERMINING MUTABILITY IN Bacillus subtilis (4)

DNA donor strain No.5 21 ind+ (Parent 168) 30 ind+ (Parent 168) w22 (Parent 23)

Mutation studied

Donor’s mutation rate

Recipient strain no.

+ his+

5.3

x

10-7

168 ind-

his- + his+

2.0

x

10-0

168 ind-

purr -+ pur+

8.1

x

10-11

168 ind-

his-

References: 26,185. Abbreviations: ind = indole; his After transformation to his- or pur-.

=

Recipient’s mutation rateb (4.3 k 2.5) x 10-7 (2.2 f 0.9) x 10-9 (7.2 k 1.7) x 10-11

histidine; pur

=

purine.

It is of interest that, after transformation of a recipient by DNA from an unrelated strain, the efficiency of the transformed recipient for subsequent transformations by the same DNA greatly increases (126, 127). These findings are consistent with the hypothesis that the transformed recipient now contains a segment of donor DNA that assures some degree of homology in the next transformation. The results of transfer, by transformation between related and unrelated strains, of information determining mutability are summarized in Table IV. It can be seen that, in this case too, the information controlling mutability was transferred unchanged in a DNA segment, probably no longer than one structural gene (124). UThe taxonomy of the species used here (Bacillus subtilis) is not advanced enough to permit decision whether the strains used (Nos. 168; ATCC 6051; ATCC 8188) belong to the same or to different species.

18

STEPHEN ZAMENHOF

This work is far from completed. It remains to be shown that the difficulty in interstrain (interspecies) transformation was not due to something trivial such as a different sequence of neighboring structural genes in the donor and in the recipient (I%?), or to a difference in a single base pair ( l a $ ) ,for such cases have also been reported. It remains to be demonstrated just how different the base sequences in the donor and recipient DNA’s were, i.e., how much of the new information that could determine mutability has been furnished by the donor DNA segment. Above all, it remains to be demonstrated that, indeed, the mutabilTABLE IV TRANSFER, BY TRANSFORMATION BETWEEN RELATED A N D UNRELATED STRAINS, OF INFORMATION DETERMINING MUTABILITY (124) DNA donor strain no.’ 168 168 168 168

Mutation studied

Donor’s mutation rate

Recipient strain no.

Transformation efficiency (%)

his2- + hisz+ try3- + try3+ try3- try3+ try3- ---f try3+

3 . 9 X 10-8 1 . 2 X 10-9 1 . 2 X 10-9 1 . 2 X 10-0

168 168 ATCC6051 ATCC8188

100 100 1 1 to 10

--$

Recipient’s mutation rateb 3.2 X 0.8 x 1.2 x 1.3 X

10-8 10-9 10-9 10-9

a All strains are classified as Bacillus subtilis. References: hisz- (183); try3- (131). Abbreviations: his = histidine; try = tryptophan. After transformation to hisa- or to try8- by the donor’s DNA. The neighboring structural genes were not transferred in these transformations, indicating that the DNA segment that was transferred was not longer than one structural gene.

ity of a site in the recipient DNA differs from that of the identical site in the unrelated donor strain. If one accepts this concept and the preliminary evidence that a segment of DNA not longer than a structural gene carries information about the mutability of bases within this segment, then the above considerations lead to the following question: in what form (in what new coding) is this information registered in the DNA molecule? It has been suggested that this information is contained in the sequence of neighboring bases (91, 4 ) . It is conceivable that a reaction [tautomeric shift (28) or depurination (26, 26) ] triggering a “spontaneous” mutation (first step) may proceed a t a different rate for a purine situated in a tract of purines than for one flanked by pyrimidines. It is also conceivable that a given base pair may separate more easily if it is flanked by two A-T pairs (two H bonds each) than by two G-C

19

NUCLEIC ACIDS A N D MUTABILITY

pairs (three H bonds each) ( 9 3 ) . However, to date, experimental evidence on this subject is scanty and indirect. It is logical to assume that the distribution of bases in D N A will be influenced by the (G C)/(A T) (i.e., G/A) ratio; a relationship between this ratio and some dinucleotide frequencies has indeed been pointed out (133).It appeared of interest to study whether depurination, one of the possible causes of some spontaneous and of some induced mutations [high temperature (2.5, 26) , alkylation (134-137) 1, also depends on the G/A ratio. This study [Table V (138)l reveals that this is not the case: the proportion of guanine or of adenine released by heating DNA under the same conditions (lOO"C, 24 hours) appears to be es-

+

+

TABLE V RELEASEOF PURINES FROM DNA's

DNA origin Sarcina lutea E. coli Calf (thymus) Streptococcus faecalis a

OF

VARIOUS G-C CONTENT UPON HEATING

Intact

DNA

Mole % G-C

G/A

G*

Ab

G/A

72 51 43 36

2.57 1.04 0.75 0.56

13.8 13.0 13.6 14.9

10.1 9.5 9.9 10.8

1.37 1.37 1.37 1.38

Purines released

lOO"C, 24 hours, 0.005 A4 phosphate buffer, pH 6.8. Per cent of total guanine or adenine present in intact DNA.

sentially the same, regardless of the species from which the DNA was derived and independent of the G/A ratio in these DNA's. Thus, there is no evidence that the over-all loss of a purine by thermal oscillations depends on the content of G-C pairs, or on the over-all density of this purine along the D N A molecule ; it could be, however, that individual purines can be influenced by local peculiarities of base distribution, or that heating under these conditions was not representative of depurination occurring a t biological temperatures. This (138) and a previous (126) study reveal that the proportion of guanine released by thermal oscillations is always higher than the proportion of adenine so released (G/A = 1.4). Such a mechanism would facilitate depurination after alkylation (see above), which was suggested as a cause of the mutagenic effect of such treatment (134-137). It has also been suggested (139, 140) that helix stability plays a predictable role in mutational alterations of DNA. Model experiments

20

STEPHEN ZAMENHOF

with synthetic polyribonucleotides have shown the helix-coil transition of an (Ap),N.poly U or a poly (A, N) Spoly U complex to be different for N = C, G, and U, presumably due to differences in stacking energy for different dinucleotides. A base interacting weakly with its two neighbor bases is likely to loop out with a higher frequency than a base interacting more strongly, Since it has been demonstrated that some mutagens [nitrous acid (87), hydroxylamine (SS)] react more strongly with denatured DNA than with native DNA, it is conceivable that such denatured (looped out) sections will also be more susceptible to these mutagens. Another possible mechanism of the influence of base sequences upon mutability is through the recognition of these sequences by enzymes. I n in vitro experiments, Kornberg and his collaborators (51, 52) have demonstrated the incorporation of guanine (a “mutational” change) into poly (dA-dBrU), i.e., a polymer of the poly (dA-dT) type containing 5-bromouracil in place of thymine. Of particular interest was the finding that only half of the guanine residues were incorporated opposite 5-bromouracil, whereas incorporation of all is predicted by the theories of the mutagenic action of this analog; some additional factors (neighboring bases) controlled the insertion of the aberrant half during the synthesis. It was reported (141) t.hat, when both uridine and pseudouridine nucleoside triphosphates were present, RNA polymerase fitted them (opposite adenine) into distinctly different places along the chain ; thus, neighboring bases in the template or in the newly synthesized chain must have influenced the choice between a nucleotide and its analog. Since analog incorporation sites are sites of potential change that may lead to mutation, such potential sites of mutation must also be under the influence of neighboring bases. One should also mention here the interesting hypothesis of Koch and Miller (142), who postulated that DNA polymerases may have certain special properties that reduce the number of potential mutations arising from ionization and tautomerization of purines and pyrimidines. Little is known about the enzymes that repair DNA and thus reduce the final proportion of mutants (see Section 11,B). It is conceivable that these enzymes may also recognize certain sequences along the DNA molecule; if they do, then the final or net “mutability” after repair would also vary along the DNA molecule. One should now mention the most obvious cause of lack of uniformity in mutability along the nucleic acid molecule. Mutagens, as a rule, exhibit preference in attacking certain bases (review in 21). This phenomenon was first demonstrated for alkylating agents, which almost exclusively attack guanine a t nitrogen in the 7 position (1.6s). Heat

NUCLEIC ACIDS AND MUTABILITY

21

(depurination) (26) affects purines only.12 Nitrous acid attacks only adenine, guanine, and cytosine (31); of these, guanine, after deamination to xanthine, may be lethal (84) and therefore such a deamination will not be scored in determination of mutability. Ultraviolet irradiation may preferentially affect sites that contain neighboring thymine residues (148, 148a) in bacterial cells, or some other bases in bacterial spores (149, 160).Hydroxylamine preferentially attacks cytosine and can even differentiate between cytosine, 5-methylcytosine, and 5-hydroxymethylcytosine (151).5-Methylcytosine is frequently situated next to guanine (152); this offers an additional basis for unequal mutability along the DNA molecule. As a result of these preferential actions, the spectra of mutability along the DNA molecule differ for %pontaneous” mutations and for individual mutagens (Table I, Fig. 2, Table 11; see also the beginning of this section) ; they should also differ for a forward and for a reverse mutation if only because in the latter a different base is present a t that site (91,92, 120, 121, 118, 119, Table VI). Some of these results must be interpreted with caution. (1) A “reverse” mutation, restoring enzymatic activity, may not be a true reversion to the parental base sequence but a second change in the same codon or even a second change in some other codon (36,83, 73). (2) Present mapping techniques, using essentially the process of recombination as a tool, may not be sufficiently accurate to resolve one codon and one base within one codon; thus, the incidence of many mutations in “one spot in the DNA molecule” may really refer to a segment containing several bases. (3) The forward mutation might not have been a true “point mutation,” but a deletion, and therefore difficult to repair by reverse mutation. In spite of these pitfalls, one must admit that the composite picture emerging from all these investigations suggests that spectra of mutabilities along the DNA molecule do vary for different mutagens. These spectra are more complex than the maximum of four classes of mutabilities that could result from the different reactivities of each of the four bases. Some of the modifying factors involved were discussed above, in “It can be argued that this will not render immune any part of the DNA molecule, because every site contains a purine, either in one or in the other (complementary) strand. However, it remains debatable whether both strands, after mutation, are replicable; if it were so, then all resulting bacterial colonies would be mosaics (of cells derived from the segregation of mutated and nonmutated strands), but they are not [26, 144-146; 147 (review)]. One explanation that has been offered is that the other (nonmutated) strand is rendered nonreplicable by overly strong action of the mutagen (26, 1 4 7 ) ; another suggestion was that, as a rule, only one strand replicates (1.46, 67) . Incidentally, whatever the explanation, this situation decreases the number of detectable mutants (see Section 111) by a factor of 2. This sort of difficulty can be bypassed by using single-stranded DNA (118).

22

STEPHEN ZAMENHOF

NT 341

E l H Y L METHANE S ULFON ATE

n EM 14

n

n n

2-AMINOPURINE

AP 2111

129 80

AP I00

n

Lu

2.6-DIAMINOPURINE

n

5- BROMOURAC IL

n

N

55

14

5-BROMODEOXYCYT ID1NE

n

R

PROFLAVINE

Alo

Albl

hlDZ

FIG.2. See opposite page for legend.

connection with the influence of neighboring bases and of enzymes. An additional interesting suggestion has been made recently (153). It is well-known that some amino acid substitutions in proteins are not acceptable if only because they would not fulfill the conformational requirements for a stable protein (see Section IV,A,4). May there not be,

-

23

NUCLEIC ACIDS AND MUTABILITY

+

1149 1562

12I4

-

I7

NI 264

I510

-

n n 370

E

80

-

d

-

NT NT 100 73

N1

71

- - a -

n

A2a

A2b

A2c

A2d

A28

A21

FIG.2. Comparison of topography of T4 phage DNA for spontaneous mutations and those induced by various mutagens. Only the first few segments of the rII region are shown. From (93).

in DNA itself, structural regions that cannot be varied without upsetting replication or some other essential functional process? If such regions exist, mutations in them might be lethal and would never be observed; these regions would truly appear to be "cold spots." No evidence on this subject has yet been presented.

OF MUTATIONS IN PHAGES13 INDUCTION

BY

TABLE VI HYDROXYLAMINE, BY ETHYLMETHANESULFONATE A N D B Y NITROUS ACID(118)

Induction frequency X 108 Forward mutation Mut,ation h+ + hi1 h+ + hi2 h+ + hi1 h+ -+ hi2 h+ -+ hi65 hiUR48-hiUR48S

NHzOH

EMSb

<30 <5 <6 <6 <5 20,000

>30 <50 200 >lo0 100 200

Summary of inductiona

Reverse mutation

HNOz NHzOH EMSb HNO, >20 >600 200 >200 400 300

2000 <20 <1 <50

100 100 <2 20 <5

1000 500 2000 300 500

-

-

-

<5

Abbreviat’ons: F, forward mutation; R, reverse mutation; (<10-6); 0, no detectable induction; -, no results available.

* EMS

=

ethyl methanesulfonate.

NHzOH F 0 0 0 0 0

++

EMS

R

F

++

+

0 0 0 0

0

-

HNOi

R

F

R

++ + ++ ++ ++ ++

++ 0 ++ ++ ++ + ++ ++ ++ 0 ++ ++ ++ - ++ -

+ +, high induction frequency (

Inferred base change in forward mutation

T+ C A+G G+A G+-4 G+A C+T

+, low induction frequency

m

r3

3

EZ

N

s

P

Z -4

NUCLEIC ACIDS AND MUTABILITY

25

In summary, none of the evidence to date seems to explain the existence of particularly “hot” spots, such as the two spots that account for 40% of total mutations of the TIIregion in phage DNA (90, 92, 93, 72) (Fig. 1). One must seek additional explanations for this and other cases. These will be discussed in the next section. 4. PROTEINS AS CARRIERS OF INFORMATION PERTAINING TO

DETECTION OF MUTABILITY If the direct detection of mutations occurring in nucleic acids were possible, this section would be of less importance for molecular geneticists: our curiosity about mutability would be satisfied by investigating changes at the site of their primary occurrence (nucleic acids). For the study of evolution, the consideration of mutational changes in proteins is very important in retrospect ( 1 , 8, 9-19), but study of the contributions of individual mutations in statu nascendi has thus far not been very fruitful because of the complexity of intervening recombinations and natural selection (see also Section I). However, since a t present the only practical way of detecting mutations is by studying the changes induced by them in proteins (see Section 111),these changes become of primary importance for the study of mutability itself. The surest proof that mutation has occurred is evidence of a perman e n P change in the amino acid sequence of a protein. I n several painstaking studies (15-20), this evidence has been obtained for mutations that occurred a t some time during the history of the species (15, 16) or of one individual (156) and are now well-established (i.e., present in the multitude of cells). The detection of such changes right after their occurrence (as a demonstration that a mutation has just taken place) requires the singling out of that particular rare cell or virus that has mutated. This is accomplished by taking advantage of the fact that a change in a protein may alter the selective advantage of the cell or virus, and thus serve for enrichment of the mutant carrying the changed protein. Such enriched cultures can now serve to demonstrate the change in the protein. The most drastic “change” is the complete absence of a protein; this, obviously, would occur as a result of a large deletion (“multisite mutation”) including the entire DNA structural gene segment [for bacteria, review in (157)l. However, it is conceivable that the same result could occur even after a point mutation, if the resulting protein I3The reading of the code can be interfered with, and this may also result in a change in the amino acid sequence (164, 166); however, such a change is transient; if nucleic acids are also affected, then the problem can be treated jointly with previously discussed mutational changes.

26

STEPHEN ZAMENHOF

bulkiness, change in polarity, stiffness a t the bends, and conformational changes in general produced by the mutation are incompatible with the structure of a stable protein (15-17). The complete absence of a protein is difficult to demonstrate; more often one can only vouch for the inability of cell extracts to neutralize antibodies against the original protein [ CRM-; recent review in (158)]. The demonstration of a protein or polypeptide that is shorter than normal (159, 160), presumably due t o point mutations in a codon resulting in chain termination (161),has recently been reported. If a complete protein is produced (CRM’) but has a single amino acid change, the mutation may still be demonstrable by primitive methods, but as a rule only if the protein has biological (enzymatic) activity. Of course, the safest and most elegant method is the combination of enzyme assay with the determination of change in the sequence of amino acids (36, 83). It is clear that all these circumstances, including degeneracy of the code and the above-mentioned selection for enrichment, may greatly distort the actual picture of mutability. The reasons for failure to detect mutations have been discussed in Section 111. Here we discuss the possibility that the existence and distribution of mutational “hot spots” may actually have little to do with the structure of nucleic acids but may rather depict the protein itself. Consider, for simplicity, a structural gene that has a completely uniform mutability along its DNA segment. If this segment codes for an enzyme in which all the amino acids of the active center of the enzyme and some residues such as cysteine are critical for binding the substrate and for protein stability, respectively, then a mutational change (forward mutation) substituting any amino acid for one present in the critical area of the native protein will result in a demonstrable mutation [inactive (36,83, 7‘3, 16.2, 163) or absent protein]. The “invariant area” of cytochrome c (16, 16) is an excellent example of such a critical region. Thus, the codons in this region of DNA (particularly the codons for cysteine) will appear to be “hot spots” for forward mutation. I n the reverse mutation, still another amino acid may be substituted in place of the wrong one (36, 83, 73) ; this, however, will not restore function of the critical area, since only one, the original amin.0 acid (e.g., cysteine), will perform properly. Thus, in such areas the reverse mutation that restores activity would be an infrequent event (“cold spot”). The converse is true of noncritical areas (83, 73) [carboxyl-terminal sequences in cytochrome c (16,16)1. Many forward mutations can and do occur without being noticed (scored as “cold

NUCLEIC ACIDS AND MUTABILITY

27

spots”) ; of those that arc noticed, revcm mutations substituting many amino acids other than the original one can restore full activity (36, 83, 73) (more reverse mutations scored, i.e., “hot spots”). Thus, in some cases the observed “hot-spotness” and “cold-spotness” may have nothing to do with the mutability of the DNA itself. True, the information about the distribution of critical areas of the protein resides in DNA; thus, in the end, it is still DNA that determines “hot and cold spots.” However, clearly this particular information in DNA pertains not to mutability itself, but merely to our skill in the detection of mutability. The degree of participation of the spectrum of mutabilities of DNA itself (Sections IV,A,2 and 3) and of the spectrum due to the structure of individual proteins in the over-all detection of mutability probably varies from case to case, and is at present difficult to estimate. The complete “immutability” (no detectable “mutability”) of a codon can indeed be explained logically by the assumption that, for this particular position, both the conformation and the activity of a protein do not suffer at all, even in the extreme case in which any of the nineteen other amino acids substitutes for the original one (zero forward mutability for all mutagens inducing single base pair changes). The cases of zero forward mutability do not lend themselves to study if only because one would not know how to select such mutants that are not different in any observable respect. On the other hand, if a mutation is detectable, it is still not clear how a codon for even the most critical (stringent) position (no substitution allowed by any of the nineteen other amino acids) could be more than nineteen times as mutable as the most permissive one (detectable only if substituted for by one specific amino acid out of the other nineteen possible ones). Actually, forward mutabilities (point mutations) covering a range of a t least 500 to 1 have been reported (72) (Fig. 1 ) . Such a range could be explained if the “hot spot” represents a composite of several “hot codons,” but not if it is a single codon. For instance, the amino acid sequences of all presently known vertebrate and yeast cytochromes c contain an identical “invariant area,” consisting of eleven consecutive amino acid sequences (16). Thus, as mentioned above, all eleven must be important and any change in any of them will be scored as a n inactivating mutation. If the accuracy of mapping of DNA is such that the corresponding codons are scored as one (composite) site, this site will appear very “hot” indeed. Thus, much more work is needed before one can venture to estimate the participation of the DNA structure and of the protein structure in the distribution of over-all detectable mutability.

28

STEPHEN ZAMENHOF

B. Mutability and Development; Protection against Mutations An interesting side problem is the possible suspension of mutability for a period in which the nucleic acids are inactive in the scheme of development or are inactive for other reasons. This situation may affect only part of the genome (e.g., in cells of higher organism) or the entire living entity (viruses without hosts, spores, dry bacteria, spermatozoa, seeds, etc.). It has recently been suggested (8) that the functional activity of a DNA region may influence its mutational activity for the following reasons: (1) Functionally inactive DNA regions tend to be condensed into a smaller area, presumably by means of basic proteins, rendering them less accessible to mutagens. (2) Certain agents (e.g., nitrous acid, hydroxylamine) mutate single-stranded DNA much more effectively than double-stranded regions (87, 88) so that any uncoiling of a given DNA region, such as may occur when DNA serves as a template for messenger RNA formation, may increase its mutagenic response. The data thus far give no evidence of such an effect. There was seen no large difference in the rate of mutation induction by hydroxylamine for the tryptophan region of repressed and constitutive bacteria (164). However, a small difference (within a factor of 3) might have escaped detection. Dry bacterial cells and bacterial spores are mutated easily by heat (26),spores are mutated easily by X-rays (165) and by UV (160), and dry DNA is easily depurinated by heat ( 2 6 ) . I n Drosophila, older spermatozoa carry more mutated genes than do younger (166-168) .14 Spontaneous mutations accumulate in bacteriophage maintained in the complete absence of DNA replication ( 1 6 8 ~ )D. r y seeds can be mutated by heat (169) ; aged pollen and seeds also include more mutant genes [170-173; 174 (review)]. It would be of interest to learn whether the seeds of NeZumbo nucifera (Indian lotus) that were viable after approximately 1000 years of storage (176) did indeed include a high proportion of mutant genes; such a study has not been done, however. It is clear that the possibility of protection depends on the nature of the attacking mutagenic agent. In the inactive or in the dry state there seems to be less protection against heat and irradiation (as either “spontaneous” or inducing mutagens), as compared with protection against chemical mutagens (again, either “spontaneous” or inducing). This latter protection may be offered by steric hindrance in the inactive regions of DNA, by basic proteins, by the second strand of DNA in the double helix, by the chemically nonreactive dry state, etc. Mutability in the inactive state, however, may conceivably be due to cryptic replication of DNA (“DNA turnover”) (see Section 11).

NUCLEIC ACIDS AND MUTABILITY

29

C. Regulation of Changes in Mutability It is clear from the foregoing that the detectable mutability of individual sites and of the cell or virus as a whole is subject to changes during evolution. Some of these changes, especially those resulting from the acceptable changes in a functional protein structure (see preceding sections), may be very slow. Other changes may be very rapid. The appearance or activation of a typical mutator gene may be the result of a single mutation. The appearance in a cell of an episome acting as a mutator may be the result of a single infection (Section IV,A,P). A single mutation may substitute a base that is more susceptible to a “spontaneous” or inducing mutagen (Section IV,A,3). Conceivably, a change in the neighboring base(s) may have a similar effect but no evidence pertinent to this subject is available. Changes of mutability may be of considerable selective advantage or disadvantage and are therefore subject to selection pressure. In general, it is advantageous for the species if the mutability level is low, because most mutations are deleterious. On the other hand, too low a mutability is disadvantageous because of the need for plastic adjustment to environmental changes (review in reference 1 ) . It thus appears that, if the mechanisms for maintenance of a constant mutability level were to occur in biological systems, they would be favored by selection and would establish themselves. The mechanisms whereby a species strives to maintain a constant level of mutability may be manifold [review in reference 1761. There may be hereditary changes that affect primarily nongenic parts of the cell, such as changes in resistance to admission of mutagen to DNA, neutralization of the action of mutagen, etc. In other cases, hereditary changes may affect primarily the DNA itself. Here, one should first mention changes in the structure of DNA that result in changes of its susceptibility to mutagen; one may speculate that the DNA molecule evolved in such a way as to be mutationally rather stable in a normal cellular environment. Other changes are those in mechanisms that repair DNA. The above changes in resistance to the mutagenic action of mutagen may go hand-in-hand with changes in resistance to the lethal action on the cell as a whole. However, the two effects may be nonparallel, especially if the essential substance, most sensitive to the lethal action of the mutagen, is not DNA. I n the former case, the resistance toward lethal action of the mutagen may act as an agent selecting for lower mutability, and, as such, may form a part of the mechanism for the regulation of mutability. Obviously, the reversibility of the above phenomena would also be of advantage for the regulation of mutability.

30

STEPHEN ZAMENHOF

There is evidence for one such mechanism for maintenance of a constant level of mutability (176).The agent used was the potent chemical mutagen, N-methyl-N'-nitro-N-nitrosoguanidine (NTG) (177-1 79). I n the case studied, two mutants of E . coli more resistant to NTG exhibited lower NTG-inducedl6 and lower spontaneous mutability than the parent strain; the higher the NTG resistance, the lower the mutability (Table VII). As a result, after replacement of the original population by these TABLE VII SPONTANEOUS A N D INDUCED MUTABILITY, AND UV RESISTANCE OF MUTANTS TO NTGa RESISTANT Strain ( E . coli)

Resistanceb t o NTG (300 pg/ml)

Resistanceb to uv (600 ergs/mmz)

Spontaneous mutant frequencyC

NTG-induced mutant frequency.

15s 15S/NTG1 15S/NTG2

< 10-0 8 X 10-4 2X

1.5X 6 . 6 X 10-6

1.6 X 5 . 3 x 10-7 1 . 3 x 10-7

2.1 x 10-8 1 . 7 x 10-4 1.7 X

a

3.2

x

10-4

From (176). NTG, N-methyl-N'-nitro-N-nitrosoguanidine.

* Expressed as fraction of population surviving the treatment. c

Mutation to azide resistance. TABLE VIII POPULATION STUDYOF STRAINS SENSITIVE A N D RESISTANT TO MUTAGENMutagen Incubation -(hours) Present

Proportion of original cells in a mixture of original (mutagen-sensitive) and mutagen-resistant strains

0 12 24

-1

x 5x 3

10-1 10-5

Absent 6 X

zx x

2

10-4 10-4

From (176); Mutagen NTG. See Table V for details.

NTG-resistant mutants (due to the selective action of NTG), the muwas tant frequency in the presence of this potent mutagen (1.7X essentially the same as in the parent strain in the abs,ence of mutagen (1.6 X (mutability compensation). The phenomenon was reversible; upon removal of the mutagen from the system, the population tended to revert to the original one (Table VIII), thus again maintaining constant mutability. The NTG-resistant mutants were more resistant to UV, as demonstrated also by Greenberg and his collaboratjors "Compare also R. F. Hill (180)for UV-induced mutations.

NUCLEIC ACIDS AND MUTABILITY

31

(181-184). Since such W resistance is believed to involve a DNA repair mechanism, it is likely that a similar repair mechanism was also responsible here for higher NTG survival and for lower NTG-induced mutability. Another suggestion from the above findings was that, since the spontaneous mutability was also lower for NTG-resistant strains, the injury (such as depurination) caused by the unknown “spontaneous” mutagens may be subject to repair by mechanisms analogous to those involved in repair of UV damage (63, 64). The above example of mutability-compensation in the mutagenresistant strains illustrates a case of complete nullification of mutagenic action. For all practical purposes, in strain 15S/NTG2, NTG ceased to be a mutagen. The results of the studies on the incorporation of 5-bromouracil may also be interpreted as an example of a step towards such compensatory mutability. I n this case, a mutant resistant to 5-bromouracil was found to incorporate less 5-bromouracil in its DNA (186). Since the mutagenic action of this analog depends on its incorporation into DNA (1%23), the decrease in this incorporation is a step toward neutralization of the mutagenic action of 5-bromouracil. Other examples are the thymine-deficient strains as such. The conditions of thymine deficiency are mutagenic (186-188) ; under these conditions, a backmutant to thymine independence (46) 185) is selected for. This mutant is “resistant” to conditions of thymine deficiency, which thus cease to be mutagenic. These two examples are, however, rather special cases; a mechanism that may involve DNA repair (see above) is likely to be more general. All such compensatory mechanisms are analogous to the Le Chatelier principle: If a system a t equilibrium is disturbed, the system will shift in such a way as to minimize the effect of the disturbance. The above considerations suggest that the appearance of a mutator gene should always be selected against. That this is not always the case has been demonstrated in a population study (102) in which a strain of E . coli (Harvard strain) carrying a mutator gene (99, 100) was grown with a strain that is genetically stable. Contrary to expectations, the strain with low mutability was overgrown by the one with high mutability (Table I X ) . This result accounts for the preservation of the highly mutable strain on transfers through nutrient media for many years (99). Several mechanisms could explain this survival of strains with higher mutability (102). The original appearance of the mutator gene might have occurred in a strain that, owing to several genes, had better survival value than all the competing strains. The mutator gene might be (or might be linked to) a gene that confers a considerable selective advantage. I n other cases, such as in media con-

32

STEPHEN ZAMENHOF

TABLE IX POPULATION STUDY OF STHAINS WITH AND WITHOUT

MUTATORGENE^

No. of transfers (every 24 hours)

___ Proportion (%) of cells of Harvard strain (with mutator) in a mixture of Harvard and of B (without mutator) a

Penicillin (units/ml)

0

1

2

3

0 20

16 52

19 -100

28 -

41 -

4 48 -

From (102). For strain with mutator gene (Harvard) see (99).

taining antibiotics, high mutability may facilitate faster appearance and establishment of resistant mutants. Once established, the highly mutable strains may provide more possibilities for further evolutionary changes than the less mutable ones. It is not unlikely that periods of accelerated evolution have in the past coincided with the occurrence and persistence of such highly mutable strains.

V. Summary and Conclusions “Spontaneous” and induced mutations in general and their quantitative aspects (mutability) are subjects of great importance. They are also subjects of great complexity, beginning even with their definitions. Many terms need clarification; the borderline between these and other phenomena (such as recombination) is not always sharply defined. Especially for consideration of quantitative aspects, it is important to recognize that in most cases mutation is not a single process but consists of several steps (primary injury to nucleic acids; translation of this injury into a final change in base sequence, with a possibility of repair in between). What one hopes to observe is the end result. Since mutations primarily affect nucleic acids, one can propose their detection by direct detection of changes in nucleic acids, regardless of whether these changes later result in detectable changes in phenotype in general and in proteins in particular. This approach appears sound, but thus far is experimentally feasible only in a very limited number of cases. The easiest way to detect mutations is still by the change in phenotype produced. This, however, often results in a great distortion of both qualitative and quantitative aspects of mutations that originally occurred. Bearing in mind all the above limitations, one can venture to study the mechanisms of inheritance of mutability; in particular, one is in-

NUCLEIC ACIDS AND MUTABILITY

33

terested in estimating whether and how nucleic acids determine the spectrum of mutabilities detectable along a molecule of nucleic acid. Such mechanisms may be manifold. Special DNA segments, called “mutator genes,” can increase the mutability of some or all other genes. This phenoiucnon, however, is usually an exception. As a rule, normal segments of DNA ought to be able to determine their own mutability. Several mechanisms that could account for the latter have been proposed but no evidence to date seems to explain the full range of spectra of detectable mutabilities. It is likely that the functional and structural requirements of protein moieties that are implicated in the detection of mutations contribute considerably to these spectra. A possibility also exists that the state of metabolic activity influences the possible protection of DNA and, therefore, mutability. Mutability itself is subject to natural selection, and several mechanisms have been suggested for the maintenance of mutabilities a t constant levels that are usually (but not always) low. The degree of participation of each of these mechanisms in the overall mutability spectra cannot be estimated a t present. Numerous obstacles exist on the path of elucidation, but I feel that this should not call for postponement of formulation of the problem. ACKNOWLEDGMENTS I am indebted to Dr. Emil L. Smith and to Dr. Patrice J. Zamenhof for critical reading of the manuscript.

REFERENCES 1. Th. Dobzhansky, “Genetics and the Origin of Species.” Columbia Univ. Press, New York, 1951. 8. T. Browne, “Pseudodoxia Epidemica: or Enquiries into Very Many Presumed Truths,” 4th ed., p. 403. Printed by A. Miller for Edward Dod and Nathan Ekins, at the Gunne in Ivie Lane, London, 1658. 3. M. Demerec, Symp. SOC. Exptl. Biol. 7 , 43 (1953). 4. S. Zamenhof, R. De Giovanni-Donnelly, and L. H. Heldenmuth, Proc. Nall. Acad. Sci. U.S. 48, 944 (1962). 6. W. J. Schull, ed., “Mutations.” Univ. of Michigan Press, Ann Arbor, Michigan, 1962. 6. S.Zamenhof, A m . J. Med. 34, 609 (1963). 7. E. Mayr, Federation Proc. 23, 1231 (1964). 8. E. Freese and A. Yoshida, in “Evolving Genes and Proteins” (V. Bryson and H. J. Vogel, eds.), p. 341. Academic Press, New York, 1965. 9. V. Bryson and H. J. Vogel, eds., “Evolving Genes and Proteins.” Academic Press, New York, 1965. 10. A, W. Ravin, A m . Naturalist 97, 307 (1963). 11. F. Lanni, Perspectives Biol. M e d . 3, 418 (1960). 18. E. Freese, J. Theoret. Biol. 3, 82 (1962).

34

STEPHEN ZAMENHOF

13. J. Marmur, S. Falkow, and M. Mandel, Ann. Rev. Microbiol. 17, 329 (1963). 14. D. Dubnau, I. Smith, P. Morell, and J. Marmur, Proc. Natl. Acad. Sci. US.54, 491 (1965). 16. E. L. Smith and E. Margoliash, Federation Proc. 23, 1243 (19Sa). 16. E. Margoliash and E. L. Smith, in “Evolving Genes and Proteins” (V. Bryson and H. J. Vogel, eds.), p. 221. Academic Press, New York, 1965.

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Specificity in the Structure of Transfer RNA KIN-ICHIRO MIURA Institute of Molecular Biology, Faculty of Science, Nagoya University, Chikusa, Nagoya, Japan

I. Introduction . . . . . . . . . . . . . . 11. Base Composition of tRNA . . . . . . . . . . 111. The Arrangement of Nucleotides in tRNA . . . . . . A. Nucleotide Distribution . . . . . . . . . . B. Anticodon Sequence . . . . . . . . . . . C. The Terminal Nucleotide Sequences. . . . . . . IV. The Three-Dimensional Structure of Transfer RNA . . . V. Studies on the Functional Sites in tRNA . . . . . . A. Modification of Specific Sites. . . . . . . . . B. Inhibition by Oligonucleotides of Aminoacylation of tRNA References . . . . . . . . . . . . . . .

. . . . . .

. .

. . . . . . .

. . . .

.

.

.

39 40 43 43 48 52 59 63 63 71 76

1. Introduction Recent progress in molecular biology has shed much light on the mechanism of protein synthesis and the general principles involved have been established by the efforts of a number of workers, using mainly cellfree systems from bacteria. The genetic code, carried in the nucleotide sequence of DNA, is transcribed into messenger RNA which, in turn, associates with several ribosome particles to form the polysomes, in which mRNA is able to perform its role as a template. Transfer RNA’s, which are aminoacylated by the aid of aminoacyl-RNA synthetases, carry amino acids to the proper position on templates. In each cell there are many tRNA’s, each specific for an individual amino acid. A “codon” (a triplet of nucleotides) in the template selects the appropriate tRNA not because of the amino acid itself, but because of the nucleotide sequence of the “coding site” in the tRNA molecule. The concept that an individual tRNA molecule can adapt to a specific code in mRNA was originally advanced in the “adaptor” theory by Crick ( I ) , and was clearly established experimentally by Chapeville e t al., ( 2 ) and Weisblum et al. (3). There are multiple tRNA molecules even for a single 39

40

KIN-ICHIRO MIURA

amino acid, and these may be separated by many fractionation methods (4). Each functions preferentially with a specific coding template (3, 5, 6 ) ; for example, one of the leucine tRNA’s of E. coli incorporates leucine into polypeptides using poly (U,C), but another leucine tRNA functions only with poly (U,G), and not with poly (U,C). The specific interaction between tRNA and mRNA is considered to be a complementary base pairing of G with C and A with U as in the DNA molecule, though there is no direct evidence for this. On the other hand, the specific interaction between a tRNA and a synthetase, which is also specific for each amino acid, would occur between the nucleotide sequence of the “synthetaserecognizing site” in tRNA and probably the peptide sequence in the functional site of the synthetase. I n these two steps, the tRNA translates codewords for amino acids in different ways. Most of the coding triplets of mRNA have been established by Nirenberg, Khorana, Ochoa, and their co-workers (7-9).However, the translation mechanism, in which tRNA plays a central role, is still an unsolved problem. In approaching this problem it is obviously important to know the structural specificity of transfer RNA. Many workers have already contributed to our knowledge of this topic and several review articles on tRNA studies have been published, for example by Berg ( l o ) , Brown (4, 11), and Hoagland (12). The present article is concerned with recent work on the specificity embodied in the structure of transfer RNA.

II. Base Composition of tRNA The numerous analyses of the base composition of bulk tRNA preparations from different sources have been reviewed by Allen (IS) and Brown (4). These results, and others not cited in these reviews (4, 13) show a similarity of base composition throughout the range of organisms tested: yeast, bacteria, protozoa (Tetrahymena),algae (Euglena),wheat germ (14, 15),pea (16), blowfly, silkworm (17),fish, and mammals. I n many bacterial species, deviations in the base compositions of bulk tRNA are small, though the compositions of their DNA’s show remarkable variety in G plus C content (18,19). A few representative analytical results are shown in Table I. Countercurrent distribution, column chromatography, and other separation techniques have been used in the fractionation of tRNA’s. Some of them are highly specific for one particular amino acid. The base composition of these fractionated tRNA’s is characteristic (20-25). Since Davis and Allen (26) in 1957 discovered the fifth nucleotide component, identified later by Cohn (27) as pseudouridine, in the soluble RNA fraction in addition to the four major nucleotides, many

m w

H

2 ;?

2* 5 Y

!2 u,

NUCLEO’PIDE COMPOSI’PION O F

tRNA’s

FROM v.4RIOUS

TABLE I SOURCES (MOLE %

e s

OF I D E N T I F I E D

NUCLEOTIDES A N D NUCLEOSIDES)

s

3

0

Source Yeast (63) E. coli (30) Wheat germ (14) Silk worm ( 1 7 ) Rabbit liver (57)

G

A

C

U

*

29.2 32.1 29.7 38.3 31.1

21.1 20.3 21.8 19.3 16.6

26.4 28.9 23.6 26.1 27.8

16.1 15.0 19.2 19.3 15.9

4.6 2.1 2.6 3.3 4.3

T

MG

MA

JIC

pGp

1.3 1.1 1.3

0.1 0.5 2.4

0.4 -

-

-

0.4

0.6

-

1.2 1.9 1.5

-

-

-

Nucleosides

4

1.3 -

$

1.3 1.1

B

z

q

g P

42

RIN-ICHIRO MIURA

minor nucleotide components have been found in tRNA (see articlcs by Goldwasser and Heinrikson, Chambers, and Srinivasan and Borek in Volume 5 of this series). Of these odd components in RNA, pseudouridylic acid has been detected in every tRNA analyzed. Its content in yeast tRNA is between 2 and 5747, that is, 2 or 3 moles per tRNA molecule on the average, and it is tlie most abundant of the minor components. In bacterial tRNA, the content of pseudouridine seems to be lower than in yeast tRNA (4, 18). About twenty different minor comTABLE I1 FROM YEASTtRNA (28) MINORNUCLEOSIDES ISOLATED Nucleoside

Mole %

Ne-Methyladenosine 2‘(3’)-0-Ribosyladenosine N6-(Aminoacyl)adenosines l-Methylguanosine N4-Methylguanosine NZ,N2-Dimethylguanosine 3-Methylcytidine 5-Methylcytidine 3-Methyluridine 5-Methyluridine (ribosylthymine) Pseudouridine Inosine 1-Methylinosine 2’-0-Methyladenosine 2’-0-Methyluridine 2’-0-Methylguanosine 2’-O-Methylcytidine 2’-O-Methylpseudouridine

0.39 0.013 0.034 0.090 0.014 0.236 0.018 0.084 0.01 0.69 4.5 0.22 0.04 0.028 0.029 0.31 0.11 0.0009

ponents have been identified by Dunn, Smith, Littlefield, and by Hall ( 2 8 ) . Many of these contain methylated bases, which are particularly abundant in soluble RNA as compared with other RNA’s of high molecular weight (29, 30). Some of the minor components are the 2’-O-methyl derivatives of the usual nucleosides (31, 3 2 ) . Hall (28) confirmed the presence of eighteen minor nucleotides in yeast tRNA as shown in Table 11. Recently, 4-thiouridine was discovered in E . coli tRNA (33, 34) while Madison and Holley (36) found dihydrouridine in yeast tRNA. Dihydrouridine does not show the ultraviolet absorption in the 250-280 mp region that is typical of the usual base components. If the average chain length of tRNA molecules is taken as seventyseven nucleotides, 1.3 mole % nucleotide corresponds to 1 mole in one

SPECIFICITY IN THE STRUCTURE OF TRANSFER RNA

43

tRNA molecule. Since many minor components are less plentiful than 1.3 mole %, respectively, not every minor component is contained in each tRNA molecule. It is still uncertain whether these minor nucleotides are necessary for the function of tRNA. It is known that methyldeficient tRNA from E . coli (i.e., cultured under conditions of methionine starvation) accepts amino acids to the same extent as does normally methylated tRNA ( 3 6 5 9 ) . However, methyl-deficient E . coli tRNA does not accept leucine with the yeast aminoacyl RNA synthetase as does normally methylated E . coli tRNA (40).

111. The Arrangement of Nucleotides in tRNA A. Nucleotide Distribution It is necessary to cleave the polynucleotide chain a t definite sites in tRNA in order to establish the nucleotide arrangement. Fortunately, two kinds of specific ribonucleases are available for this purpose. Pancreatic ribonuclease IA (abbreviated as RNase I in this article) is highly specific for pyrimidine nucleotides and cleaves the bond between the 3‘-phosphate of a pyrimidine nucleotide and the 5’ site of the adjacent nucleotide in RNA (41). RNase TI, discovered by Sat0 and Egami (42) in Taka-diastase, splits the bond between the 3’-phosphate of guanylic acid and the 5’ site of the adjacent nucleotide in RNA (43) (see article by Egami et al. in Volume 3 of this series). When these enzymes were used for RNA digestion, their specificities were confirmed for both individual and combined use (44). The specificity of the ribonucleases for the unusual nucleotide components found in tRNA (43) is also a matter for discussion. Although many workers have explored other specific splitting methods, including modification of the RNases or of the substrate tRNA, no satisfactory procedures have yet been obtained. The oligonucleotides produced from RNA by the ribonucleases can be separated by column or paper chromatography. Among the various methods employed, chromatography on DEAE-cellulose columns using concentrated urea in the eluting system, as developed by Tomlinson and Tener (45, 5 8 ) , and the two-dimensional mapping procedure involving electrophoresis and chromatography on filter paper, as developed by Rushizky and Knight (467, have been exceedingly useful. Quantitative analysis of the oligonucleotides obtained by ribonuclease digestion from unfractionated tRNA show that the nucleotide sequences of tRNA are nonrandom, though the amounts of mononucleotides in the digest do not deviate significantly from the value calculated on the basis of a random distribution (25, 44, 47-49). Analysis of the oligonucleotides in the RNase I plus RNase TI digest of unfractionated tRNA

44

KIN-ICHIRO MIURA

(Table 111) shows that the amount of every ApNp is more than 2.3 mole (48). This means that, on the average, each kind of tRNA molecule may have a t least one sequence of every possible ApNp, since the average chain length of a tRNA molecule is about seventy-seven nucleotides, and one place in a tRNA molecule corresponds to 1/77 (1.3%) of the total nucleotides. The amount of every ApApNp is between 1 and 0.1% of total nucleotides; some molecules, therefore, may not have every possible sequence of ApApNp. No sequences of more than four adenosine residues were found in yeast tRNA (48, 25). Sanger e t al. (51) showed that the amounts of the isomers,

% of the total nucleotides

TABLE I11 OLIGONUCLEOTIDES IN THE RNASEI AND RNASE TI DIGESTOF YEASTtRNA (MOLARPROPORTIONS OF THE TOTAL NUCLEOTIDES) (48) ~

Amount (%) Nucleotide

Observed Calculated as random dietribution 20.9 20.4 12.5 2.7

21.5 20.5 13.9 3.5

4.5 2.3 4.2

3.7 3.6 2.4

0.8 0.3 0.6

0.7 0.7 0.4

even among trinucleotides, are not equal in certain cases; for example, the ratio of CUG/UCG or of CAG/ACG is about 2 in E . coli tRNA. Aronson (52) analyzed the amount of oligonucleotides in the RNase I digest of tRNA's from various bacterial species with widely varying G plus C contents in their DNA, and found considerable differences in their nucleotide distribution, although their base compositions were quite similar to each other (see Section 11).Miura and Matsuzaki (17) analyzed the nucleotide arrangement in tRNA from the posterior silk glands of two kinds of silkworm. In the posterior silk gland of Bombyz mmi a glycine-rich fibroin (46% glycine, 29% alanine) is synthesized, whereas in that of Philosamiu cynthiu ricini an alanine-rich fibroin (48% alanine, 32% glycine) is produced. The amounts of oligonucleotides in the RNase I and RNase T, combined digests of the two kinds of tRNA are similar to each other, although they differ slightly from yeast

SPECIFICITY IN THE STRUCTURE OF TRANSFER RNA

45

tRNA. Sanger et al. (51) compared E. coli t R N h with yeast tRNA by mapping P3,-1abeled oligonucleotides in the RNase TIdigest of tRNA’s from the two sources. Both tRNA’s showed many similarities in their nucleotide distribution, but many obvious differences were also detected. The sequence GCUCAG, for instance, seemed to be present in more than half the different tRNA’s of E . coli, although i t was not so common in yeast tRNA’s. These findings suggest strongly the presence of structures common to many different kinds of tRNA in one species. Nihei and Cantoni (64) digested tRNA by an exopolynucleotidase, snake venom phosphodiesterase, and analyzed the composition of the nucleotides released a t 19%, 5376, and 99% hydrolysis. They concluded that the methylated purines and pseudouridine are concentrated in the central portion of the tRNA chain. Miura (48) analyzed the amount of pseudouridine in the RNase I digest and in the RNase I plus RNase TI digest of yeast tRNA. In the former, 48% of the pseudouridine appeared as mononucleotide coming from the sequence ( P Y ) ~I~n . the latter, 78% of the pseudouridine appeared as the mononucleotide, derived from the sequences ( P Y ) ~ S and (G),P. The amount of (G)n@can be shown to be 30% by subtracting the amount of pseudouridine in the first digest from that in the second digest, the difference between total f and the amount in the latter digest (22%) corresponding to the frequency of the sequence (A),@. Hence i t may be concluded that the neighboring position to the 5‘-phosphate of pseudouridylic acid in yeast tRNA is not restricted to any one nucleotide. Staehelin (56) showed that the distribution of the rare nucleosides in tRNA is not random, that they cluster in several sequences in the RNase I digest of unfractionated yeast tRNA, as in 2Me,G-@ and 1MeG-2MeG-C. Since the amounts of these two sequences are about 0.3 residue per chain, they cannot be present in each tRNA molecule, but it is suggested that more than one individual molecular species of tRNA may include them. Zamir and coworkers (66) found the sequence T f C G in the RNase TI digest of unfractionated tRNA’s from yeast, E . coli, and rat liver in the amount of one residue per chain. Sanger et al. (61) confirmed this result by a different method of analysis. As the amount of T in tRNA is the same, all T may be included in this one sequence. The tRNA’s specific for alanine, valine, and tyrosine contain this sequence (56). It is strongly suggested that the sequence G T W G occurs in every tRNA molecule and performs some common function, such as binding to ribosomes or to the aminoacyl-tRNA synthetase. Sanger et al. (61) found H,U-A-G in unfractionated tRNA of yeast or E . wli in about one-third the amount of the T W G . It is more common than UAG in these tRNA’s and it may be contained in many tRNA molecules.

46

KIN-ICHIRO MIURA

The nucleotide distribution of partially purified tRNA preparations Lipshita-Wiesner and Chargaff was analyzed by Holley e t d. (%I), ( 2 3 ) , Staehelin et al. ( 5 9 ) , Doctor et al. (GO),Armstrong et al. ( G I ) , and Bergquist and Scott ( 2 5 ) . In all cases, remarkable differences in the amount of particular oligonucleotides in the RNase digest were found among different tRNA fractions. For example, the fraction rich in alanine tRNA contains a high proportion of GU as compared with the tyrosine tRNA fraction (the ratio is about 2: 1) (57, 60) , as shown TABLE IV OLIQONUCLEOTIDES IN THE RNASE I DIGESTSOF tRNA PER MOLE OF tRNA) (60)

0

FROM

YEAST (MOLES

Nucleotidea

Original mixture

Tyrosine tRNA

Alanine tRNA

CP UP *P GPCP APCP APUP GPUP APAPCP (APGP)CP APAPUP (APGP)UP GPGPUP GPGPCP (APGPGPICP (APAPGP)UP (APGPGP)UP

12.1 7.0 1.1 2.3 1.3 1.2 2.3 0.3 0.7 0.7 1.5 1.3 0.9 0.2 0.4 0.6

12.7 7.9 0.9 1.4 0.2 0.2 1.9

15.3 7.1 1.4 1.7 1.1 0.2 4.4 1.1

-

0.8 0.4 1.2 0.3 0.2 0.5 0.1

-

2.7 1.6 0.9 0.5

-

Parentheses enclose nucleotides of unknown sequence.

in Table IV. These results suggest that the structure of tRNA differs among tRNA’s specific for individual amino acids. On fractionation of tRNA, sometimes “degenerate” molecules are separated. Rushizky and co-workers (6s) separated three serine tRNA’s by countercurrent distribution, and Bergquist and Robertson (64) obtained five serine tRNA’s by partition chromatography on Sephadex. They digested the fractions with RNase I and determined the amounts of oligonucleotides. Though many features of nucleotide distribution were common to many of the fractions, obvious differences were found in some sequences, as shown in Table V.

47

SPECIFICITY I N THE STRUCTURE O F TRANSFER RNA

TABLE V OLIGONUCLEOTIDES IN THE RNASE I DIGESTSOF Two SERINE tRNA’s
Serine I1 RNA Serine I11 RNA 10.60 0.60 0.78 3.90 1.30 0.78 1.33 0.52

11.00 0.49 0.81 3.82 1.04 0.88 1.48 0.44 6.S6 1.09 1.30

1.06

1.37 1.25

a

1.67

0.68

2.60 0.26 8.60

2.28 0.25 7.80

The base compositions are the same, i.e.: Cp, 27.7.; Ap, 20.8; q p 3.3; Up, 19.2;

Gp, _ .28.7%. b

Parentheses enclose nucleotides of unknown sequence.

Many workers have concentrated their efforts on the purification of individual tRNA molecules in order to determine the complete nucleotide sequence. The sequence for yeast alanine tRNA was first established by Holley et al. (66) (Fig. 1 ) . After purifying alanine tRNA by countercurrent distribution, they cleaved the RNA molecule into two large fragments by limited digestion with RNase T1 (short treatment a t Me

p ~ - G - G - C -WG-U-G-G-C-G-c G-

DiH

Din

D‘Me

-G-u A-G-Gc-G-G-b-A-G-C-G-C-d.

9

c~w-’-E)-w’8-o-’-E)-~-!-~-’-E)-i-n-n-3-3 -G t ekv %c-C-GG-T-Y-C-G-A-U-U-C-C-G-

G-A-C-UCG-U-C-C-A-C-C-A~

FIG.1. Whole nucleotide sequence of an alanine tRNA from yeast

(65).

DiH = dihydro-; DiMe = dimethyl; Me = methyl.

OOC). Both the fragments were degraded further by RNase digestion, and reconstruction of the whole sequence was achieved by isolating the overlapping sequences. Many minor nucleotides in this tRNA were useful markers for sequence determination, but they were not concentrated in the middle part of the chain. McCully and Cantoni (66) and Spencer e t al. (67) had earlier considered the presence of long runs of comple-

48

KIN-ICHIRO MIURA

mentary sequences connected through a hairpinlike structure formed by a folding of the chain back upon itself, thus creating a loop in the middle of the molecule where minor components might be abundant ( 5 4 ) .This, however, is not the case (see Section IV). Alanine tRNA may have a complex structure with many coinplementary regions allowing hydrogen bonding of A to U and of G to C and also some “looped-out” sequences. Precise surveys on the overlapping of oligonucleotide sequences in the RNase digests for serine tRNA by Zachau e t al. (68) and by Cantoni e t al. (69) are now in progress. Much attention is also being paid to the oligonucleotide sequences in the RNase digests of other specific tRNA’s. Although some of these results have been mentioned above, other characteristics of the structure of these specific tRNA’s, especially of the anticodon structure and the terminal sequences, are described in the following sections.

B. Anticodon Sequence An adaptor sequence in tRNA is considered to be an (‘anticodon,” a sequence complementary to the codon, or code triplet, in messenger RNA in terms of the conventional base pairing of A to U and G to C. If this is so, phenylalanine tRNA, for example, would have AAA as an anticodon, because phenylalanine tRNA attaches to the ribosomepoly U complex specifically (62) and phenylalanine is polymerized in a cellfree system catalyzing synthesis in the presence of poly U (70, 7 1 ) . Adenylic acid clusters can be detected in the type (A),N ( N # A) when RNA is digested by the simultaneous use of RNase I and RNase TI. In our laboratory, yeast tRNA was digested by the combined action of RNase I and RNase TI and the resulting nucleotides were separated by the two-dimensional mapping procedure of Rushizky and Knight (46). The oligonucleotides in the digest were acid-soluble, and every nucleotide spot observed was smaller than a tetranucleotide. Sequences of more than four adenylic acid residues, if any, are less than 0.1% of the total nucleotides. Only one faint spot of the oligonucleotide supposed to contain three adenylic acid residues was detected. It is assumed from the position of this spot that the neighboring nucleotide to the AAA triplet (i.e., the N in AAAN) would be some kind of uridylic acid. The amount of the spot presumed to be AAAN was approximately 0.05% of total nucleotides. Therefore, only a small fraction of the total tRNA’s may have such a sequence. The probability of the occurrence of one characteristic sequence in a given tRNA molecule is 1/77 x 1/40, or 0.03%, on the supposition that the average chain length of tRNA is about seventy-seven nucleotides, that the tRNA mixture used in the

SPECIFICITY I N THE STRUCTURE OF TRANSFER RNA

49

experiment contains equal amounts of twenty kinds of tRNA’s, and that each variety of tRNA consists of two kinds of molecules to cover the degeneracy of the code. Very small amounts of AAAU, AAAG, AAAC, and an unidentified tetranucleotide containing are detected in the digest of a large amount of yeast tRNA by Dowex-I chromatography ( 7 g ) . The presence of AAAU, AAAG, and AAAC in tRNA has also been reported in a few papers (26,4 7 ) . Phenylalanine tRNA was separated from other tRNA’s in an effort to establish the presence of t,he AAA anticodon. According to Miyanaki et al. (7.31, the tRNA’s specific for individual amino acids may be successfully fractionated by column chromatography on DEAE-Sephadex with appropriate variations of the eluting solvents. By the procedures shown in Fig. 2, phenylalanine tRNA was concentrated about 20-fold with respect to its aminoacyl acceptance per optical density unit. When the fraction rich in phenylalanine tRNA was digested with RNase I and RNase TI, and the resulting nucleotides were mapped, the results (74) in Table VI were obtained. The amounts of some of the oligonucleotides differed from those found in digests of unfractionated tRNA. Two tetranucleotides appeared in this map. One of them could correspond to the AAAU analog, which was observed as a faint tetranucleotide spot in the map of the digest of unfractionated tRNA. Another had not been observed in the map of the digest of unfractionated tRNA. These two spots were pooled from several maps, eluted with N/100 HCI, and dried in vamo. These samples were digested with RNase T, and their nucleotide composition determined by paper chromatography. It was concluded that one is AAAU and the other (A,A,MeI)U*;U* is a U-analogue. These two oligonucleotide sequences thus appear to be characteristic of phenylalanine tRNA. The second spot was also found in the digest of phenylalanine tRNA, which was partially purified (84) on a methylated serum albumin column ( 7 5 ) , followed by concentrating by Zamecnik’s dye method ( 7 6 ) . I n the sequence of yeast alanine tRNA, as determined by Holley et al. ( 6 5 ) , one can find a few anticodon sequences, e.g., CGG, GGC, and IGC. Cantoni et al. (69) detected a few anticodon sequences (A,G,A) in a purified serine tRNA. One of them was neighbored by pseudouridylic acid as PyAGAik. An analysis of serine tRNA by Dutting et al. (68) shows similar sequences, as AAGAq. I n addition, they observed IGA as well as a few (G,A,G) sequences (68). I n the pancreatic RNase digest of valine tRNA, GAC and IAC were detected by Ingram et al. (61) and Bayev et al. ( 7 7 ) .These anticodon sequences, detected in specific tRNA’s, are listed in Table VII.

*

50

KIN-ICHIRO MIURA

An anticodon sequence enclosed by two dihydrouridylate residues in alanine tRNA suggested that this might be a functional site, distinguished by neighboring odd bases from nonfunctional parts. However, the sequence H,U-2’0MeG-G-H2U has been found in serine tRNA

I

I

1 A,,

CPM/Tube

DMFA-(NH,l, SO,

CI

t,.i

x I@

5-

2.0

0.8 M KCI

3l-

20

40

I

00

60

100

I

.Ll

I

I L

20

40

60

140

00

5.0

Borate - KCI

2.0 1.0

20

40

60

80

100

I20

Tube No.

FIG. 2. Fractionation of phenylalanine tRNA by DEAE-Sephadex (73). (a) Gradient elution by (NH,)&O, from 0.75 to 1.6 M in 0.02 M potassium acetate buffer (pH 5.3) containing 1% (V/V) dimethyl formamide (DMFA). (b) Gradient elution by KCl from 035 to 0.40 M in 1 M potassium phosphate (pH 6.2) containing 5% (V/V) dimethyl formamide. (c) Gradient elution by KC1 from 0.60 to 0.70 M in sodium borate (pH 7.0).

by Diitting et al. (68). This doublet bracketed by two dihydrouridylate residues does not correspond to the anticodon triplet. I n phenylalanine tRNA, the anticodon sequence AAA is not adjacent to dihydrouridylate. Consequently H2U-CGG-H,U in alanine tRNA might be an anticodon sequence by chance. I-G-C-MeI-4 in alanine tRNA seems to be another possibility, since similar sequences have been detected in phenyl-

51

SPECIFICITY IN THE STRUCTURE OF TRANSFER RNA

TABLE \'I OLIGONUCLEOTIDES IN THE RNASE I AND RNASE TI DIGESTOF PHENYLALANINE tRNA-RrcH FRACTION FROM YEAST(MOLES OF THE TOTAL NUCLEOTIDES) (74) Nucleotide

Amount (%)

A

1.3

GP CP UP

19.0 21.2 14.0

TP *P IP

1.2 2.8 0.9

PGP

I .R

Nucleotide ApGp APCP APUP

+ ApPvIeGp

Amount (%) 7.4 2.4 4.1

0.6 0.4 1.8

APAPAPUP (ApApMeIp)U*.p

0.4 0.4

IJridine analogue

alanine, serine, and valine tRNA's as listed in the right column of Table VII. It is significant that the rare components I, Me1 and ?P are prominent in these sequences. I could be read as G or A in degenerate codes. If we take YNN'Z.k as a generalized form for these sequences, the three nucleotides on the left could correspond to the anticodon sequence and TABLE VII IN tRNA ANTICODON SEQUENCES Amino acid

Anticodon sequences"

Ser (68)

The sequence of the nucleotides in parentheses has not been determined. N(6) Isopentenyladenosine. c Uridine analogue. (1

b

52

KIN-ICHIRO MIURA

the two rare nucleotides on the right could be markers indicating a functional site. The first residue Y represents I in alanine tRNA or serine tRNA, and it might be considered as a degenerate code indicator. In serine tRNA, the position Z, corresponding to Me1 in alanine and in phenylalanine tRNA, is given as N(6) isopentenyl-A by Diitting et al. (77a, 7 7 b ) . As the amounts of I and Me1 in the unfractionated yeast tRNA do not amount to 1 mole per chain on the average (28, 78) (see Table 11),Y and Z might be different bases in different tRNA’s. Much more detailed determination of nucleotide sequences of specific tRNA’s is required to give more clear-cut generalized information.

C. The Terminal Nucleotide Sequences When tRNA is digested with alkali, most nucleosides appear as 2’- and 3’-mononucleotides, with small amounts of nucleoside 3’3’-

bisphosphate and nucleoside. I n the alkaline digest small amounts of oligonucleotides containing 2’-0-methyl ribose are also found (31, 32), which are resistant to alkaline digestion. The alkali-resistant dinucleotides, however, are not only those containing 2’-0-methyl ribose, as some “normal” dinucleotides have been shown to be resistant under certain digestion conditions (79). The nucleoside bisphosphate and nucleoside, which comprise 1-1.5 mole % of the total nucleotides, originate in the two terminals of a tRNA molecule. Although the major part of the bisphosphate found is pGp, others were found by Ralph .et al. (SO),and by Bell et al. (SI), as seen in Table VIII. It has not been determined whether the 5’-phosphate terminal nucleotide of the intact tRNA is always guanylic acid. Ralph and co-workers (80, 82) tried to label the 5’-terminal phosphate of a tRNA molecule with C14-aniline or C14-methylphosphoromorpholidate. Progressive pancreatic ribonuclease digestion yielded a labeled terminal oligonucleotide as C14-aniline-pGp (Pup).Pyp. Separation was achieved by the aid of DEAE-cellulose, and a considerable proportion of pGpCp was identified as the 5’-terminal dinucleotide sequence. Since the oligonucleotide pGp (Pup),Pyp is resistant to spleen phosphodiesterase because of its 5’-terminal phosphate, when tRNA is digested with pancreatic ribonuclease and spleen phosphodiesterase , the internal oligonucleotides are digested to 3’-mononucleotides, and the terminal oligonucleotide remains intact. When the resulting nucleotides were separated by mapping procedures, the dinucleotide region was found to contain much pGpCp but only small amounts of pGpUp (83). This observation agrees with the results of other workers (84, 8 5 ) . The results of quantitative analyses for terminal mono- and dinucleotides (81) are listed in Table VIII. McLaughlin and Ingram (86) found

53

SPECIFICITY IN THE STRUCTURE OF TRANSFER RNA

no pGpGpCp in the trinucleotides of the pancreatic ribonuclease digest of mixed tRNA’s. They concluded that the frequencies of the oligonucleotides in unfractionated tRNA are apparently not random. A few examples of the terminal nucleotide sequences of specific tRNA’s are listed in Table IX. The other terminal of a tRNA molecule is considered to be the common sequence CpCpA (86-90). Amino acids become attached to the TABLE VIII 5’ TERMINAL SEQUENCES OF YEASTtRNA (81)

Terminal mononucleotides in alkaline digest Amount (%)”

Terminal nucleotides in RNase digestb

Amount ( % ) b

-

PGP PUP PAP PCP

78 10 7 5

PUP PCP

11.5 6.5

PGPCP PGPUP PAPUP PAPCP

26 0.4 0.7 0.1

% of nucleotide diphosphates derived from the 5’ terminal of tRNA. % of tRNA molecules.

TABLE I X TERMINAL SEQUENCES OF SPECIFICtRNA’sO Amino acid Ser (68) Ala (66) Val (61,77)

5‘ terminal sequences in RNase I digest PGPGPCPPGPGPGPCPPGPGPUP-

3’ terminal sequences in RNase TIdigest -Gp C pCp A -GpUpCpCpApCpCpA -(APAP,AP CP,UP)CPCPA 5

Sequences in parentheses are unknown.

3’-hydroxyl of this terminal adenosine residue, although it has also been claimed that some amino acid residues also become attached to the 2’ position (50, 91-9s). A final decision on this point has not yet been made (see article by Zachau in Volume 4 of this series). Equilibration between the 2’ and 3’ isomers of aminoacyl-tRNA may be sufficiently rapid to supply whichever one is required by the enzymes involved in peptide bond formation (50).In any case, the terminal sequence CpCpA seems to be used by all amino acids and for a large variety of biological species. Some tRNA preparations do not contain the terminal pA or

54

KIN-ICHIRO MIURA

pCpA sequence but this deficiency can be remedied by incubation with ATP and CTP in the presence of the appropriate enzyme (94). The sequence CpCpA was chemically proved by Neu and Heppel using a method that strips bases one by one from the terminal sequence (96). Regarding the sequences attached to this terminal CpCpA, Lagerkvist and Berg (98) and Herbert and Wilson (97) showed that the fourth nucleotide from the 3' CCA terminal of tRNA from E. coli, yeast, and rat liver is heterogeneous. Berg et al. (98) determined the sequences adjoining the CpCpA by the principle shown in Fig. 3. A specific amino

I . . P x ~ Y ~ z ~ c ~ leucine c ~ A -1 . .pYpZpXpCpCpA . .pXpZpYpCpCpA Periodate amine -catalyzed hydrolysis,alkali

I

PXPYPZ

I

. . PYPZPXPCPCP . . PXPZPYPCPCP

1

I

PPi

I

. .PYPZPXPCPCP . . PXPZPYPCPCP

CTP-a'p

t

RNase

FIG.3. A method for labeling the two terminal Cp residues of a definite tRNA (98).

acid (in this case, leucine) is attached to the tRNA. The ribose of the terminal adenosine residues of tRNA not aminoacylated is deleted by periodate and amine treatment, leaving all tRNA's except leucyl-tRNA with a primary phosphate a t the 3' terminus. When leucine is removed a t an alkaline pH, the terminal adenosine of the leucine tRNA is revealed. On pyrophosphorolysis, pA and pC are released from the leucine tRNA, but all other tRNA's are unchanged. When P3WTP is incorporated into the leucine tRNA molecule, the terminal p*Cp+C sequence is formed. By such treatment, only leucine tRNA is labeled. When the treated tRNA mixture is digested with RNase T, and the resulting oligonucleotides are separated by DEAE-cellulose, the structure of the labeled oligonucleotide may be studied. In this way it has been

55

SPECIFICITY I N THE STRUCTURE O F TRANSFER RNA

shown that one terminal sequence of E . d i isoleucine tRNA is GC(U,C)ACCA, and that the two terminal sequences GCACCA and GUACCA occur. Bergquist (99) showed by this method that the terminal sequence of serine tRNA from yeast is G(A,C)CCA. Heterogeneity in tlie nuclcotide sequence ncar the aminoacyl-accepting terminal of tRNA was further studied independently in a different way by Ishida and Miura (100, 101) and by Herbert et al. (102, 103). A specific amino acid labeled with C14 or H' was put on tRNA by aminoacyl RNA synthetase. The aminoacyl-tRNA was digested with RNase

BOO

-LY8

600

-

.

-200

-m400

-

200

-

-

-

- i - 100 -100 -

0-

G O -0 0

1 0

20

30

50 60 Fraction number 40

70

80

90

_ _ _ - .. . . . --.,FIQ. 4. Elution pattern of HNase TI digests of L C J lysine-charged, 1G"J phenylalanine-charged, [C^1 valine-charged, and [CI41 leucine-charged tRNA's of bakers yeast on DEAE (100,101). Four independent experiments are superimposed. _I ~ .._

TI and the resulting oligonucleotides were separated on a DEAE-cellulose column. The terminal oligonucleotide originally adjacent to the guanylic acid residue nearest the terminal in the chain was detected as a peak containing radioactive material ; thus: NpGp(Np),CpCpA-amino

1RNaae TI

(Np),Gp

acid-Cl4 (or

+ (Np),CpCpA-amino

H3)

acid-Cl4 (or

H3)

where N = nucleosides other than guanosine and n = 0, 1, 2, 3, . . . . I n yeast tRNA, only one peak was observed for lysine and one for phenylalanine, but two peaks were found for valine and for leucine as shown in Fig. 4. For yeast phenylalanine, however, Bergquist (99) 'resolved two terminal sequences by the same method, and for yeast valine tRNA, similar results have been reported by two other laboratories

56

KIN-ICHIRO MIURA

(lO.2-lO4). Thiebe and Zachau (105) showed the presence of two kinds of valine tRNA from yeast after fractionation by countercurrent distribution or on a methylated albumin column. The two different terminal sequences indicate the degeneracy among these tRNA's. This was confirmed by Grachev et al. (104) using the finding that one of the valine tRNA's fractionated by countercurrent distribution is sensitive to hydroxylamine modification in regard to its ability to accept amino acids. One peak of terminal oligonucleotides of valine tRNA's disappeared on hydroxylamine treatment prior to amino acid attachment. Two valine tRNA's may also be separated from yeast by DEAESephadex column chromatography (106). Kawata and Takemura (107)

I

1

Fraction No.

FIQ.5. Chromatography of a mixture containing RNase TI digests of yeast [H31 valyl-tRNA (open circles) and rat [C"l valyl-tRNA (closed circles) ( 1 0 1 ) .

in our laboratory showed that one of them contained one terminal oligonucleotide sequence whereas the other yielded the other terminal oligonucleotide sequence on column chromatography on DEAE-Sephadex. Thus, degenerate valine tRNA molecules differ from each other a t least in their nucleotide sequence near the CpCpA terminal. The fractionation of tRNA by countercurrent distribution or by methylated albumin column chromatography suggests that the same specific tRNA's from different organisms differ from each other in structure (108, 109). Ishida and Miura (101) compared the sequences near the CCA terminal of the same specific tRNA's from different organisms. As shown in Fig. 5, valine tRNA's from the rat display two different kinds of the terminal sequence. These two sequences are also distinct

57

SPECIFICITY I N T H E STRUCTURE OF TRANSFER RNA

from the two terminal sequences of the yeast valine tRNA’s. The variety in these terminal sequences of the same specific tRNA’s from different organisms is shown in Figs. 6 and 7 for leucine and alanine tRNA’s. Whereas the nucleotide sequences near the CCA terminal of alanine tRNA from yeast, E . coli, rat, and silkworm differ from each other, those of bakers’ yeast, Saccharomyces cerevisiae, and torula yeast, Torulopsis utilis, are similar (110). The nucleotide sequences in the area near the CCA terminal are species-specific, though they may be identical in closely related organisms. I n spite of the fact that the valine tRNA’s of the rat and yeast differ from each other in their terminal nucleotide sequences, the aminoI

I

I

I

I

I

I

I

I

Fraction No.

FIG.6. Chromatography of RNase T, digest of a mixture containing rat [H’I leucyl-tRNA and [ C 4 1 leucyl-tRNA’s of yeast and E. coli (101).

acyl-RNA synthetase of the rat and yeast may replace each other. For leucine or alanine incorporation also, similar exchangeability of the synthetases and the tRNA’s among different organisms is found, although the terminal structures in the tRNA’s may again be different. It is therefore suggested that the synthetase-recognizing site in tRNA is not located near the aminoacyl-accepting terminal. Marcker (111) showed that the nucleotide sequences near the CpCpA terminals of methionine tRNA and N-formylmethionine tRNA differ from each other. N-Formylmethionine is now considered to be a possible chain initiator in peptide formation (112, 116). According to Marcker and Sanger (113)’ formylation of methionine occurs after the attachment of methionine to tRNA, but all the methionine in the methionine tRNA’s is not formylated. The methionine tRNA capable of formylation for methionine transport and the corresponding methionine tRNA that can

58

KIN-ICHIRO MIURA

i

i

600 c'4 1200 H3

o

C'4-Ala Torula Yeast H3-Ala Bakers' Yeast A.F. M

0.6 0.4 0.2

20

0

60

40

80

H3

CI4

100

C ' ~ - A I OYeast

300 300

fl

o

A.F.

H3-AIa E.coli

M 0.6 0.4 0.2

Tube

/

No.

c ' ~ - A IYeast ~ 0

H3-Ala Liver

M 0.6

0.4 0.2

0

2U

40

60

80

100

Tube No.

0

20

40

60

80

100

Tube No.

FIQ. 7. Chromatography of RNase Ti digest of a mixture containing [H'I alanyl- and LC"1 alanyl-tRNA's from various sources (110).

SPECIFICITY IN THE STRUCTURE OF TRANSFER RNA

59

not be forriiylated could be distinguished in their terminal sequences by paper electrophoresis of the RNase TI digest of tRNA's charged with S35-methionine ( 1 1 1 ) .

IV. The Three-Dimensional Structure of Transfer RNA A molecule of tRNA seems to have a definite tertiary structure arising from the many intramolecular hydrogen bonds among the bases, stabilized by metal ions such as Mg++.Although snake venom phosphodiesterase releases the nucleotides one by one from the 3' terminal of a polynucleotide chain, the enzyme removes the terminal pA and the next pC, in the presence of Mg++ion, but no further digestion of tRNA occurs ( 1 1 4 ) . Nishimura and Novelli (115) claimed that the presence of Mg" prevents the digestion of tRNA by B. subtilis ribonuclease, and that some tRNA's retain their ability to accept amino acids (cf. Section V,A) . This appears as if the definite secondary structures essential for their activity are preserved, even if some looser regions have been split by the enzyme. From the analysis of the tRNA digest by venom phosphodiesterase, under optimal conditions for exonuclease activity, Nihei and Cantoni (54) concluded that the minor nucleotides accumulate in the middle part of the tRNA molecule, forming a looplike structure, with the other parts folded back to make a tight double-stranded structure resembling DNA. Thus tRNA would assume a hairpinlike structure as a whole. A similar model was also considered from the results of X-ray diffraction analysis by Spencer et al. ( 6 7 ) . However, as already mentioned in Section 111, the nucleotide sequence of yeast alanine tRNA determined by Holley et al. (65) cannot form such a simple folded structure. Moreover, Spencer and Poole (117) reported recently that the low molecular weight RNA material of the previous analysis (67) was subsequently considered not to be tRNA, but a mixture of degradation products of ribosomal RNA, which might have a considerably ordered structure. Therefore, the higher-ordered structure of tRNA would be more complex than a simple hairpin model. Englander and Englander (118) made hydrogen exchange measurements on yeast tRNA by the use of the tritium-Sephadex method, and found about seventy-seven hydrogen bonds per an average seventynucleotide tRNA molecule, indicating the involvement of about 82% of the nucleotides in a helix form. Felsenfeld and co-workers (119, 120) estimated 76% helix and 71% hydrogen bonds per molecule from the hypochromicity of tRNA. If the N-1 position of the adenine moiety is blocked by hydrogen bonding, the oxidation by monoperphthalic acid is inhibited. Seidel and Cramer (121) calculated the number of bases in-

60

KIN-ICHIRO MIURA

volved in a non-hydrogen-bonded part (a looped structure) by an estimation of the rate of N-1 oxidation of adenine. Augusti-Tocco and Brown (122) found that guanylic and uridylic acids in the nonhelical parts in a polynucleotide react with N-cyclohexyl, N'-P- (4-methylmorpholinium) ethyl carbodiimide iodide, and 4% of the total nucleotides in a tRNA molecule react in the presence of Mg++.These data suggest that there are many hydrogen bonds in a tRNA molecule, but that all the bases do not participate in hydrogen bonding.

1.3-

1.2 -

///'

I

RNA

1.1-

1.0-

I

t

1

20

-

1

30

I

1

40

1

I

50

I

I

60

I

1

70

I

I

00

I

I

O

90

Temperature PC) FIQ.8. Variation of the ultraviolet absorption of nucleic acids at 260 mp with temperature in 1.6 x lo-' M NaCl plus 1.5 x lo-" M sodium citrate (RDV = rice dwarf virus) (123).

We have found that the RNA isolated from the rice dwarf virus (RDV) is double-stranded (123, 124). We compared tRNA with RDVRNA in many experiments under the same conditions. The ultraviolet absorption of RDV-RNA rises sharply in a narrow temperature range as does DNA, while that of tRNA varies gradually with temperature over a wide range as does ribosomal RNA or denatured RDV-RNA, as shown in Fig. 8. Free amino groups react with formaldehyde. The reac-

61

SPECIFICITY IN THE STRUCTURE OF TRANSFER RNA

Tronsfer RNA

0.8

0.8

Ribosome RNA

0.7

0.6

2 0.5

x $

0.4

D

a3 a2

0.I "

0

220 "0 260 280 300 320

220 240 260 280 300 320

Wovelength (rnp)

Wovelength ( m p )

0.9 I.6

RDV- RNA Denotured

0.7 0.6 >

e 0.5

P

P" 0.4 0.3 0.2

0.I

220 240 260

280 300 320

Wavelength ( m p )

Wavelength ( m p )

FIG.9. Variation of the ultraviolet absorption of nucleic acids on treatment with 1.8% formaldehyde in 0.1 M NaCl at 37°C for 30 minutes and 22 hours (RDV = rice dwarf virus) (123). tion may be followed by the increase and the shift to a longer maximum wavelength of ultraviolet absorption. As shown in Fig. 9, tRNA reacts as well with formaldehyde as does ribosomal RNA, whereas doublestranded RDV-RNA and DNA do not, even in the absence of Mg++. RDV-RNA is more resistant to dilute ribonuclease than either tRNA,

62

KIN-ICHIRO MIURA

ribosomal RNA, or heat-denatured RDV-RNA. These results suggest that tRNA differs considerably from completely double-stranded RNA, even if one considers the difference in the chain length between the tRNA and RDV-RNA. Also, the studies on optical rotatory dispersion of tRNA and on circular dichroism show a difference between tRNA and DNA, and give an asymmetrical configuration for tRNA (125128). Keller (129) studied the digestion of tRNA by snake venom phosphodiesterase precisely and observed three steps with different digestion rates. This suggests heterogeneity in the higher-ordered structure of tRNA. Kisselev et al. (130) also suggest such heterogeneity from the behavior of formaldehyde-modified tRNA on digestion by ribonuclease. Marciello and Zubay (131) found three groups of bases according to their reactivity with formaldehyde. Fresco et al. (132) studied the change in ultraviolet absorption of purified alanine, tyrosine, and valine tRNA's on raising the temperature, finding two separate phases in the absorbance vs temperature profile. Fresco et al. (132) suggested that a tRNA molecule contains a t least two helical regions, which might have different base compositions. A model of a tRNA molecule that contains several short double-helical regions has been described (65, 133, 134). Recently, a triple-stranded model resembling a gem-clip has been proposed as a possible secondary structure of tRNA (136). Although a definite tertiary structure of tRNA has not yet been defined, each specific tRNA, which has a specific base composition (see Section 11), may have a specific tertiary configuration, because the melting profiles of a few specific tRNA's differ from each other (132, 136) . The secondary structure of tRNA is broken by heating, acid or alkali treatment, and urea treatment. The change, however, is reversed when tRNA is replaced a t the original temperature or p H (128, 137). These treatments do not affect the amino acid-accepting ability and the transfer activity of the tRNA (138).Arch et al. (139) examined the activity of tRNA from a thermophilic bacteria, B. stearothermophilus, a t high temperature, since the synthetase specific for isoleucine does not become inactivated even a t 80°C. The ATP-PPi exchange reaction accompanying isoleucine activation occurs even a t 8O"C, while the isoleucine acceptance of the tRNA suffered extensive loss a t 80°C. On the other hand, a T,, of 80°C is obtained from ultraviolet absorption measurements. From these experiments, it is suggested that the secondary structure of tRNA is essential for amino acid acceptance. A change in the higher-ordered structure of tRNA thus seems to have some effect on its activities. According to Tissieres et al. (140),

SPECIFICITY I N THE STRUCTURE OF TRANSFER RNA

63

tRNA free of amino acid inhibits RNA polymerase, whereas tRNA charged combined with amino acid does not. Since a conformational change of tRNA was observed between these conditions ( l 4 l ) ,the inhibitory effect of tRNA on RNA polymerase might depend on a specific tertiary structure of tRNA. If this is the case, i t is one instance where a conformational change in a certain biopolymer is related to metabolic control. An ambiguous behavior in the fidelity of code-reading by tRNA is sometimes observed under the influence of pH, temperature, solvent, etc. (142-146). In these cases, changes in the specific tertiary structures of tRNA and/or its partner would cause the misreading. Precise information on the tertiary structure of tRNA as well as primary structure are required for an understanding of its function.

V. Studies on the Functional Sites in tRNA

A. Modification of Specific Sites There are many papers dealing with the modification of nucleic acid bases without scission of the polynucleotide chain and the resulting change in the biological activities of nucleic acids from viruses or bacteria. I n order to elucidate the functional sites of tRNA, i t seems logical to examine the change in activity of such modified tRNA. 1. FORMALDEHYDE TREATMENT

Formaldehyde reacts with the free amino groups of nucleic acid bases. Penniston e t al. (147) compared the inactivation rates of the aminoacylation activity and of the transfer activity onto polysomes for phenylalanine incorporation, using yeast and E . wli tRNA, and found that these inactivation rates were similar to each other. The fact that Mg++inhibits the inactivation shows that a certain higher-ordered structure is required for both the activities, as mentioned in a previous section. 2. NITROUSACIDTREATMENT

Nitrous acid deaminates adenine, guanine, and cytosine. Zillig et al. E. coli tRNA with nitrous acid and found the acceptance of amino acids by tRNA to be inactivated a t different rates for several amino acids. Phenylnlnnyl-tRNA synthesis was strongly inactivatcd, compared with leucine, lysine, and valine. It iiiiglit be that the active site for the synthetase on phenylalanine tRNA contains many purine residues, since HNOa reacts with purines faster than with pyrimidines. Carbon (149) observed that the tRNA transfer to polysomes is inactivated by nitrous acid treatment more than is the aminoacylation step. (148) treated

KIN-ICHIRO MIURA

3. BROMINATION AND METHYLATION

Yu and Zamecnik (150) brominated tRNA, observing that changes in uracil, in cytosine, and, to a lesser extent, in guanine caused inactivation of amino acid acceptance. The inactivation rates of tRNA from yeast and from E . coli differ from each other. The order of sensitivity in yeast was: phenylalanine > valine > lysine; on the other hand, in E. coli it was: lysine > valine > phenylalanine. Based on these experiments, Yu and Zamecnik (150) claimed that the sites responsible for the recognition of the synthetase differ among organisms, so that these sites might be distinguished from the coding sites for messenger RNA, if one assumes, for the latter sites, the universality of the complementary sequence to a codon in mRNA. They further compared the inactivation of amino acid acceptance and of amino acid transfer of tRNA upon bromination (151). The former activity was more sensitive than the latter. They suggested that the sites responsible for these two activities may differ from each other, and that the acceptor activity requires a definite secondary structure. Weill e t al. (152) studied the change of the activities of tRNA, applying the fact that bromination does not affect adenine and that methylation by dimethylsulfate does not attack uracil. The inactivation of transfer activity seems to correlate with the modification of the anticodon. For example, methylation inactivates phenylalanine transfer but not that of lysine, while bromination inactivates lysine transfer. However, the accepting activity was not related to modification of the anticodon, as methylation affected lysine attachment to tRNA and bromination that of phenylalanine. From these results, Weil et al. (152) considered that the active sites differ from each other. Further, in the presence of poly U, brominated tRNA favored the incorporation of amino acids other than phenylalanine into polypeptide (153). It is conceivable that brominated C and U might act as A. If such a change occurred in the anticodon sequence of tRNA, the incorporation of amino acids other than phenylalanine seems reasonable. 4. SEMICARBAZIDE TREATMENT

If only one specific base is modified, one can expect to get more clear information as to the functional sites of tRNA than in the cases of the above-mentioned modifications. Hayatsu, Ukita, et al. (154) found that semicarbazide combines selectively with the amino group of cytosine in RNA under mild conditions. Muto e t al. (155) studied the effect of the semicarbazide modification of tRNA on amino acid acceptance and found that the extent of reduction of the activity of individual tRNA’s varied as shown in Fig. 10. The order of inactivation

SPECIFICITY I N T H E STRUCTURE OF TRANSFER RNA

65

of the amino acid-accepting activity was: valine > phenylalanine, alanine, glycine > tyrosine. Of these, the tyrosine-accepting activity was exceptionally resistant to the modification. Holley et al. (20) analyzed the base composition of yeast tyrosine, valine, and alanine tRNA’s, separated by countercurrcnt distribution, and found the cytosine contents to be 26.7%, 27.5%, and 29.9%, respectively. On the supposition that the average chain length of the tRNA molecule is seventyseven nucleotides (48), tyrosine tRNA contains twenty-one cytosine

FIG.10. Effects of semicarbazide (SC) modification on the amino acid-accepting activity of Torula yeast tRNA (a) and E . coli tRNA (b). The tRNA was treated with 2 M semicarbaeide at 37°C for various periods except the plot of 79% semicarbazide/cytosine (C), for which the tRNA was treated with 3.3 M semicarbazide for 72 hours (164,166).

residues, valine tRNA twenty-one, and alanine tRNA twenty-three, respectively. The differences in the cytosine contents of these tRNA’s are thus only one or two residues. Therefore, the degree of the inactivation by the semicarbazide modification seems not to depend on the cytosine content of the molecule. Inactivation of 50% of the amino acid acceptance requires modification of ten cytosine residues for tyrosine tRNA, whereas modification of only two cytosine residues is needed for valine tRNA and four for alanine tRNA, provided each individual tRNA is modified to a similar extent. For every tRNA, the more the modification of cytosine proceeded, the more profound was the inactivation of amino acid acceptance, although the rate of the inactivation was different for each tRNA. The

66

KIN-ICHIRO MIURA

inactivation could be caused, even if the sites responsible for the recognition of aminoacyl-RNA synthetase involved no cytosine, by the modification of bases contained in sequences other than the recognition sites through an effect on tertiary structure. Since semicarbazide does not react with DNA, non-hydrogen-bondcd rytosinc rcsiducs in tRNA would be the first to be modified by semicarbazide. When tRNA reacts with semicarbazide a t high temperature, with destruction of its secondary structure, the modification proceeds quickly. On the other hand, when tRNA is modified by semicarbazide a t low temperature and with the addition of Mg", only a restricted part of the cytosine residues seems to react with the reagent, so that no loss of amino acid-accepting ability is observed for some tRNA's, while others are inactivated (155). The CC sequence in the amino acid-accepting terminal sequence (CCA) would react easily with some modifiers, as this part is believed to be free of the tightly hydrogen-bonded structure (114). In order to distinguish the modification of this part and other parts contained in the functional sites, enzymatic repair of the CCA sequence with semicarbadde-CTP was tried (162). However, semicarbazide-CTP was not incorporated into the terminal sequence. The enzyme responsible for the CCA sequence repair may have strict specificity for CTP. The rates of inactivation of aminoacylation of tRNA by semicarbazide for a few amino acids differed in yeast tRNA and E . coli tRNA as shown in Fig. 10. A derivative of semicarbazide, Girard P reagent: H~N-NH-CO-CH~-&(CH~)&OH~ c1-

also combines only with cytosine residues (156). The modification of tRNA by this reagent causes inactivation of amino acid acceptance a t similar rates for tyrosine, alanine, and valine. It is thus quite different from semicarbazide modification. This reagent might more effectively modify the common C residues essential for accepting activity for all amino acids, such as the C in the CCA sequence a t the amino acidaccepting terminal. Another possible reason for a common inactivation might be a change in the electrical charge of a tRNA molecule owing to the modification by this reagent. 5. HYDROXYLAMINE TREATMENT

It is known that hydroxylamine modifies cytosine a t pH 6 and uracil a t pH 9 selectively (157,158). The specificity of this modification for a definite base was confirmed when tRNA was treated under these conditions (159-161). This modification does not split the polynucleotide

SPECIFICITY IN THE STRUCTURE O F TRANSFER RNA

67

chain (159-161). Takanami and Miura (161) examined its effect on the amino acid transfer of intact tRNA and of tRNA without the CCA sequence, which was obtained after inoculation with so-called “pH 5 enzyme.” The CCA terminal sequence of the modified tRNA was repaired enzymatically with CTP and ATP. After treatment with nitrous acid or semicarbaxide, repair was not achieved. The C in the CCA terminal sequence as well as the internal C residues participate in the interaction betwecn ribosomes and tRNA. The terminal CCA sequence might act as a locating site on ribosomes. Hydroxylamine modification a t p H 9 leads to the conclusion that internal U in tRNA is also required for binding to ribosomes. pH 9.0 I N NH20H37’C (Time, hr)

FIG. 11. Effects of hydroxylamine modification a t pH 9 on the amino acidaccepting activity of Torula yeast tRNA. The tRNA was treated tre!ated with M NHzOH at pH 9.0 at 37°C (166).

Regarding the amino acid-accepting ability of hydroxylaminetreated tRNA, the rates of inactivation are different for some amino acids as in the case of semicarbazide modification. Tyrosine acceptance by yeast tRNA was not as much affected by the treatment with hydroxylamine a t pH 6 as was the acceptance of other amino acids (166). As this is in good agreement with the results of semicarbaxide treatment, the modification of cytosine residues does not seem to destroy directly the acceptor activity of tyrosine tRNA of yeast. Hydroxylamine treatment a t pH 9-10, resulting in a modification of uracil, inactivates the amino acid-accepting ability of yeast tRNA as shown in Fig. 11. It is significant that lysine acceptance by tRNA is the

68

KIN-ICHIRO MIURA

most destroyed by the uracil modification (155, 159, 160) while phenylalanine tRNA was the least sensitive one of those tested. The anticodon sequence of the lysine tRNA is considered to be UUU and that of phenylalanine AAA. Including the cases of other amino acids, the degree of inactivation by the modification seems to correlate with the amount of U in the anticodon sequence (155, 159, 160). It is claimed that the anticodon sequence in tRNA acts not only as the code-reading site for messenger RNA in the transfer step but also as the synthetaserecognizing site in the accepting step of amino acids. The inactivation curves of tRNA’s in this treatment (Fig. 11) are single-hit curves. Kiselev and Frolova (159) showed also that the inactivation for lysine acceptance is a single-hit event. An attack on one of the uracil residues in an anticodon sequence would induce a loss of synthetase recognition. CernB e t al. (160) compared the inactivation of the accepting activity of uracil-modified tRNA’s from yeast, r a t liver, and E. COG. The degree of inactivation of this activity for individual amino acids varies among the species. But, in every case, the inactivation of lysine tRNA was greater than that of phenylalanine tRNA. Cernh e t ul. (160) observed no marked difference between the transfer of an amino acid from normal and from modified (hydroxylamine treatment a t both pH 6 and p H 9) tRNA to proteins. Grachev e t al. (104) found that one of the degenerate valine tRNA’s is quite resistant in its valine-accepting activity after the hydroxylamine treatment, while the other is sensitive (see Section 111,C). 6. ULTRAVIOLET LIGHTAND X-RAYIRRADIATION

When RNA is irradiated with ultraviolet light, uracil and cytosine are modified. However, the change in cytosine is readily reversed to natural cytosine, so that one can achieve a modification of uracil alone by such irradiation. Zachau (162) found that the UV-induced inactivation rates for amino acid acceptance of yeast tRNA are Lys > Ser > Phe, and stated the possibility that the anticodon sequence in tRNA participates in the recognition of the synthetase as well as in the code-reading of mRNA. Wada e t al. (163) also observed that lysine acceptance of tRNA’s from yeast and E. coli is greatly inactivated by ultraviolet irradiation, compared with the acceptance of other amino acids. Kawade e t al. (164, 165) compared survival activities of phenylalanine acceptance and transfer of E. coli (or yeast) tRNA after ultraviolet irradiation, and found that the inactivation of these activities is a single-hit event and that the rates of the inactivation are different for each activity. They concluded that the functional sites of acceptance

SPECIFICITY I N THE STRUCTURE OF TRANSFER RNA

69

and transfer in phenylalanine tRNA do not overlap as regards the uracil residues contained in them. Even if the anticodon is contained in the functional sites for both the synthetase recognition and the code-reading of mRNA, the number of uracil residues not contained in an anticodon sequence but essential for function, for example, for binding with ribosome or with enzyme, may be different in the steps of acceptance and transfer of an amino acid. Therefore, the possibility that the anticodon sequence is required for the two functions of tRNA is not eliminated. Wacker and Chandra (166)showed, by ultraviolet or X-ray irradiation of tRNA, that the anticodon sequence in tRNA is essential for the code-reading on mRNA.

7. PHOTOOXIDATION Uehara et al. (167) found that adenine and guanine are photooxidized by visible light in the presence of riboflavin. Tsugita e t al. (268) observed that the acceptance of phenylalanine and lysine by E. coli tRNA is inactivated with first-order kinetics by this treatment. They also observed the loss of amino acid acceptance of tRNA by visible light in the presence of lumichrome or methylene blue. Simon and van Vunakis (169) found that the photosensitized oxidation in the presence of methylene blue causes the preferential destruction of guanine residues in nucleic acid. They also studied the effect of this modification on synthetic polynucleotide as a messenger RNA (170). Recently, Kuwano et al. (171) found that in the presence of methylene blue or carcinogenic 4-nitroquinoline-N-oxide (about M ), visible light irradiation induces a loss of the ultraviolet absorption of guanine residues in tRNA, without breaking the polynucleotide chain. By this treatment, amino acid acceptance was damaged a t various rates, that of proline especially. The anticodon of proline tRNA is considered to be GGG, from the Watson-Crick type of conjugation between the codon in mRNA and the anticodon in tRNA. For other amino acids, the rate of inactivation of the acceptance activity also seems to correlate with the content of guanine in the anticodon. These results also support the hypothesis that the anticodon sequence is included in a functional site responsible for amino acid acceptance. 8. CYANOETHYLATION

As mentioned in Section 11, many kinds of “minor” nucleotides occur in tRNA. What role do these play in the activity of tRNA? I n order to elucidate this problem, it is desirable to develop selective modification methods for each minor component. Ofengand (172) and Yoshida and Ukita (173)and Chambers et al. (17%) found independ-

70

KIN-ICHIRO MIURA

ently that pseudouridine is cyanoethylated by acrylonitrile. Ofengand (172) reported an 87% loss of amino acid-accepting activity of tRNA after 1 hour treatment with 1 M acrylonitrile (pH 9.1) a t 60°C. We (174) studied the same activity of cyanoethylated tRNA for a few amino acids. As shown in Fig. 12, pseudouridine in tRNA is modified by acrylonitrile (at pH 8.6) according to the time of incubation in 0.01 M salt solution, while in 0.51 M salt solution the modification stopped at about 33%. Under both conditions, the amino acid-accepting ability of tRNA was destroyed with time of treatment a t different rates for alanine, tyrosine, and valine. Since, actRNA modified in 0.01 M salt conc.

100

100

--3

100

3

-

0

8

6

21 c .?

g 50

y"--

2

2

50s

-._o9 -%r

........_ y

A.

c

c

n;'; tRNA modified in (X5lMsalt conc.

50

Q---r

0

._ B >

$ 2 m

U 0.

2 5 0 24 34 48 Reaction time (hr)

s -

. I -

c .c

0

5

m

5 10

24 34

40

Reaction time (hr)

FIG. 12. Effects of cyanoethylation on the amino acid-accepting activity of Torula yeast tRNA. The tRNA was treated with acrylonitrile in 0.01 M salt solution (a) and in 0.51 M salt solution (b) at pH 8.5 at 37°C (174).

cording to Holley e t al. ( g o ) , the content of pseudouridine would be four residues in valine, three in tyrosine, and two in alanine tRNA, respectively, the rate of inactivation of amino acid acceptability appears to be correlated with the pseudouridine content. However, the inactivation of amino acid acceptance may not depend only on the modification of pseudouridine as a small part of the uracil is also modified by acrylonitrile a t about one-tenth the rate of the pseudouridine modification, as shown in Fig. 12, and inosine residues are also cyanoethylated. HYDROLYSIS WITH ENZYMES 9. PARTIAL After the free phosphate a t the 5' terminal of tRNA is removed by phosphomonoesterase, the amino acid-accepting ability remains as before the treatment (83, 176).

SPECIFICITY IN THE STRUCTURE OF TRANSFER RNA

71

Spleen phosphodiesterase splits nucleotides from the 5’ terminal of tRNA one by one. When about 5% of the total nucleotides has been released, amino acid acceptance is almost nil (175, 176). Nishimura and Novelli (115) found that, when tRNA is treated M Mg++, with RNase from B. subtilis or RNase TI in the presence of the inactivation of amino acid acceptance of tRNA proceeds a t different rates for each amino acid; for some amino acids no inactivation is seen. Some tRNA’s that do not lose amino acid acceptability after the RNase treatment lose their transfer activity (e.g., valine, leucine, and alanine tRNA from E. coli) while some tRNA’s do not lose either the activities of acceptance or transfer (e.g., phenylalanine, lysine, and tyrosine tRNA’s) (177). From these facts, Nishimura and Novelli (177) considered that the sites essential for the two steps of amino acid acceptance and transfer are separate from each other. I n the cases where these RNases do not destroy the tRNA activities, the tertiary structure should be maintained firmly by Mg++in spite of some loss in primary structure. If such is the case, a certain rigid tertiary structure of tRNA seems required for its function. The inactivation rates of the acceptor activity of tRNA by these RNase treatments are different depending on the sources of the tRNA’s (178).

B. Inhibition by Oligonucleotides of Aminoacylation of tRNA Each tRNA molecule combines with a definite amino acid with the aid of a specific aminoacyl-RNA synthetase. Attempts have been made to modify those structures by which a tRNA molecule recognizes a specific synthetase, as described above (Section V,A). A different approach for this purpose was devised by us. This is a study of the inhibitory effect of various kinds of polynucleotides on the aminoacylation of tRNA (179). It was assumed that inhibition would be observed in the case where the polynucleotide contains the specific structure corresponding to the synthetase recognition site in tRNA, by blocking the active site in the synthetase. The following observations have been made. Transfer RNA, oxidized by periodate to destroy the acceptance of amino acid, inhibited the synthetase. Similar results have been reported by others (18el82). Ribosomal RNA showed no inhibitory effect, but ribosomal RNA partially degraded by mild alkaline trcatment had an inhibitory effect. A mild alkaline digest of tRNA was also found to be inhibitory. The inhibition by these poly- (or oligo-) nucleotides was competitive with the aminoacylation of tRNA, and it did not depend on the terminal phosphate of the oligonuclcotide. For valine incorporation into tRNA, oligonucleotides longer than tetranucleotides in the ribonuclease digest

72

KIN-ICHIRO MIURA

of RNA showed an inhibitory effect, whereas dinucleotides showed no effect and trinucleotides showed little effect (see Table X) . Deutscher (183) studied the effect of a several kinds of synthetic polynucleotides on aminoacyl-RNA synthetases in a mammalian system, and observed that specific kinds of polynucleotides inhibit the attachment of a specific amino acid to tRNA, for glutamic acid, serine, and leucine. TABLE X INHIBITORY EFFECT OF OLIGONUCLEOTIDES FOR VALYbtRNA FORMATION (179) Nucleotide length of inhibitora Control (no inhibitor) 2 3 4

5

6

7

Concentration of inhibitor (0.D.ssomp)

0 9.8 15 5.2 10 16 1.1 2.2 3.3 1.4 2.9 4.3 0.9 1.7 2.6 0.8 1.5 2.3

Incorporation of valine into tRNA (%) 100 99 99 100 91 89 100 70 12 13 14 13 14 13 13 13 12 10

a This corresponds to the fraction number of the oligonucleotides eluted from DEAE-cellulose by a salt solution containing urea [D. Bell, R. V. Tomlinson, and G . M. Tener, Biochem. Biophys. Res. Commun. 10, 304 (1963)l. The number also shows the average chain length of polynucleotides.

The inhibitory effect of several kinds of synthetic oligonucleotides on the incorporation of amino acid into tRNA was studied, using a yeast system. I n our experiments (184), the rate of the formation of phenylalanyl-tRNA was strongly inhibited by the addition of oligo A and only slightly inhibited by oligo U, while oligo C showed no inhibition (Fig. 13a). The inhibition of phenylalanyl-tRNA formation by oligo A was of a typical competitive type, as seen in the Lineweaver-Burk plots of l / v against l/s for various concentrations of the “inhibitor” (Fig. 13d). The rate of lysine incorporation into tRNA was strongly inhibited

73

SPECIFICITY I N THE STRUCTURE O F TRANSFER RNA

0 2 4 6 8 Amount of the Inhibitor Added (O.DzW\ per Tube

I

~

I

I

I

I

I

2 4 6 8 0 Amount of the Inhibitor Added ( O . C J . ~ ~per ~ ) Tube

I

Oligo A *

.

a

0.4

I (C)

,

1

,

1

1

1

1

1

1

2 4 6 8 Amount of the Inhibitor Added(O.D,,, )per Tube

0

I

I

I

2

1/s

FIQ. 13. Inhibitory effect of oligonucleotides for phenylalanyl-tRNA (a), lysyltRNA (b), and prolyl-tRNA (c) formation. (d) shows competition between tRNA and oligo Ap for yeast phenylalanyl-tRNA synthetase (Lineweaver-Burk plot) I: no inhibitor; I1 oligo A (inhibitor), 4.8 O.D. units per 0.6 ml. (184).

by oligo U in a competitive manner, only slightly inhibited by oligo A, and not inhibited by oligo C, as shown in Fig. 13b. The incorporation of proline into tRNA was inhibited strongly by oligo G, slightly inhibited by oligo C, but not inhibited by either oligo A or oligo U (Fig. 13c). If a tRNA molecule contains a complementary sequence for a codeword of messenger RNA as suggested in the adaptor hypothesis of Crick ( I ) , phenylalanine RNA may contain AAA as a coding site for messenger

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RNA, since the code letter of messenger RNA for phenylalanine is UUU (70, 71).The presence of such an anticodon sequence was observed in a specific tRNA molecule, as mentioned in Section III,B. The fact that the incorporation of phenylalanine into tRNA is inhibited by oligoadenylate clusters might be considered as follows: The enzyme recognition site in phenylalanine tRNA would be also an adenylate cluster. A slight inhibition of phenylalanine incorporation into tRNA by oligo U would be caused by the formation of complementary complexes for the adenylate clusters in tRNA. The formation of such complexes could reduce the velocity of the reaction. The inhibition of phenylalanine incorporation by oligo U did not proceed over 15% with increasing amounts of oligo U. I n the case of incorporation of lysine and proline into tRNA, the oligonucleotide complementary to the messenger RNA code letter showed competitive inhibition, and the same sort of oligonucleotide as the messenger RNA code letter caused slight inhibition, since the messenger RNA code letters for lysine and proline are AAA and CCC, respectively (7-9). If the anticodon part, that is, the coding site in tRNA, consists of complementary nucleotides to the messenger RNA code and the code letter is universal for all species, we might assume, depending on the above-mentioned results, that the coding site acts also as an enzyme-recognizing site when a specific amino acid is incorporated into the terminal site of the specific tRNA by the specific aminoacylRNA synthetase, or two anticodon sequences in a tRNA molecule may act as a coding site and an enzyme-recognizing site, respectively. Usually a cross reaction between a tRNA and a synthetase from a different source shows species specificity, suggesting that the enzymerecognizing site in tRNA differs from species to species. Loftfield and Eigner ( I % ) , however, found that the Michaelis constants are quite similar for the aminoacylation of tRNA by the use of heterologous enzyme as compared with the homologous enzyme. They concluded that the enzyme-recognizing site may be identical throughout all species and it may be unnecessary to distinguish the enzyme-recognizing site from the coding site in tRNA. But as we have mentioned, modification of tRNA of different species by bromination causes inactivation of aminoacylation a t different rates. From these results, Yu and Zamecnik (150, 161) considered that the enzyme-recognition site in tRNA varies from species to species and is separate from the coding site in the same molecule. Other workers came to similar conclusions after studying the effect of modifications of tRNA by nitrous acid (149), by methylation and by bromination (15.2), or by ultraviolet light irradiation (164).These treatments, however, are not specific for one kind of nucleotide. They may

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conceivably inactivate tRNA not only by a direct modification of the enzyme-recognition site, but also through changes in the tertiary structure essential for the tRNA activity arising from modification of parts other than the enzyme-recognizing site. Such a situation is suggested from the inactivation experiments by semicarbazide modification of cytosine residues (154) or by ribonuclease treatment (17‘7), and this possibility is discussed even in those reports that concluded that the enzyme-recognizing site and the coding site are different. That the enzyme-recognizing site and the coding site in tRNA are identical was also suggested by a few modification experiments. Penniston et al. (147) compared the inactivation rates of the aminoacylation activity and of the transfer activity to polysome for formaldehydemodified tRNA, and found that these inactivation rates were similar to each other. Zachau (162) modified the uracil residues in tRNA with ultraviolet light, which makes uracil dimerize intramolecularly, finding that the incorporation of lysine into tRNA decreased considerably more than that of phenylalanine. H e considered that the anticodons of lysine tRNA and of phenylalanine tRNA might be UUU and AAA, respectively, and they might also act as the enzyme-recognition site, so that the ultraviolet inactivation is more effective for the inhibition of lysine incorporation. Similar experimental results (156,159, 160, 171) have also been obtained by a specific modification of uracil residues in tRNA by treatment by hydroxylamine a t p H 9, and of guanine residues by photooxidation (see Section V,A). These data agree with our results of the “inhibition” experiments and support its interpretation. The fact that the kinds of inhibitor oligonucleotides are consistent with anticodon sequence in the case of yeast are also found in the system of E . coli (186).However, even if the site responsible for the recognition of the synthetase in tRNA is universal for every organism, other parts than the site itself would have different primary structures to cause inevitably the differences in tertiary structure that are related to the interaction with the synthetase. I n fact, the structures of a specific tRNA and synthetase seem to be different for various organisms (see Section 111), so that the efficiency of amino acid attachment to tRNA must vary depending on the combination of tRNA and synthetase from different organisms. The diversity in the inactivation rate of modified tRNA among different organisms (see Section V,A) may depend also on the variety in the structure. Hayashi (187) fractionated tRNA with a protamine-coated Kieselguhr column to concentrate tyrosine tRNA; these fractions were oxidized by periodate and the “inhibition experiments” were tried. The fraction rich in oxidized tyrosine tRNA inhibited tyrosine acceptance;

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the oxidized fraction that contained no tyrosine tRNA did not inhibit. Similar observations have been made by Torres-Gallardo and Kern (182).However, when the oxidized tRNA from E. coli is used in the inhibition experiment with the yeast system, the oxidized tRNATyr (E. coli) did not inhibit, whereas the oxidized tRNA that did not contain the tyrosine tRNA from E. coli showed inhibition. I n the E. coli system, too, oxidized yeast tRNA gave similar results. I n some heterologous combination, a paradoxical phenomenon arises as fractions other than tyrosine tRNA inhibit tyrosine attachment to tRNA. From only the above-mentioned results, it may be too early to conclude whether the synthetase-recognition site in tRNA depends on the biological species, or whether the enzyme-recognition site in tRNA is distinguished from the mRNA coding site. When tRNA interacts with the synthetase or with mRNA on polysomes, there are some essential structures for these interactions or bindings besides the enzyme-recognizing site or the mRNA coding site. If one considers further the tertiary structure of both the tRNA and the synthetase the diversity in the structure essential for this interaction would increase. It has been observed that a slight change in temperature or salt concentration, etc., or an addition of organic solvent causes misreading of mRNA (142, 14.9) (“ambiguity” of the code). Similar phenomena would occur also in the interaction of tRNA and synthetase and its appearance may differ depending on the sources. According to Sarin and Zamecnik (146),addition of an organic solvent affects the activities of tRNA both on acceptance and transfer of amino acids. Therefore, one must also observe the effect of the environment on these activities of tRNA, especially in the case of heterologous combinations of tRNA and synthetase. These problems should be studied more precisely. ACKNOWLEDGMENTS The author is indebted to Professor Fujio Egami, Professor Yonosuke Ikeda, Professor Tyunosin Ukita, Dr. Shosuke Takemura, Dr. Mitsuru Takanami, Dr. Masazumi Miyazaki, Dr. Hiroshi Hayashi, Mr. Torao Ishida, Mr. Akira Muto, Mrs. Yutsuki Hayashi, Mr. Yushin Fujimura, Miss Keiko Matsueaki, Mr. Michihiko Kuwano, Dr. Toshio Okamoto, Dr. Hikoya Hayatsu, Mr. Akimitsu Yoshida, and Mr. Kiyomi Kikugawa for their discussions and able collaboration in a number of the works presented here and for their hospitslity.

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177. S. Nishimura and G. D. Novelli, Proc. Natl. Acad. Sci. US. 53, 178 (1965). 178. S. Nishimura and G. D. Novelli, Biochim. Biophys. Actu 80, 340 (1964).

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179. H. Haymhi and K. Miura, J. MoZ. Biol. 10, 345 (1964). 180. J. Preiss, P. Berg, E. J. Ofengand, F. H. Bergmann, and M. Dieckmann, Proc. Natl. Acad. Sci. US.45, 319 (1959). 181. A. G. So and E. W. Davie, Biochemistry 2, 132 (1963). 18.8. J. Torres-Gallardo and M. Kern, Proc. NutZ. Acad. Sci. U S . 53, 91 (1965). 183. M. Deutscher, Biochem. Biophys. Res. Commun. 19, 283 (1965). 184. H. Hayashi and K. Miura, Nature 209, 376 (1966). 186. R. B. Loftfield and E. A. Eigner, Actu Chem. Scand. 17, Suppl. 1, 117 (1963). 186. H. Hayashi and K. Miura, Cold Spring Harbor Symp. Quant. Biol. 31 (1966),

in preas. 187. H. Hayashi, J . MoZ. Biol., 19, 161 (1966).

Synthetic Polynucleotides A. M. MICHELSON, J. MASSOUL&, Institut de Biologie Physico-Chimique, Paris, France AND

W. GUSCHLBAUER~

Dkpartment de Biologie, Centre $Etudes Nuclkaires de Saclay, Gif-sur-Yvette, France

I. Introduction . . . . . . . . . . . . . . . . 11. Preparation of Polynucleotides . . . . . . . . . . . 111. Techniques for Investigating the Physical Chemistry of Polynucleotides . . . . . . . . . . . . . . . . A. MoleculariWeight Determination by Ultracentrifugation, Diffusion, and Viscosimetry . . . . . . , . . . . . B. Thermal Stability . . . , . . . . . . . . . C. Characterization of-Nucleotide Material by Ultraviolet Absorption Spectra . . . . . . . . . . . . . . . D. Optical Rotatory Dispersion and Circular Dichroism . . . E. Titrimetry. . . . . . . . . . . . . . . . F. Infrared Spectroscopy . . . . . . . . . . . . G. X-Ray Diffraction, Low-Angle X-Ray Scattering, and Light Scattering . . . . . , . . . . . . . . . . H. Nuclear Magnetic Resonance (NMR) and Electron Spin Resonance (ESR) . . . . . . . . . . . . . . . I. Electron Microscopy. . . . . . . . . . . . . J. Equilibrium Density Gradient Centrifugation . . , . . IV. Homopolynucleotides . . . . . . . . . . . . . A. Polyguanylic Acid , , . . . . . . . . . . . B. Polyinosinic Acid. . , . . . . . . . . . . . C. Polyuridylic Acid. . . . . . . . . . . . . . D. Polyadenylic Acid . . . . . . . . . . . . . E. Polycytidylic Acid . . . . . . . . . . . . . V. Polynucleotide Complexes . . . . . . . . . . . . A. Complexes between Poly G and Poly C , . . . . . . B. Complexes between Poly A and Poly U . . . . . . . C. Thermal Dissociation of Poly A . Poly U and Poly A * 2 Poly U D. Stability of Poly A . Poly U and Poly A . 2 Poly U at Alkaline E. F. G. H.

pH . . . . . . . . . . . . . . . . . . Stability of Poly A . Poly U and Poly A 2 Poly U at Acid pH Base Pairing in Poly A . Poly U and Poly A . 2 Poly U . . . Thermodynamics of the Interaction between Poly A and Poly U Alternating Copolymers. . . . . . . . . . . , 9

'Helen Hay Whitney Foundation Fellow. 83

84 84 85 85

87 87 92 94 94 95 97 97 98 98 98 99 101 101 103 104 104 106

108 110 110 111 111 113

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VI.

VII. VIII. IX. X. XI.

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I. Association of Poly A and Poly I . . . . . . . . . J. Associations Involving Hypoxanthine and Cytosine. . . . Role of Sugar Phosphate Backbone . . . . . . . . . A. The Phosphate Residues . . . . . . . . . . . B. TheSugsr. . . . . . . . . . . . . . . . C. Right- or Left-Handed Helices, Parallelism, andiAntiparalle1ism. . . . . . . . . . . . . . . . . . Reversibility . . . . . . . . . . . . . . . . Displacement Reactions. . . . . . . . . . . . . Polynucleotide Analogs . . . . . . . . . . . . . Theory and Practice of Ilelix-Coil Transitions . . . . . . Factors Governing Structure . . . . . . . . . . . References. . . . . . . . . . . . . . . . .

116 116 119 119 120 122 122 123 125 130 131 134

1. Introduction Synthetic polynucleotides are of considerable interest not only from the biological viewpoint of code cracking but also as grossly simplified models for the study of the numerous physical properties manifested by nucleic acids. This review is limited to physical studies of structure in synthetic polynucleotides, including homopolymers of naturally occurring nucleotides and of various analogs, but is by no means exhaustive. For earlier reviews see reference 1.

II. Preparation of Polynucleotides Perhaps the least understood of enzymes catalyzing the formation of high molecular weight polymers is polynucleotide phosphorylase ( 2 ) . Although the function and detailed mechanism of action of polynucleotide phosphorylases remain somewhat obscure, such enzymes, isolated from various bacterial sources, have proved to be of great utility for the preparation of polyribonucleotides from the appropriate ribonucleoside 5’-pyrophosphates. Large quantities of polynucleotides can be prepared relatively easily from commercially available diphosphates. I n general, primers are not mandatory, and the enzyme shows a very low substrate specificity with respect to the purine and pyrimidine base. A wide variety of polyribonucleotide analogs can thus be prepared by the action of polynucleotide phosphorylase on the analog ribonucleotide pyrophosphates. Examples include NMe-UDP, 5Br-UDP, 5F-UDP, N6-hydroxyethyl-ADP, and isoadenosine diphosphate (3,4 ) . Whereas homopolymers of A, C, I, and U have been available for some considerable time, the preparation of poly G remained extremely difficult until recently. Suitable modification of the incubation conditions has rendered the preparation in quantity of this polymer equally feasible (6). I n general, the products are somewhat polydisperse.

SYNTHETIC POLYNUCLEOTIDES

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Although preparation of an analog diphosphate followed by polymerization is useful in many cases, an alternate approach lies in the modification of a preformed “natural” polynucleotide. Of course, any such modifications must be achieved under experimental conditions that do not lead to extensive degradation of the polynucleotide chain. Bromination and methylation of various nucleic acids have been described ( 6 ) and from a preparative viewpoint the chemical methylation of homopolyribonucleotides has been extensively studied (7). Other treatments include the action of ultraviolet light (8),hydroxylamine (9),and nitrous acid ( l o ) , and the acetylation of cytosine amino groups with acetic anhydride (11). Apart from polynucleotide phosphorylase, other enzymes catalyzing polynucleotide formation have been used. DNA polymerases are of use not only for replication of given DNA’s but also for synthesis (under suitable conditions) of poly (dA-dT) (12) (alternating) and poly dG * poly dC (homopolymers) (15) in addition to a variety of deoxy analogs. PolydA has also been prepared using a primer ( 1 4 ) . Similarly, RNA polymerase can give rise to polyribonucleotides of known sequence by the use of small, chemically synthesized oligodeoxynucleotides ( 1 5 ) . As a result of “slippage” during transcription, high molecular weight polynucleotides are obtained. Physical studies of such products have not been reported in detail as yet.

111. Techniques for Investigating the Physical Chemistry of Polynucleotides This section very briefly reviews some technical aspects of the physical chemistry of polynucleotides. For more detailed descriptions of the techniques mentioned, reference is made a t appropriate points to pertinent books, reviews, and specialized articles.

A. Molecular Weight Determination by Ultracentrifugation, Diffusion, and Viscosimetry

Ultracentrifugation in combination with diffusion and viscosity measurements has been widely used in the field of nucleic acids and proteins to obtain information on the size and shape of such macromolecules. Several books (16-19) and specialized reviews (ZO-24) describe these techniques and their applications. A priori, any molecular weight determination on a polymer th a t is not physically and chemically uniform will yield an average molecular weight. I n the field of the nucleic acids, two kinds of molecular weight

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are frequently used : the number-average molecular weight, M,, and the weight-average molecular weight, M,. They are defined as follows :

and

where f ( M )dM represents the weight fraction of material with molecular weight between M and M d M , Ni the number of molecules of kind i (i-mers) present in the mixture, Mi the molecular weight of the i-mer, and g the number of grams of i-mer. The ratio Mw:M , is a measure of the homogeneity of a polymer sample (see Table 17-1 in ref. 19). I n the ideal case where all molecules have the same chain length and thus the same molecular weight, M , equals M,. For nucleic acids this can actually be the case and for tRNA this ratio is close to unity (25).I n the case of synthetic polynucleotides, this ratio will always be very much larger. The number-average molecular weight is generally determined by osmotic pressure (,%a). More frequently noted in the literature are molecular weights obtained by sedimentation velocity or sedimentation equilibrium measurements. While the latter measurements give the M,, the former give the actual molecular weight a t the sedimenting peak. Quite frequently only the sedimentation from velocity measurements is quoted. This has the following connection with the molecular weight (in a simplified form)

+

(%?): s20,w

=

M(l

- Pp) Nf

(3)

where szo,w is the sedimentation coefficient (at 2OoC and corrected for water), M the molecular weight, the partial specific volume of the solute, p the density of the solution, N = 6.02 X loz3 (Avogadro’s number), and f the frictional coefficient, which is a function of size, shape, and hydration of the sedimenting polymer. Because of the greatly different values f can take depending on these properties, the sedimentation

SYNTHETIC POLYNUCLEOTIDES

87

coefficient will mean quite different things for structured and unstructured polymers. Certainly there exist correlations between szo and molecular weight for random-coiled and structured polynucleotides ; these calibration plots, however, are not necessarily applicable to any other polymer. Thus it is now well-established that, for instance, oligonucleotides of guanosine can easily associate and assume a considerably higher molecular weight than if each polymer chain were free; even GMP can form polymerlike gels. It has been noted that, in a sample of the copolymer of U and G, poly (U,G), with an szo,u, of 5.6, some 80% of the materials is dialyzable in 4 N urea. Thus the sedimentation constant can frequently be a source of error when highly polymerized polynucleotides are used.

B. Thermal Stability What is called, in nucleic acid chemistry, the “melting point” is an operational term for a temperature-dependent change in structure. Therefore the expression “transition temperature” is much more appropriate. It describes the temperature a t which a helical polynucleotide assembly rearranges itself either to a different helical arrangement or to the randomly coiled state. This thermal transition, long known in polymer and protein chemistry, was first shown to be a characteristic property of nucleic acids by Thomas (66).Since then, T, determinations have been used routinely to characterize nucleic acids and polynucleotides. This transition from one helical to a different helical form or to the coiled state is accompanied by changes in various physical properties of the polynucleotide, for example in ultraviolet light absorption or optical rotatory dispersion (ORD). Changes in infrared spectra, circular dichroism, and buoyant density also occur (see below). It has been established that the T, is dependent on ionic strength, divalent cations being more effective than monovalent salts (by a factor of about 104). Further, it is now quite clear that, below a limiting molecular weight, chain length becomes a determining factor. As already established for DNA, the content of G and C influences the thermal stability of polynucleotides. Theoretical considerations of helix stability in polynucleotides are discussed in Sections X and XI.

C. Characterization of Nucleotide Material by Ultraviolet Absorption Spectra Nucleic acids generally possess an absorption band around 260 mp. This absorption band (which is determined by the purine and pyrimidine bases) is changed when the polynucleotides form secondary structures. The hypochromicity and hyperchromicity are functions of various en-

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ergy transitions arising from interactions among the bases. These interactions are weakened when the secondary structure is destroyed, and the absorption near ,A increases (hyperchromicity) . Hyperchromicity can be caused by increased temperatures, ionic changes, or radiation. Most frequently the thermal dissociation is studied. In the case of perfect alignment, as in DNA, an abrupt change in absorption occurs over a very narrow temperature range. This so-called “cooperative” effect shows that base pairs are not broken independently, but that an all-or-none process takes place; i.e., in a hydrogen-bonded helical complex containing homopolynucleotides, the entire chain is dissociated (by heat or changes in pH, for instance). The temperature a t which this fusion takes place is called the melting temperature or T,. It is generally defined as the temperature of the mid-point of the thermal transition. Felsenfeld and Sandeen (27) have introduced another definition

where ADi is the change in absorption in the temperature interval around Ti. This definition is particularly useful for cases with broad melting profiles. I n cases where the melting step occurs within a few degrees, T, and T%will be essentially equal. 1. DIFFERENCE SPECTRA It is now established that hyper- or hypochromicity a t a given wavelength is a function of the bases involved in the polynucleotide structure. It is for this reason that determinations of T,, till recently confined to measurements a t 260 mp, are now being extended over a wider range of wavelengths. To illustrate the possible differences as a function of composition, the difference spectra of the following reactions are shown in Figs. 1 and 2.

+ + + +

poly A * poly U + poly A p01y U poly A 2 poly U -+ poly A 2 poly U poly G . poly C + poly G poly C 2 p l y G poly C -P 2 p01y G p01y C

-

One can readily see that transitions involving complexes between A and U (128)are vastly different from those involving G and C (29), and further, that ,double- and triple-stranded transitions also present significant differences. Such differences have permitted the detailed study of thermal transitions of both complexes involving poly A and poly U. Difference spectra have also been used for the examination of secondary structure in nucleic acids (127,3e56). All these studies involve some method of fitting the difference spectra of A U and G C to a

+

+

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SYNTHETIC POLYNUCLEOTIDES I

I

I

I

I

I

E

E

4

2 x

W

2

0

1

1

,

1

240

1

1

260

1

280

,

30

FIQ. 1. Molar difference spectra for poly A . polyU and poly A * 2polyU between 20 and 85°C [after Massouli6 et d.(98)l.

r

I

I

240

I

I

I

260

200

1

300

A (mp)

FIG.2. Difference spectra (not on a molar basis) for poly G . poly C and 2 poly G poly C [after Pochon and Michelson (39)1.

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difference spectrum of a given nucleic acid. These investigations have given good results for DNA, the structure of which is known to be completely helical, and have given rise to suggestive results for various RNA’s for which the secondary structures are as yet unclear. The seemingly trivial question of the “strandedness” of RNA’s, known in the case of a few RNA’s to be double-stranded, apparently cannot be solved by difference spectra (36,36). 2. MIXINGCURVES(METHODOF CONTINUOUS VARIATION) This method, originally described by Job (37) permits the determination of the stoichiometry of complexes formed between two components.

Fraction U

FIG.3. Typical mixing curve at 260 mp between poly A and polyU in 0.2 M Na+ [after Massouli6 (In)].

It consists in studying a characteristic property as a function of the concentration of the two components A and B in the mixture. I n the case of polynucleotides the hyper- or hypochromism of the mixture is followed. I n practice, a series of mixtures is made in each of which the total concentration of both components is constant, and only the ratio [A]/[B] is changed. (As in most other instances, all concentrations refer to monomer units of the polymer.) A mixing curve is then obtained by plotting the change in absorption a t a given wavelength against the ratio of concentrations (Fig. 3). I n the case of no interaction between

SYNTHETIC POLYNUCLEOTIDES

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the two components the mixing curve becomes a straight line connecting the two points for the pure components. A sharp intersection between the two branches shows the stoichiometric mixture. Curved or flattened intersections indicate that the mixture is not thermodynamically equilibrated. Frequently such curved intersections sharpen considerably if the measurements are repeated some time later. Felsenfeld (38)investigated theoretical models for the interaction of linear polymers. His studies led to the conclusion that sharp intersections correspond to the geometrically (and thermodynamically) most favorable arrangement ; i.e., the maximum number of base pairs are formed. This implies automatically a “slipping” mechanism in which the chains already associated disintegrate and reform base pairs locally. This hypothesis has been confirmed by studies with oligonucleotides. “Mixing curves” were first used with polynucleotides to show the interaction of polyA and polyU to give a complex resembling DNA (39) and a second complex involving the binding of two polyU strands for each polyA strand ( 4 0 ) . Since then, the mixing curve has become a standard criterion for evaluating polynucleotide complexes. Recently the use of mixing curves has been refined by use not only of measurements near A,,, but also other, more characteristic wavelengths. It is thus possible to determine the formation of the two-stranded complex poly A poIy U a t 283.5 mp where the three-stranded complex shows neither hypo- nor hyperchromicity ( 4 l , 4 2 ) . a

3. CHARACTERIZATION OF NUCLEOTIDES AND POLYMERS BY ULTRAVIOLET ABSORPTIONSPECTRA

The fact that the various bases and their corresponding nucleosides and nucleotides possess different absorption spectra has long been used for the determination and characterization of nucleotide material. For summaries of spectra we refer to the work of Fox and Shugar (43, 44), of Voet et al. ( 4 5 ) ,and of Beaven, Holiday, and Johnson ( 4 6 ) .Recently, Vengstern and Bayev (47) have published a series of thirty-three spectra, including those of oligonucleotides a t various pH values. Kerr and Seraidarian (48) and Steiner (49) exploited the differences in absorbancy of nucleotides a t various pH values and used the absorbancy a t characteristic wavelengths to determine approximate base compositions in mixtures. Reid and Pratt (50) first used computers to resolve spectra of nucleic acid hydrolyeates into their components. A similar method, though less accurate, has been employed by Vasilenko et al. ( 6 1 ) . Recently, several attempts have been made to use ultraviolet absorption spectra for the deteflmination of base compositions of nucleic acids and

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even to determine certain nucleotide sequences. Lee et al. (5g) have used the least square method of Reid and Pratt (50) with a theoretical error estimation and claim to be able to resolve up to nine different components simultaneously. Guschlbauer et al. (53), using ultraviolet absorption spectra of polynucleotide and RNA hydrolyzates before and after UV irradiation (which changes the spectra of the pyrimidine nucleotides) , have been able to determine base compositions to an exactness of 1% or less. Pratt et al. (54) further developed the computational aspect by using linear programming to determine exactly the composition of oligonucleotides without having recourse to hydrolysis. D. Optical Rotatory Dispersion and. Circular Dichroism

Electromagnetic radiation, upon interaction with matter, can be absorbed, refracted, scattered, or diffracted. If polarized radiation, which is split into a left and a right circularly polarized component, interacts with a molecule no change will be observed if the component beams are transmitted with equal velocity. If the refractive indices of the medium for the left and the right circular polarized light of a given wavelength are different (only very small differences suffice), the two beams are transmitted with unequal velocities and are out of phase upon recombination after passage through the medium, and the plane of polarization is rotated. Thus optical rotation is observed if a medium transmits two circularly polarized component beams with unequal velocity. If the absorption of the left and right circular polarized light is also different, the recombined light beam, after passage through the medium, becomes elliptically polarized. This phenomenon is called “circular dichroism” and causes the so-called “Cotton effect” (Fig. 4). An extensive discussion of the interrelation of absorption, optical rotation, and circular dichroism will be found in the book of Kauzmann ( 5 5 ) . Biot (56) first observed (in 1817) that optical rotation depends on wavelength. The dispersion of the optical rotation was first stated mathematically by Drude in 1906 (67) in an empirical way:

where [a]is the specific rotation, K a constant, h the wavelength a t which rotation measurements were made, and hq the wavelength of the absorption maximum of the chromophore at which the dispersion curve shows zero rotation (Fig. 3). By using the [a]values outside the Cotton region one can determine A,, with reasonable accuracy. This is done by plotting

SYNTHETIC POLYNUCLEOTIDES

93

l / [ a ] against X2. If this plot does not yield a straight line, higher-term equations of the form

Optical rotatory dispersion (ORD) has been widely used for structure and absolute configuration determinations in organic chemistry [for a summary, see the book of Djerassi (68)1. Neither Eq. ( 5 ) nor (6) will, however, satisfy optical rotation changes caused by structural assymetry, such as secondary or tertiary structure.

FIG.4. Optical rotatory dispersion of poly BrU in 0.1 M NaCI, 0.05 M Na cacodylate, pH 7, 0.01 M MgClz at 2 and 20°C [after Michelson ( 1 1 ) l .

Moffitt and co-workers (59-61) have derived several empirical equations; the most widely used one is the following:

Here [m’]is the mean residue rotation, n the refractive index, M , the mean residue weight, and and &, two constants, a,, consisting of two terms, one depending on the intrinsic residue rotation, the other, as well as a,, depending on the helical configuration. It still must be kept in mind that this equation is empirical and numerous theoretical studies

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(62-64)have been undertaken to interpret the Moffitt equation, which has proven extremely useful in protein and polypeptide chemistry. I n recent years ORD measurements have been extended to polynucleotides and nucleic acids (66). The availability of several new instruments that record ORD curves automatically has produced a number of basic publications from the laboratories of Yang (66-68), Tinoco (69-70),and Zamecnik (71-73). As mentioned before, the differential absorption of left and right circular polarized light will give rise to circular dichroism. If this differential absorption can be determined, the rotational strength (RJ can then be calculated (74):

where EL and E R refer, respectively, to the molar extinction coefficients for left- and right-handed circularly polarized light a t frequency v ( h is Planck’s constant, c the velocity of light, and N Avogadro’s number). Circular dichroism was virtually unavailable until recently. Grosjean and Legrand (76) designed an electrooptical apparatus for the measurement of circular dichroism, and Holzwarth (76) has developed an adaptation for the Beckmann DK-2A spectrophotometer. The apparatus of Grosjean and Legrand (Roussel-Jouan dichrograph) as well as an adaptor for the Cary 14 spectrophotometer for circular dichroism measurements are now commercially available. Brahms (74)has described a series of pioneering investigations on the circular dichroism of polynucleotides and nucleic acids (77-84).

E. Titrimetry

I

The simplicity and reproducibility of spectrophotometric titration has resulted in a wide use of this technique for polynucleotides and their complexes, and for nucleic acids. The titration is performed in a spectrophotometer cuvette with a fitted glass electrode, a magnetic stirrer, and a thin tube connected to a milliliter syringe (86). Changes in absorbancy and in pH are recorded as a function of added titrant, thus providing a convenient method for the determination of apparent pK values. These are characteristic of the material and of the salt concentration.

F. Infrared Spectroscopy If the incident electromagnetic radiation has a wavelength in the micron range, its energy will be sufficient to cause vibrations in the molecular structure of a molecule. Infrared ( I R ) spectra measure the separation between vibrational energy levels. The theoretical background

95

SYNTHETIC POLYNUCLEOTIDES

of IR spectra can be found in the books of Tanford (19), Kauzmann (55),and Bellamy (86). I n polynucleotide chemistry, the use of IR spectra is rather limited, although they are very widely and routinely used in organic structure research. Apart from studies by Angel1 (87), Tsuboi et al. (88-90),and Miles (91-100) on bases, nucleosides, and nucleotides, Miles et al. have used this technique to study polynucleotide structures. Detailed discussion of their results is found in Section V, dealing with polynucleotide interactions. Many less popular techniques have also been utilized for the study of polynucleotides. Their lack of popularity is often due either to the fact that the equipment involved is very expensive or that extensive experience is needed to obtain meaningful results. Frequently a purity of material is required that is not easily obtainable from biological sources. However, the application of these techniques is growing as more problems in biochemistry necessitate novel approaches.

G. X-Ray Diffraction, Low-Angle X-Ray Scattering, and light Scattering These three techniques have many physical principles in common but the first two provide more precise information. Only a very short introduction to these techniques is given. A brilliantly written and frequently amusing review with examples from “everyday” Cambridge life for noninitiated members of the nucleic acid field has been written by Crick and Kendrew (102).Kendrew (105) has also presented the mathematical foundations of X-ray diffraction. Low-angle X-ray scattering has been reviewed in this series by Luzeati (104). Luzzati (106) and Kratky (106) have discussed the mathematical and technical details of this method. Further treatises on these subjects are those of Porod ( l o r ) , Witz (108), and Stokes (109). Light scattering has been extensively discussed by Tanford (19) and by Geiduschek and Holtzer (110). Any electromagnetic wave can be scattered, X-rays by the electron density of a crystal (or any other molecule for that matter), light by whole (and preferentially large) molecules. This can be deduced from s = 2 sin

e/x

(9)

Here s is the direction of the propagation of a scattered ray. For a sphere with radius r the intensity distribution of scattered radiation I ( s ) will be given by sin sr - sr cos sr

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It can be shown that, for all practical purposes, only measurements in the range where

rs = X

(11)

are useful. Since s is limited by definition [see Eq. (9)] to values equal to or smaller than 2/h, this means that the resolving power will be reached with r = X/2. This limits the applicability of X-ray scattering to the angstrom range, and that of light scattering to the range of thousands of angstroms. 1. X-RAYDIFFRACTION

If molecules are arranged in ordered patterns, as in a crystal, X-rays will be scattered by the electron density of the molecules and will be caught on the photographic plate in an ordered manner. This diffraction pattern is called the “reciprocal lattice.” Each spot will correspond to a wave in a wave analysis of an electron density (Fourier analysis). The position of every spot is indicative of the wavelength and the direction of the wave. Crystal patterns show ordered arrays of spots, which is not the case with powder or fiber diagrams. I n powder diffraction patterns, the totality of the small crystallites gives rise to a pattern of all crystallites together. They show sets of more-or-less sharp concentric rings, the radii of which correspond to the spacings of the principal lattice planes in the crystals. Fiber patterns are generally less perfect than crystal patterns. They can be thought of as a crystal that continuously revolves around one (the helix) axis. From fiber diagrams, one can generally obtain the crystallographic repeat unit along the fiber axis. The dimensions of the unit cell can only rarely be obtained. Frequently, this information, as well as other crystallographic parameters, can be obtained by model building, calculation of the X-ray diffraction pattern (Fourier synthesis) of the constructed model, and comparison with the pattern obtained. By this method, often all possibilities except one can be excluded. 2. LOW-ANGLE X-RAYSCATTERING

The distinction between X-ray diffraction and low-angle X-ray scattering is by no means arbitrary, but is based on the fact that pure liquids (and many crystals) show no X-ray pattern below an angle of about 10”. This is apparent from Eq. (9). For a value of B = 10” and X = 1.5 A, r will be about 4 A. This value of r will increase rapidly with decreasing 8 , thus leaving the region of atomic dimensions. On the other hand solutions of high molecular weight substances will show this low-

SYNTHETIC POLYNUCLEOTIDES

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angle X-ray pattern. This phenomenon, first discovered by Guinier (1111, has recently become very useful in the study of macromolecules. The low-angle X-ray pattern can be exploited to determine the radius of gyration of the macromolecule and, if measurements are made on an absolute basis, yields the mass per unit length, a parameter highly useful in determining the “strandedness” of a polymer helix (11.2). For the mathematical details of this difficult and demanding method we refer to the reviews by Luazati (105), Kratky (106), and Wita (108). 3. LIGHTSCATTERING

This technique, widely used for the determination of M , in protein chemistry, has shown relatively little value for polynucleotides. Extensive reviews are found in the paper by Geiduschek and Holtzer (110), and in Chapter 17 in the book of Tanford (19). Peterlin (113) has compared light scattering with low-angle X-ray scattering.

H. Nuclear Magnetic Resonance (NMR) and Electron Spin Resonance (ESR)

Both these techniques are of recent date and their application to polynucleotide studies is still very limited, partly because of the costly equipment involved. Jardetaky and Jardetzky (114) first used NMR to study basic resonance assignments of nucleosides, while Schweitzer et al. (116) studied various substituted purines. Jardetzky (116) and McDonald et al. (117) also have applied NMR to the study of the structure of poly A, poly U, poly C, poly I, poly A * poly U, and DNA. I n a recent study, McDonald et al. (118) used NMR for the investigation of the secondary structure of tRNA. They were able to distinguish between the rotational liberation of the bases and that of the ribose units during denaturation. ESR has been used mainly by Blumenfeld and by Rahn et al. (119, 119a) for the study of basic assignments of purines, pyrimidines, and nucleosides.

1. Electron Microscopy Technical improvements in electron microscopy have greatly increased the interest for investigations of macromolecular structure. Hall (120) has pioneered this technique in the nucleic acid field. Recently, Beer and his collaborators (121-124) investigated the possibility of selective labeling of individual bases in the polynucleotide chain with a view to sequence determination.

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J. Equilibrium Density Gradient Centrifugation Centrifugation in a density gradient of cesium chloride was introduced by Meselson, Stahl, and Vinograd (125) for the determination of the buoyant density of deoxynucleic acids. Doty, et d.have established (126-129) a linear relationship between G plus C content and buoyant density. The fact that polyribonucleotides band a t considerably higher buoyant densities has inhibited the use of this technique for the study of polyribonucleotides and their complexes. However there have been several studies on hybrids between deoxyribo- and ribopolynucleotides (130, 131).

IV. Homopolynucleotides The various random coil and structured forms of polynucleotides containing a single naturally occurring purine or pyrimidine base are discussed in this section. Under suitable conditions of pH or salt concentration, all such polymers form hydrogen-bonded, multistranded, secondary structures.

A. Polyguanylic Acid The highly marked tendency of guanosine residues to aggregate and form gels was long evident to those who worked chemically with guanosine and guanosine phosphates. However, the first reliable investigation producing clear-cut results was that of Gelled ‘et al. (132,133) who reinvestigated an observation made by Bang in 1910 on gel formation with guanylic acid (134),Formation of organized structures, with a T, z 12OC in concentrated solutions of G5’P and G3’P (but not of G2’P), a t pH 5 was followed by changes in UV absorption and optical rotation. X-ray diffraction studies of fibers drawn from the respective gels indicated a linear aggregate of stacked tetramers with rotation about the helix axis in the case of the 5’-phosphate. ORD studies of 5’-GMP gels have also been reported (135). I n view of the above remarks, i t is not surprising that oligoguanylic acids aggregate readily. Such effects have been described for chemically synthesized oligoriboguanylates and oligodeoxyguanylates, one of the most striking results being a very marked increase in pK value (136). More detailed studies of oligoriboguanylates showed that, whereas the aggregation is rather slow, the thermal stability (T, = 23OC for GpG in 1 M NaC1) is marked (1S7). Various alleged preparations of polyG have been described but i t appears that the only reliable preparation involves E . coli polynucleotide

SYNTHETIC POLYNUCLEOTIDES

99

phosphorylase in the presence ( 5 ) of Mn++ (instead of Mg++)a t 60°C. This polymer possesses a secondary structure with an extremely high stability, indicated both by the thermal resistance to transition to a random coil, and by the large shift in pK value (to 11.3) of the bases (29, 138). Whereas alkaline dissociation is abrupt and cooperative (characteristics of the transition, structure + nonstructure) , acidic titration is noncooperative and shows no indication of loss of secondary structure (29). Indeed, the optical rotatory dispersion curve remains virtually unchanged a t pH 2 (no precipitation occurs) (139) and even with a proton on each base the T, is > 100°C in 0.15 M NaCl a t pH 2.5 (I@). Unlike oligo G’s (or small poly G’s obtained by a brief alkaline treatment but liberating no oligonucleotides), which form very viscous solutions a t 7 mg/ml (possibly as a result of multistranded aggregation) , poly G does not show either extreme viscosity or a tendency to aggregate and precipitate at neutral pH, as has been reported for an alleged poly G (141). There is a t present no evidence indicating the number of strands involved in the secondary structure. It is probable that poly dG also has a very strong secondary structure, which inhibits action as a template for RNA or DNA polymerase. Whereas poly d C is active with both these enzymes, poly dG is not (14.2,

-

143)

B. Polyinosinic Acid P o l y I forms a secondary structure that probably contains three strands with an interbase distance of 3.4 A (14.4). As with other helical structures containing uncharged bases, the T, increases as a linear function of the logarithm of the salt concentration. Poly dI is slightly more stable than poly I (T, = 50°C compared with 42°C in M NaCI) (146). The increase in thermal stability for a tenfold increase in salt concentration is 31.5”C1 compared with a typical double-stranded complex such as poly dI * poly dC, which shows only a 17°C increase (146). This suggests that poly dI, like poly I, is a triple-stranded complex, the effect of salt being presumably more marked with the higher concentration of negative charges present in a triple compared with a double helix (Fig. 5). The ORD of polyI is exceptional in that the general appearance is reversed: two troughs and one peak are present between 240 and 300 mp instead of two peaks and one trough shown by all other polynucleotides (and nucleic acids) examined (141). Formation of a complex with poly C or poly A inverts the ORD profile. Since an inverse ORD profile does not necessarily indicate a change in the “handedness” of the helices, this may be attributed to the difference in base interactions (141).

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01

I

-3

GUSCHLBAUER

I

I

-2

w.

-I

1

0

tl

log [Nd]

FIG.5. Variation of T, with ionic strength of various two-stranded complexes.

Symbol 0-0

(>-Q

Q-Q

+-+ x-x

A-A

v-v 0-0

0-0 A

rG . dC

D

rG rC

v

Complex

References

dG . dC dI .dBrC dI . dC dI * dI * dI rI rI rI rI . dC rI rC dI . rC rI rMeC

199, 869 199,807 199,206 146,806 606,868 606 806 606

-

888

+ EDTA

867

SYNTHETIC POLYNUCLEOTIDES

101

C. Polyuridylic Acid Secondary structure in poly U is apparent a t low temperatures (147, 148, 150). The formation of this structure can be followed not only by changes in ultraviolet absorption, but also by the variation of the Cotton effects (149) with temperature, and by UV circular dichroism ( 7 8 ) . Nevertheless, nothing is known about the number of strands in the structure or whether (if a double-stranded structure is formed) the strands are parallel or antiparallel. It is considered that polyU is completely devoid of any kind of organized structure a t temperatures above 15°C (148). However, the same authors found no anomalous titration behavior. This is not the case; polyU shows a very marked variation of pK as a function of ionic strength as well as an anomalous ORD. Recent work indicates the presence of considerable base stacking in singlestranded poly U (150).

D. Polyadenylic Acid Titration of polyA shows a very marked shift in pK value of the bases to a higher value. In acid solution (pH 5) , poly A possesses hydrodynamic properties that are characteristic of a rigid molecule (151), and ultraviolet absorption-temperature profiles indicate a cooperative loss of secondary structure over a narrow temperature range (152-155). The stability of the structured form is greater the lower the pH or concentration of salt. Decrease in salt concentration increases the apparent pK value and hence the pH difference between the pK and the ambient pH (Fig. 6). The T, is a direct function of this difference. Crystallographic studies indicate a double-stranded structure with parallel chains and an interplanar distance of 3.8 A. Hydrogen bonds occur between the 6-amino group and N7 of the bases; the proton a t N1 is not involved but probably stabilizes the double helix by electrostatic interaction with phosphate groups (156, 157). Formation of the “acid” form of poly A has also been followed by small-angle X-ray scattering techniques. The rodlike molecules have a mass per unit length similar to that in fibers of DNA (158).As might be expected, significant differences in optical rotatory dispersion and viscosity, as well as ultraviolet absorption, can be seen for the transition between the molccule a t neutrality and the double-helical “acid” form of polyA (151). I n neutral solution, poly A shows hydrodynamic properties characteristic of B random coil (42, 69, 74, 80, 159, 158). Whereas in doublestranded secondary structures involving the amino group, the bases are protected from attack by formaldehyde, the neutral form of poly A read-

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A . M . MICHELSON, J .

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AND

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PY

FIG.6. Variation of T, with pH of homopolymer complexes at various ionic strengths. Symbol

+-+ x-x

v

A-A

A-A

v-v .-a

0-0 0-0

Complex

“a+]

References

dC * dC dC . dC dBrC . dBrC rC . rC rC ’ rC rMeC . rMeC rA . rA rA ’ rA rA . rA

0.1 M

146 268

~

0.4M 0.4M 0.15 M 0.1 M 0.15 M 0.03 M 0.15 M 0.5 M

268 268 167 260

166 166 166

SYNTHETIC POLYNUCLEOTIDES

103

ily reacts with this reagent (159).Nevertheless, a variety of optical properties such as the variation of ultraviolet absorption with temperature (159),the optical rotatory dispersion (69,160, 16l>,and circular dichroism (7‘8, 83) indicate, as was previously demonstrated with oligonucleotides, that there is a marked interaction (169-164) among successive bases in the chain leading to a single-stranded, stacked, helical structure. The same conclusion has also been drawn from NMR studies of poly A (117).Small-angle X-ray scattering techniques also indicate, over short regions, a rodlike structure with a mass per unit length corresponding to one nucleotide per 3.5 A (158).Nevertheless, the structure, as shown by the hydrodynamic properties, is nonrigid (152).The disorganization of structure with increase in temperature is noncooperative (49,69,159,159) and is probably independent of hydrogen-bond formation between adenine bases (hydrogen-bonded water molecules are, of course, present). Further, the transition appears to be independent of the salt concentration (159).The structure is random with respect to total conformation, but ordered in terms of short-range interactions (158). The thermodynamic analysis of such a conformation has received attention lately (49,169, 163,164),since an evaluation of the “stacking” energy can be thus obtained. The standard free-energy change obtained for single-stranded oligo- and polyadenylic acid a t neutral p H and 0°C is about 1 kcal/mole in favor of base stacking, about one in eight of the bases being unstacked (83) a t this temperature, thus giving considerable flexibility to the molecule (159,162). At 20°C, about two thirds of the bases are in the stacked conformation a t any given instant. [See also the paper by Ts’O, Helmkamp, and Sander (165)who arrive at a similar proportion of structural interaction by less refined techniques.]

E. Polycytidylic Acid As with poly A, an “acid” double-helical form of poly C can be obtained (165,166). However, the structure is somewhat different. Studies of thermal stability as a function of p H and ionic strength of the medium suggested that the system of hydrogen bonds involves a shared proton between each pair of cytosine bases (167).X-ray diffraction studies of fibers of polyC drawn from acidic solution led to the same conclusion (168).The chains are parallel with an interplanar distance along the helix axis of 3.11 A. The thermal transition can be followed by changes in the ultraviolet absorption and in optical rotation (169).The corresponding structure is also formed by polydC and, a s in the case of the polyribonucleotidc, the thermal stability of the secondary structure is greater the lower the pH and the ionic strength of the medium (145).

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Again there is an uptake of one proton for two cytosine residues on passing from the neutral form to the ordered helical structure. It may be noted that the pH of formation in a given solvent is higher than that for polyrC and, in conformity with this, the thermal transition occurs a t higher temperatures than with polyrC (respective pK’s in 0.05 M Na+ are 5.8 and 7.4) (Fig. 6). The structure of polyC a t neutral pH has also been examined. As with poly A, heating an aqueous solution a t pH 7 produces a noncooperative continuous increase in ultraviolet absorption (166) in the region of A,,,. Circular dichroism studies suggested that a t neutral pH poly C possesses a helical conformation ( 7 4 ) . Optical rotatory dispersion studies were also claimed t o prove the presence of a highly ordered secondary structure a t pH 7 (169). Since similar results were obtained with formaldehyde-treated poly C, hydrogen bonds were considered to be eliminated. I n fact, treatment with formaldehyde does not necessarily destroy hydrogen-bond-forming capacity. As suggested in earlier publications, it was considered that base stacking could contribute to hypochromism. The statement that p l y C a t p H 7 exists as a highly ordered asymmetric structure is erroneous if taken in a hydrodynamic sense and the conclusion that so-called hydrophobic forces are the predominant stabilizing factor is, rather strangely, immediately contradicted in the sentence following this claim when it is stated that hydrophobic forces are less effective a t lower temperatures. If this were so, then clearly the postulated helical structure should be more stable a t high temperatures than a t low, contrary to experience. It was also inferred that the helical structure is not necessarily completely rigid, but an interrupted helix made up of smaller helical regions. This static concept is untenable and, as in the case of poly A, one must consider poly C to be a random coil with continuously fluctuating regions of helicity arising from base-base interactions in constant change.

V. Polynucleotide Complexes A. Complexes between Poly G and Poly C Although the interaction of polyA and polyU has been studied in detail for some considerable time, studies on the formation of a complex between poly G and poly C have only recently become possible. Much of the earlier work relates to complex formation between oligo G’s (or short polydisperse poly G) (170) and poly C. Lipsett has examined the interaction of GG and GGG with poly C (171, 172). At neutral pH, a complex with the stoichiometry 1G:lC is rapidly formed, followed by slower formation of 2G: 1C. However a t slightly acidic pH ( A pH 6 ) a

105

SYNTHETIC POLYNUCLEOTIDES

third complex, probably containing a t least some protonated C residues, is formed with a ratio 1G:2C. The interaction of poly G (as opposed to oligo G ) and poly C gives somewhat different results (29). Mixing curves under various conditions and a t various wavelengths indicated forination of a 1:l coinplex only, with a stability greater than that of poly G, as shown by increased thermal stability and also by alkaline dissociation, which occurs cooperatively a t pH 12.27 for poly G * poly C in 0.15 M NaCl (compared with 11.43 for poly G). I n order to achieve separation of the strands a t neutral pH, 80% methanol was used, giving a T, of 89°C. Under these conditions, the dissociation is irreversible, until the methanol is removed. (The complex with poly BrC is even more stable.) The triple complex, 2G:lC, could not be obtained. However, when the polyG was briefly treated with alkali to reduce the molecular weight, the complex 2G:lC could be readily obtained. Similarly mixing curves of poly G with short poly C (average chain length z 15) indicate formation of a 2 G : l C complex. A possible explanation is simply that, with two extremely long polymers, rigidity of a double-helical structure prevents addition of a third strand. However, if breaks are present in one or the other of the two strands, flexibility is increased and the addition of a third strand is facilitated. Another approach to the preparation of poly G poly C has been described (173, 174). Poly C was used as template for RNA polymerase (Micrococcus lysodeikticus) in the presence of G T P to give a product, apparently double-stranded, containing 1G :1C. Other than that the polyG formed appears to consist of short strands, the properties of this complex resemble those of the nonenzymatic preparation. The alkaline dissociation followed by neutralization yields a complex distinguishable from the original, possibly due to formation of aggregates containing 2G:lC (173). The optical rotatory dispersion of the above preparation of poly G poly C has been described (149). Somewhat surprisingly, a sharp transition occurs a t 90°C in 0.1 M Tris, with irreversible changes in magnitude. Similar changes were also observed in the UV absorption a t 90°C. In view of these results, serious doubts must be cast on the integrity of the material used, particularly since the same authors state that poly G is highly insoluble in aqueous solution, a statement that is as surprising as it is incorrect. ORD studies with the complex obtained by interaction between the two homopolynucleotides confirm an extreme stability and the complex exists in a range of pH between 2.0 and 12.2 (139). Other studies have shown that, a t pH 2.5, poly G-poly C is fully protonated (two protons

-

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per base pair) yet nevertheless has a T, of approximately 8OoC ( 1 4 0 ) . Hence acidification does not destroy the complex, in contrast with the alkaline dissociation. This can readily be explained in terms of the base pairing characteristics. As might be expected, thermal dissociation of poly Gepoly C a t pH 2.5 is irreversible. At this pH both poly G and poly C possess structures much more stable than that of poly G-poly C. Complexes of poly G with poly BrC (or with poly iodo C) a t p H 2.5 are even more stable than is poly G-poly C. The complex of polydC and polydG can be obtained from dCTP and dGTP by means of DNA polymerase. The product generally contains 5 0 4 0 % of dG (that is, an excess) and shows a biphasic melting curve, the first step of which corresponds to the melting of polydC ( 1 4 6 ) . Alkaline dissociation is reversible with respect to ultraviolet absorption, but the viscosity does not return to the original value (175). It may be noted that poly dC.poly dG is markedly less stable than the corresponding ribonucleotide complex. Circular dichroic spectra (81, 84) of poly dCapoly dG are inverted by comparison with those of DNA or poly A-poly U. This has led to the perhaps unjustified suggestion that the structure is a left-handed double helix rather than right-handed (176). It would be of interest to have dichroic spectra of poly I (structured form).

B. Complexes between Poly A and Poly U The interaction of poly A and poly U to form helical complexes has been the most extensively studied of this type of reaction. The polymers can be prepared readily, and they react spontaneously in saline solution (177). It was thought a t first that only one kind of association could arise, a double-helical structure in which adenine and uracil would be paired like adenine and thymine are in DNA, and indeed it was shown that the complex between poly A and poly U has the crystallographic properties of a helix, the dimensions of which are very close to those of DNA (178-180). Moreover, the viscosity of the solution increases during formation of the complex and the UV absorption at, , ,A decreascs while the optical rotatory power increases (177, 181). The hydrodynamic and optical properties of the complex resemble those of DNA. The stoichiometry of the association was studied by the continuous variation method (39, 40). An equivalent number of A and U residues entered the complex, which was therefore termed poly A-poly U. The method of continuous variation also showed that a second complex, containing two U residues for one A, was found when an appropriate mixture of polyA and polyU was allowed to react for a longer time, The reaction is rapid in the presence of divalent cations, or in a high enough

107

SYNTHETIC POLYNUCLEOTIDES

(>0.1) ionic strength, but it occurs even with low salt concentration (0.005 M Na’). It is thought that the two uracils are bound to the adenine in polyA-2polyU and not to each other. One of them is presumably in an orthodox Watson-Crick pairing, while the other is hydrogen-bonded through the remaining hydrogen of the 6-amino group and N7 of the adenine (98). This can be achieved in two ways, as is discussed below. Since the properties of poly A-poly U are of importance for an estimation of the contribution of one kind of base pair to the corresponding properties of DNA, the two complexes poly A poly U and poly A - 2 poly U received intensive study. Considerable confusion about the equilibrium of the complexes arose as a result of a coincidence in certain optical properties of the two species (182). The hypochromicity a t 259 mp does not distinguish between the following possibilities :

-

(1) A lA/lU mixture first gives rise to poly A*polyU which a t equilibrium rearranges itself into poly A.2 poly U liberating one half of the poly A. This process would lead to the same complex, poly A-2 poly U, a t equilibrium in all mixtures, since i t is the stable form of the 1A/2U mixture (40, 183). (2) Poly A-polyU is the stable complex formed in a lA/lU mixture. Such was generally assumed to be the case. The second case has now been proved by a variety of techniques such as NMR (116) and infrared spectroscopy (99) using concentrated solutions, and, in dilute solution M in polymer phosphate), by ultraviolet absorption a t wavelengths other than 259 mp, either a t discreet wavelengths (41, 42, 187) or by considering whole difference spectra (28, 187), by the use of formaldehyde to measure noncomplexed poly A (184, 187), and by the stimulation of ethidium bromide fluorescence by double- (but not single- or triple-) stranded structures (186). It is particularly interesting to consider ultraviolet absorption a t two “selective” wavelengths, X1 and Xz. X,I a wavelength close to 280 mp, is such that poly A * poly U shows no hypo or hyperchromicity; i t is not distinguished from its unreacted components, and poly A * 2 poly U is the only complex shown by the continuous variation technique a t this wavelength. At Xz, which is close to 283 mp, only poly A * poly U can be seen; i t is hyperchromic a t this wavelength (41, 42) (Fig. 1 ) . By considering simultaneously two “mixing curves” a t X1 and A?, one can determine the amount of poly A * poly U and poly A 2 poly U in any mixture (41). This leads to a rather odd conclusion regarding the evolution of a lA/lU mixture (186). It is true that poly A * poly U is formed rapidly, but in high enough salt concentration the formation of

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poly A * 2 poly U is also rapid enough to take place during the association of the two strands of poly A and poly U. One can visualize this in the following manner: if the strands are not yet perfectly matched, there must be some free ends of A and some free ends of U. The latter will then tend to engage themselves in three-stranded segments-perhaps by folding back on the two-stranded structure. As a result of this, 1 or 2 hours after mixing, in 0.2 M Na+ solution, the totality of U residues has entered the complexes, but approximately 10% of them are in poly A 2 poly U. A slower rearrangement takes place, eliminating this, and eventually one finds only poly A poly U. This requires that the two poly U strands be antiparallel. Alternatively, the slow reaction involves poly A * poly 2 U formed by addition of a second strand of poly U to poly A * poly U. In this case, the poly U strands can be parallel or antiparallel.

-

-

C. Thermal Dissociation of Poly A Poly U and Poly A 2 Poly U (41,@1 187) By following the variation of ultraviolet absorption a t 259 mp, one can demonstrate the cooperative dissociation of poly A * poly U or poly A * 2 poly U a t the dissociation temperature, Tm.But simple dissociation, leading directly to free poly A and poly U, is not the only transition observed. For instance, a t low salt concentration (< 0.1 M Na+) and in the absence of divalent cations, poly A 2 poly U is dissociated in two steps:

-

poly A 2 poly U + poly A poly U 9

+ poly U + poly A + 2 poly U

It is necessary therefore to define several kinds of dissociation temperatures. The first one will be noted T m ( S - z ) , and the second one Tm(z-l), the figures designating the number of strands associated below and above the T,. I n low salt concentration, poly A poly U melts a t Tm(z-l), and between T,(3-2) and Tm(z-l)there is a temperature range where this complex is the only stable one. The absorption spectra of the lA/lU mixture and poly A poly U in the two states, poly A * 2 poly U below Tm(s-z) poly U between T,(3-z) and Tmcz-l,, intersect with that of the dissociated a t the selective wavelengths X1 and Xz. This is polymers (above Tmc2-1)) a simple way to demonstrate the existence of XI and Xz and to determine their values. Hence in low salt concentration one must consider two transition temperatures, T m ( 3 - 2 ) and Tnz(2-1,. Above 0.1 M (Na+), the corresponding transitions do not occur, and poly A * 2 poly U dissociates directly into the component polymers a t a temperature noted Tm(3-1).In contrast t o the situation in (Na+) less than 0.1 M , where the dissociation of poly A-poly U is direct and that of poly

-

-

+

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SYNTHETIC POLYNUCLEOTIDES

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A 2 poly U givcs poly A . poly U, we find in higher salt concentrations that poly A * poly U rearranges itself into poly A - 2 poly U, which in This transformation is reversible and occurs turn dissociates a t Tm(3-1).

-

This transition: at a transition temperature Tm(Z-3). 2[p0ly A * poly

U]

poly A . 2 p01y U

Tm(I-:)

+ poly A

was first demonstrated by infrared spectroscopy (100) and later by ultraviolet spectroscopy (41, 42). If the dissociation of poly A * poly U were direct, the absorption a t X1 would not vary during increase in temperature. One actually sees two equal and opposite variations a t Tm(2-3) and Tm(3-1).

70

30

10 log [No+]

FIG.7. Experimental phase diagram of interactions between poly A and poly U (41, 42, 187).

The variation of the four transition temperatures, Tm(3-2), Tm(2--l), and Tm(3--1), has been studied as a function of the salt concentration in neutral solution. It may be noted that the dissociation temperature of DNA varies linearly with the logarithm of the salt concentration. I n the same ivay, T m ( 2 - 1 ) , Tm(3--2),and Tm(3--1)increase with log (Na+) and their variation is perfectly linear, within the limits of experimental accuracy. On the contrary, Tm(2-3) decreases and this decrease is not linear. The stability diagram contains four lines, which do not intersect because, even though the transitions are cooperative, they are not perfectly sharp, and each T, defines a transition zone rather than a boundary. However, these four lines divide the plot into four regions (Fig. 7): Tm(2--3),

110

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A. M. MICHELSON, J . MASSOULI~ AND

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I corresponds to the situation found a t low temperature; both complexes exist, depending on the ratio of A and U residues. ( b ) Region 11, below 0.1 M (Na+), and a t temperatures intermediate between Tmc3-2, and Tm(P-l), is a domain of exclusive stability of poly A poly U, except perhaps a t high U/A ratios. (c) Region 111, above 0.1 M (Na+),and also a t intermediate temperatures, is a domain of exclusive stability of poly A 2 poly U. ( d ) I n region IV, the polymers are not associated.

(.a) Region

-

-

This diagram is valid in a certain range of pH around pH 7.

D. Stability of Poly A Poly U and Poly A 2 Poly U at Alkaline pH Poly A-2 polyU dissociates in two steps upon titration with alkali a t p H 10 and 10.4 (0.2 M Na+) (181).At alkaline pH, the uracil bases lose the protons involved in hydrogen bonding with adenine and therefore ionization and dissociation of the complexes are linked phenomena. The stability of poly A 2 poly U is diminished more than that of poly A . poly U. T m ( 2 - 3 ) is not changed in this p H range because the corresponding transition does not involve a variation of the number of free U residues. But decreases and, above a p H value that depends upon the salt concentration, the process of dissociation of the complexes goes through Tm(3-z) and Tmcz-l), even a t high salt concentration (187, 188).

E. Stability of Poly A Poly U and Poly A 2 Poly U at Acid pH (187, 188)

At acid pH, polyA forms a helical protonated structure, and this enters into competition with poly A poly U and poly A 2 poly U. I n the acid form, poly A reacts very slowly with poly U (161), so that dissociation of the complexes appears irreversible, and acid titration shows a hysteresis loop (189). The stability of the complexes can nevertheless be studied by first allowing association to take place a t pH 7, then bringing the p H down to a desired value. As a result of competition with the helical form of poly A, Tm(z-3), Tmcs-l), and Tmcz-l) decrease a t pH's such that the T,,, of helical polyA is sufficiently high. (It should be rememis not changed, bered that this T , increases as the pH decreases.) Tmcs-,) because no polyA is liberated a t this transition. In low salt (0.03M Na+) the situation is rather complicated. Below pH 5.4, one finds only poly A * 2 poly U in all mixtures a t room temperature. Between p H 5.4 and pH 5, the dissociation of this complex is still in two steps, proceeding through poly A poly U. However, below pH 5, the poly A 2 poly

-

-

-

-

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SYNTHETIC POLYNUCLEOTIDES

U dissociates directly, as shown by a comparison of the absorption variations that occur a t the different transitions.

F. Base Pairing in Poly A Poly U and Poly A 2 Poly U The existence of polyA * 2 poly U clearly demonstrates that adenine and uracil can be associated in at least two ways. B y studying the increase of the carbonyl stretching frequencies of the uracil bases in poly A * poly U and poly A 2 poly U, Miles (98) has found that two different carbonyl groups must be involved in the hydrogen bonding present in the latter complex. His observations support the conclusion that poly A * polyU has a Watson-Crick type of bonding (190), while in poly A * 2 poly U, the second U is hydrogen-bonded through the C2 carbonyl to the 6-amino group of adenine. This differs from the bonding pattern originally suggested but is in accord with that observed in mixed base crystals (191). Both strands of poly U would be antiparallel to poly A if the same configuration of the glycosyl linkage is realized in all the chains. (Configuration here refers to rotation of the base about the glycosyl linkage.)

-

G. Thermodynamics of the Interaction between Poly A and Poly U One can give a quantitative explanation of the stability diagram presented in Fig. 7 (187) with the very simple formula,

AG

=

A(Tm- T )

where AG is the difference of free enthalpy between the separated polymers and the complex. Jf AH and A S were independent of temperature, A would be equal to AS, but it has been shown experimentally that this is not the case (162, 163, 192). By introducing the variation of T, with log (Na+) in the formulation of AG, AG can be expressed as a function of T and (Na+). This can be done for poly A * poly U and poly A 2 poly U, and involves two differentlA constants. The ratio of these constants can be determined from the relationship that is realized a t Tm(2--3). It is then possible to deduce the existence of the fourth T,, Tmc-3-2),and to calculate its value (Fig. 8). It can be seen that this very simple and even naive formulation is justified a posteriori: i t gives a rather good representation of the different equilibria a t the T,’s. A similar expression, in which p H is introduced, can also be written to define the equilibria among poly A poly U, poly A 2 poly U, the helical form of poly A, and the separated polymers a t acid pH. However,

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in this case it is not possible to give a quantitative prediction of the variation of the Tm’s. Experimental determinations of the heat of reaction of poly A and polyU show that AG increases with temperature (165). This variation is certainly due principally to changes in the properties of poly A. I n this polymer, the nucleotide residues show interactions that are noncooperatively disorganized by increase in temperature. This introduces a temperature-variable term in AG. Attempts to obtain a “true” AG by subtraction of this contribution have been made (4SJ 163),but i t is still doubtful if a reliable estimation has been obtained.

log [No*]

FIQ.8. Calculated phase diagram of interactions between poly A and poly U: 0,experimental values (187).

---, calculated;

The value AGA obtained for one A residue of poly A compared to the temperature of mid-variation in the properties of poly A seems to imply a rather high value for the corresponding ASA. The same order of magnitude as that of A8 for the formation of a rigid pair of bases is found. Moreover, AG for the reaction,

also varies somewhat with temperature (162) and the variation of AG for the reaction, poly A

+ p01y U

-+

poly A * p01y U

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SYNTHETIC POLYNUCLEOTIDES

with temperature is not the same a t different salt concentrations (1931, whereas the properties of polyA do not seem to depend upon the salt concentration, but only upon temperature (159). The simplified expressions used above are also in contradiction with the experimentally determined variation of AG with salt concentration. Instead of an increase in A G with increasing (Na'), the contrary is observed. Obviously, compensating factors are present and perhaps concomitant variations of AS and AG can explain this difficulty (194). However, the experimental decrease of A G when (Na') increases is difficult to understand. A priori, high ionic strength stabilizes the helical associations by shielding the negative charges of the phosphates and diminishing their repulsion. Thus poly A 2 poly U is more sensitive to variation of ionic strength than is poly A poly U. A tenfold variation of [Na'] corresponds to

-

=

21"

and

-

AT,(a-l, = 25"

By analyzing the dissociations of the complexes brought about by ionization of polyU a t alkaline pH, Warner et al. (181) were able to estimate indirectly AS and AG for the rupture of one pair or one triplet of bases.

H. Alternating Copolymers We here consider ribo- and deoxyribopolynucleotides containing alternating sequences, e.g., poly (rA-rU) , poly (dA-dU), and poly (dA-dT) . All these polymers possess an organized helical structure (195-197). This structure is like that of DNA, a double-stranded helix, and the two strands are antiparallel. The crystallographic properties of the lithium salt of poly (dA-dT) are identical to those of DNA in the B form. However the sodium salt differs from the known forms of DNA (198). Since each strand possesses a periodical self-complementary sequence, it can fold back over itself. Branched helices can thus be formed, and have been observed with the electron microscope (195). This particular behavior is probably the cause of several distinct properties of these alternated polymers. Instead of rising continuously during the thermal dissociation, the viscosity of a solution passes through a minimum a t a temperature slightly below the T, (T, being defined by changes in the ultraviolet absorption). Upon cooling, the absorption returns to the original value, but the viscosity does not. ( b ) The breadth of the thermal transition (again defined by the ultraviolet absorption) increases with the salt concentration. The reverse

(6)

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-

variation is observed for poly A poly U, poly A 2 poly U and for DNA. In these cases, it may reflect the heterogeneity in length for poly A and poly U, or in composition for DNA. The breadth of the thermal dissociation of poly dG * poly dC does not vary with the ionic strength. I

I

I

I

log

I

"a+]

Fro. 9. Variation of T, with ionic strength of alternating copolymers containing A, U, T, and BrU. Symbol

Complex

References

@-a

poly r(A-BrU) poly d(A-BrU) poly r(A-U) poly A * poly U poly d(A-U) POIY d(A-T)

200 196 200 41, 42, 86 200 196, 200

c)-@

0-0

0-0 0-0

0-0

The T,'s of the dissociation of poly d(A-U) and poly r(A-U) increase linearly with log (Na') ; A T , is equal to 21" for a tenfold variation of (Na+), exactly as for Tm(z-l) in the case of polyA poly U. The effect of ionic strength (Figs. 9 and 10) is therefore the same on all these two-stranded helices containing A-U base pairs. It is also nearly the same for other two-stranded helices: poly dI * poly dC, 17°C; poly dG poly dC, 18OC; poly dI poly dBrC, 20°C (199). The polyr(A-U) helical form is 10°C more stable than poly A poly U (187); it is also

-

-

-

115

SYNTHETIC POLYNUCLEOTIDES

10°C more stable tlian the helical poly d(A-U), the T , of which coincides of poly A poly U. with Tm(2-1, An important difference may be noted between polyr(A-U) and poly d(A-U). The relative variation of absorption a t 260 nip is much

-

a-a x-x v-v A-A

+-+

rA . rU rA . rT dA . dT rA . dT dA . rU

41, 48, 86 21 8,222, 268,267 129, 266 266 266

larger for polyr(A-U) (70%) than for polyd(A-U) (45%) (600).The value for poly r (A-U) is not abnormal when compared to that for poly A poly U which is 61% [between 20 and 85°C (68)1. The differences between the deoxy and the ribo alternating polymers, +

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also observed between poly d (A-BrU) and poly r (A-BrU) (200), could arise from slightly different helical structures. It is unlikely that the unstructured chains should have very different optical properties, though this remains a possibility.

1. Association of Poly A and Poly I Polyinosinic acid is generally considered as equivalent to poly G. It does indeed associate with poly C, as one would expect, but it also associates with poly A. Both a two-stranded and a three-stranded complex can be formed, polyA * polyI and polyA * 2 poly I (201). In this respect, polyI behaves as an analog of poly U. However, the complexes between polyA and polyI dissociate a t lower temperatures than the corresponding A and U complexes (146). Poly A 2 poly I does not seem to be formed a t all a t low ionic strength (0.01 M Na+) (20201). Although this system has been but little investigated (202),i t seems from continuous variation curves that poly A 2 poly I in the conditions studied (0.05 M Na+, pH 6.8) is the equilibrium form even in a l A / l I mixture. No data are available yet to construct a stability diagram, similar to that described for poly A * poly U, and the effect of p H on the stability of the complexes is not known. The hydrogen-bonding pattern of poly A 2 poly I is probably similar to that of poly A 2 poly U (98). Here, however, the same acceptor and donor groups of inosine have to be used for both polyI strands. It follows that, if they have, as is most likely, the same configuration of the glycosyl linkage with respect to base rotation, these strands must be antiparallel.

-

J. Associations Involving Hypoxanthine and Cytosine In neutral saline solutions, poly I and poly C associate into a doublestranded helical complex, poly I * poly C (2Q3).No triple-stranded structure has been detected. Poly I poly C has a hydrogen-bonding pattern homologous to that of G and C, but it cannot be considered entirely analogous to poly G * poly C since the T, (60.2"C in 0.15 Na') is much

-

lower (146). This T, increases linearly with log (Na+) (204, 206) (Fig. 5). The acid titration of poly I poly C is complex (206). Poly C forms a semiprotonated helical structure in acid solution but even a t pH's such that this poly (C, C') melts higher than poly C * poly I, the complex is not dissociated. In contrast with the effect of decrease in p H on poly A poly U, the T, of poly I poly C rises and the complex is protonated. In an ionic strength of 0.05 M Na+, the titration presents two cooperative steps. The first, a t pH 4.9, is reversible; the second, a t p H 3.7, shows hysteresis and the reverse titration takes place a t p H 4. Analysis of the complex peak, isolated from various mixtures of poly I

-

-

117

SYNTHETIC POLYNUCLEOTIDES

and polyC by sucrose gradient centrifugation, shows that the ratio of I to C is always 1. Poly (C, C + ) ,the acid foEm of poly C, does not react appreciably with poly I a t pH 6. At neutral pH, the T,, of poly I * poly C (ionic strength 0.05 M ) is 5O"C, a t pH 4.6 (where half of the cytosines are protonated) it is 53"C, and a t pH 3 (cytosines fully protonated) it is 62°C (no titration of poly I occurs in this p H range). It would be interesting to know whether

FIQ.11. Possible types of base pairing between inosine and cytidine. The neutral form is that found in poly I * poly C at neutral pH. Forms A-C represent three possible configurations involving protonated cytidine residues. The solid circles represent the position of the carbon atom (Cl') of the ribose residue [after Giannoni and Rich (906)l. the increase of T, with decreasing p H occurs as a regular variation or, as seems possible, in more or less abrupt steps, a t the pH's of protonation. Three hydrogen-bonding schemes have been considered for poly I polyC+ (Fig. 11). One of them, A, has been eliminated because the slow rate of reaction of formaldehyde with poly I poly C+ implies that the amino groups must be involved in hydrogen bonding. I n C, the cytosine is rotated about the glycosyl linkage as compared to the neutral poly I poly C pairing. This means either an inversion of the direction of the C strand, which is impossible without dissociation of p o ly I * poly

-

-

-

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A. M . MICHELSON, J. MASSOULI~ AND

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C and therefore formation of poly (C, C+), or else a rotation of the base in the double helix which also presents difficulties. This leaves model B for poly I poly C+. The structure a t pH 4.6 is well-defined, different from both the fully protonated and the neutral forms, as indicated by the cooperativeness with which it loses or gains one proton for two cytosines. The assumption has been made that it is an incompletely protonated polyI poly C. It must be an original structure in which two kinds of base pairs coexist, perhaps alternating. A completely different possibility is that the bases are arranged in triplets, poly I * poly C * poly C+ (Fig. 12). A structure of this type would explain why poly (C, C+) and polyI can associate a t pH 4.6. The authors do not seem to have checked the stoichiometry of the complex in this state, but only at

-

-

FIG. 12. Possible type of base interaction between poly I, poly C, and poly C'.

pH 3. However, they report a higher degree of association a t pH 4.6 than at neutral pH or pH 3 and they suggest that this may arise from the combination of individual molecules of protonated poly C combined with both polyI and polyc. However, a random process does not account for the cooperative nature of the transition. Poly dI polydC has a lower thermal stability (205) than poly I * poly C; the T, is 14°C lower (in 0.1 M Na+). The relative variation of absorption that takes place a t 246 mp during dissociation is also less than for poly I poly C, 52% compared to 67%. In this series we now find, besides the double-stranded structure poly dC poly dI, a triple-stranded one, 2 poly dI * poly dC (207). It is difficult to establish a stability diagram in this case because two more structures complicate the picture: the acid form of poly dC (145) and the helical structure of polydI. The acid form of polydC a t pH 6.9 still melts around 45°C in 0.41 M Na+ (or 60°C in 0.1 M Na') but can be eliminnted by raising the pH above 7.5. The helical structure of poly dI melts lower than poly d I poly dC, but the difference of the T,'s

-

-

-

119

SYNTHETIC POLTNUCLEOTIDES

diminishes as the salt concentration rises. The triple-stranded 2 poly dI * poly dC can be found only in high salt concentration, that is, in a limited range of temperature and ionic strength. Increased thermal stability of the complex formed between polydI and poly BrC allows an easier study of 2 poly dI * poly BrC (kW7). I n 0.41 M Na+ a t 42°C (above the dissociation of poly dI), the continuous variation method demonstrates the existence of both poly dI poly dBrC and 2 poly dI * poly dBrC, according to the ratios of the polymer residues. At 20°C the reaction of polydI with poly dBrC proceeds only to the formation of poly d I * poly dBrC. At low salt concentrations, 2 poly dI * poly dBrC dissociates in two steps: 2 poly dI poly dBrC -P 9

poly dI poly dBrC +

+ poly dI + 2 poly dI + poly dBrC

(the first dissociation takes place a t temperatures lower than 20°C, below 0.1 M Na'). Above a certain salt concentration (0.35M Na+), however, the dissociation of the three-stranded complex is direct: 2 poly dI . poly dBrC + 2 poly dI

+ poly dBrC

We thus find a set of equilibria similar to those for the associations of poly A and poly U, and it appears that i t might be completed by a fourth transition, a rearrangement of poly d I * poly dBrC above 0.35 M Na+: 2(poly dI .poly dBrC) -+ 2 poly dI poly dBrC

+ poly dBrC

The hydrogen bonding in such three-stranded complexes is unknown but it probably involves a triangular scheme in which each base is hydrogenbonded to the other two, unlike those proposed for poly A * 2 poly U and poly A 2 poly I . It is not unlikely that a triple-stranded complex, 2 poly I * poly C, is possible in the ribopolynucleotides a t high enough salt concentrations.

VI. Role of Sugar Phosphate Backbone A. The Phosphate Residues We examine here the effects of the sugar phosphate "backbone" on the stability of helical structures. The negative charges of the phosphate residues in general tend to repel each other, and the repulsions may be minimized in certain helical structures. In most associations of polynucleotides, the negative charges carried by the two or three strands result in a repulsion that decreases the stability of the structure. Exceptions to this are the acid helical forms of polyA and poly C. In poly A, the proton carried by N1 is not involved in hydrogen bonding

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between bases, but hydrogen bonding between one phosphate and the amino group of an adenine in the opposite strand has been postulated. The positive charges of the bases give an attractive force with the phosphate and stabilize the association of the two strands. Hence the T , of the acid form of poly A decreases as the salt concentration increases. The variation is much more complex for poly (C, c')for which the T , is a decreasing function of (Na+) above about pH 4 and an increasing one below. Opposite variations occur on each side of the pK of cytosine because only half of the bases must be titrated in the helix. The effect of salt concentration on the T , of polyI poly C+ has not been investigated. It may be expected that the T,,, of poly I * poly C' will be higher a t low ionic strength. I n most neutral complexes, shielding of the negative charges increases the stability; the effect is greater in triple-stranded helices. Divalent cations such as Mg++,a t low concentrations, also increase the stabilities of most neutral complexes. I n the presence of excess Mg++, T,(s-l) of poly A-2polyU is nearly independent of Na+ but decreases (187)above 0.01 M Mg++.Concomitantly, the breadth of the transition increases very markedly: one fourth of the optical density change takes place in a 0.2"C interval with no Mg++in 0.5 M Na+, and in a 12.5"C interval with 0.02 M Mg".

-

B. TheSugar We have already noted many differences between ribo- and deoxyribonucleotide structures : The helical semiprotonated forms of poly dC and poly dBrC are more stable than those of poly C and poly BrC (199). The T,, of poly (dA-dU) is 10°C lower than that of poly (rA-rU) (200). The T, of poly dI-poly dC is 14°C lower than that of poly I.poly C (in 0.1 M Na+) (205). Poly dG poly dC is dissociated a t considerably lower temperatures than poly G poly C ($9, 176). The T,'s of poly d T and poly dU are lower than those of the corresponding polyribonucleotides (147, 2008, 209). In cases where DNA-DNA and RNA-RNA helices of identical sequence and composition have been compared, the T,'s of the latter are about 8°C higher (200) (if thymine did not replace uracil in DNA, the difference would be larger).

-

-

Apart from the differences in stability, in the few cases examined, hypochromicity is larger in the ribose series. These differences are per-

121

SYNTHETIC POLYNUCLEOTIDES

haps partly a consequence of RNA helices being in the A form while DNA exists in solution in the B form. Evidence for this lies in the observation that neither RNA helices nor hybrid helices bind actinomycin D, which is bound only by DNA in the B form and not in the A form (210, 211).

Hybrid RNA-DNA helices melt lower (4OC) than the corresponding DNA-DNA helices (130,131, 212). Few polynucleotide hybrids have been extensively studied. Poly dG poly C has been found in a CsCl density gradient (128). It melts higher (83°C in 0.0013 M Na’) than poly dG * poly d C but lower than poly G * poly C. Two hybrids, poly dG poly C and poly G poly dC, have been synthesized with the enzyme “nucleic acid hybrid polymerase” (213).Their Tmlsare not very different, and both are about 20°C higher than that of poly dG poly dC. However, the stoichiometry of all these hybrids has not been wellestablished, and their study was not very extensive. The order is thus

-

-

-

poly G * p01y C

> poly G . poly dC > poly dG * poly C > poly dG . poly dC

A thorough examination of all four combinations of poly I and poly C has been carried out (206).The order of stability is as follows: poly I * poly C

> p01y I

*

poly dC

> p01y dI

poly dC

> poly dI

*

poly C

The stability of a hybrid structure is therefore not necessarily intermediate between those of the two corresponding nonhybrid associations. The crystallographic characteristics of all these complexes differ and do not resemble those of RNA and DNA helices. Postulates concerning A and B forms may be very oversimplified or even wrong, but it remains that, depending on the nature of the sugar phosphate backbone, geometrically different double helices can be formed, always using the same base-pairing pattern. It is difficult to make detailed explanations of the role of the sugar.2 However, the large differences in the stability and the optical properties of homologous deoxy, ribo, and hybrid complexes illustrate the importance of this part of the polymer in the formation of helices. It is sometimes overlooked, but one must expect the contribution to be decisive, since the sugar phosphate backbone has direct interactions with the surrounding aqueous medium (117). It may be relevant to the transcription of information from DNA to messenger RNA that a DNA-DNA helix is more stable than a hybrid. This may ensure that the process can take place, leaving the DNA un* Recent comprehensive ORD studies of ribo and deoxyribopolynucleotides have provided plausible explanations for many of the stability differences (261).

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changed while producing free RNA. This point has received support recently, using the various complexes of polyI and poly C as templates (205, 214). The remaining double helix is the more stable one, regardless of the template used.

C. Right- or left-Handed Helices, Parallelism, and Antiparallelism I n all cases where crystallographic studies have been made, polynucleotide helices have been found to be right-handed: DNA (2161, poly (dA-dT) (198), polyI poly C (216), poly A poly U (190), the acid forms of poly A ( 1 5 7 ) ,and poly C (168). Like DNA, poly (dA-dT), poly A * poly U, poly I * poly C, and poly A polyI all possess antiparallel structures, whereas, in the acid forms of poly A and poly C, the strands are parallel. Can a generalization be made for all homopolynucleotides? It is reasonable to assume that the two strands in such a structure must be equivalent ( 2 ) .This implies that each base pair is symmetrical, with an axis of symmetry. The helical 'axis will therefore also be a dyad axis in this type of structure built by association of two identical strands. It follows that the strands must always be parallel. With the same assumption that the strands will be absolutely equivalent, three-stranded homostructures must have a threefold axis of rotation. Such structures must also be parallel. (One can also reason as follows: a t least two strands must be parallel. If the third one is to be equivalent to the other two, it will be parallel to them.) It may be noted that, with respect to formation of organized secondary structure in homopolynucleotides, the rate will be independent of polymer concentration if an antiparallel structure is involved (folding of a single strand). In the case of parallel strands in a double (or triple) helix, the rate of formation of secondary structure will be concentrationdependent.

-

-

VI I. Reversibility Most of the associations studied are formed spontaneously when two polynucleotides are mixed in saline solution, and it follows that, after dissociation by elevation of temperature or by a change of pH, reassociation occurs when conditions are brought back to the original. However, hysteresis phenomena are found when polynucleotide complexes are titrated (189, 206) and, in certain conditions, thermal dissociations appear to be irreversible. Hysteresis loops have been observed in the acid dissociation of poly A poly U, poly A * 2 poly U and poly I poly C, and similar observations would be possible for poly A poly I and poly A * 2 poly I (but not poly G poly C because protonation does

-

-

-

SYNTHETIC POLYNUCLEOTIDES

123

not lead to dissociation) ( 2 9 ) . In all these cases, acidification allows the formation of a new structure, the protonated forms of polyA or poly C. When the complex is dissociated, the poly A (or poly C) strands associate, and in this form they do not react easily with complementary polynucleotides. Thus the original association is reformed a t a higher pH than the pH at which it was broken. There are no hysteresis loops in the alkaline titration of the same complexes, since alkaline, stable, ordered structures do not exist for these polynucleotides. Apparently irreversible thermal dissociations can also be observed when polynucleotides are first allowed to associate under favorable conditions, then transferred to different conditions (of salt concentration or pH) and heated. For example, preformed poly dI poly dC has a T , of 51°C a t pH 6.1 in 0.286 M Na'. The acid form of poly dC in the same solution melts a t 67"C, and polydI, also associated a t low temperature, melts a t 32°C. The thermal dissociation of poly d I * poly dC is irreversible and leads, when the solution is cooled, to a mixture of the separate ordered structures of poly dC and poly dI ( 1 4 5 ) . Another example lies in the thermal dissociation of poly G * poly C a t pH 2.5 (after preparation of the complex at pH 7). Recooling does not lead to reformation of the complex

-

(29)

f

Competition with protonated ordered structures is not the only case in which irreversible transformations, indicative of frozen equilibria, can be obtained. Thus poly dI and poly dBrC seem to give only polydI poly dBrC a t 20°C in 0.41 M Na+. But a t 42"C, as poly d I is dissociated, the reaction proceeds and 2 poly dI * poly dBrC is formed. This complex, which melts at 8loC, is then stable when brought down to 20°C (207).

-

VIII. Displacement Reactions The question of competition between two different associations is of interest. Such a competition arises when one polynucleotide (K) is able to associate with two different complementary polynucleotides (L and M). If the association K L is more stable than K M, we can expect L to displace M from its less stable complex. It is widely assumed that the higher the T,, the more stable the complex, and the T, has even been considered as a measure of absolute stability. If this is correct, we can predict which of two polynucleotides will displace the other. As far as neutral complexes are concerned, many examples have been found in which the complex of higher T, is the end product of a displacement, and no example to the contrary has been reported. But i t is not necessarily true for polynucleotide associations of very different types. For example, the reaction between poly A and poly U a t acid pH, though slow, goes in the opposite direction (187, 188). Poly A is displaced from

-

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A. M. MICHELSON, J . M A S S O U L I ~AND ~

w. GUSCHLBAUER

the association with itself even though this has a higher T, than poly A * poly U. A few examples of displacement reactions between neutral helical structures follow.

+

poly A . 2 poly I 2 poly C T, = 39°C (0.11 M Na+, pH 7)

-+

2(p0ly I * poly C) T, = 58°C

+ POIY A

This reaction (202) has been demonstrated by various methods such as ultraviolet absorption, infrared spectroscopy, and sucrose gradient sedimentation. Another example is (146): poly dI poly dC T,,,= 43°C (0.1 M Na+)

+ poly dBrC -+

poly dI . poly dBrC T, = 70°C

+ poly dC

Here poly dC may be in a structured form (T,,, = 48OC a t p H 7, in 0.1 M Na+),although no corresponding transition can be detected in the dissociation profiles. I n these two cases, therefore, it may be argued that the driving force in the reactions is the tendency to accommodate the maximum number of bases into rigid helical structures. However, a complete set of displacement reactions between the polydeoxy, polyribo, and hybrid associations of poly I and poly C has been reported (206).

+ poly dC poly I . poly dC + poly C T, = 52.3"C poly dI poly C + p01y I T, 35.4"C p01y dI poly dC + poly I

poly dI . poly C T, = 35.4"C

*

=

T,

= 46.1"C (0.1 M Na+, 30°C)

-+

-+

-+

poly dI . poly dC T, = 46.1"C

+ poly C

+ poly dC p l y I poly C + p01y dI T, = 60.2"C poly I . p01y dC + poly dI p l y I * poly C T, = 60.2"C *

T,

=

52.3"C

These reactions were demonstrated by examination of the melting profile a t given intervals of time. The fluorescence of ethidium bromide in the presence of double-stranded structures also provides a useful technique in certain cases ( 2 0 6 ~ ) . Moreover, a helix-helix rearrangement has also been shown by the same authors (206): poly dI p01y C 1

+ poly I

*

poly dC + poly I . poly C

+ poly dI

*

poly dC

Another example of reaction between two multistranded secondary structures lies in the association of poly A and poly X a t p H 5 (221).

125

SYNTHETIC POLYNUCLEOTIDES

Another series of displacements occurs when poly C and its halogenated analogs compete for association with poly I or poly G (217). In 0.15 M Na+, pH 7, the T, values are the following: poly I . poly c poly I poly BrC poly I . p01y IC

60.2"C 89.2"C 91.3"C

According to expectation, poly BrC and poly IC displace poly C. Equilibrium is reached in 1 hour a t 20°C. This system offers an opportunity to compare the stabilities of two structures, the Tm's of which are nearly equal. Such a comparison is of interest in view of the use that can be made of T, as a measure of stability. If one can write AG = A ( T , - T) , a direct comparison of AG values is possible for different complexes, provided the A constants are equal. The values are, of course, different for two- and three-stranded helices, but it may be supposed that, for similar double-stranded structures, A will be the same. Poly I poly BrC and poly I * poly I C constitute a most favorable case. If the A constants are really equal, polyI * poly I C should have a slightly lower free energy than poly I poly BrC, a t any temperature. This is confirmed by a slow displacement of poly BrC by poly I C ; half of the poly BrC is displaced, in dilute polymer solution ( l k 4 M in nucleotides), in 1 month a t 20°C. Conversely, poly BrC does not displace poly IC. Some expected displacement reactions have not been found to occur a t measurable rates, even though the T , differences must be large in both cases (217):

-

poly G poly C poly I * poly C

+ poly BrC

+ p01y G

-f,

-f,

+

poly G . poly BrC poly C] p01y G * poly C plyI

+

However the reverse displacements do not occur either, and it has been argued that large activation energies linked to the breakage of G-C or G-G bonds prevent these systems from reaching equilibrium.

IX. Polynucleotide Analogs A wide variety of polynucleotides containing analogs of the purines and pyrimidines occurring in nucleic acids has been examined (217~). Included in this group are also homopolyniers of methylated nucleotides, which occur in rather minor proportions as natural constituents. When hydrogen-bonding possibilities are blocked, as for example in poly N1-methyluridylate (86, 218) poly N1-methylinosinate, poly Nl,N7dimethylinosinate, and poly N1-methyladenylate ('7) , then complex formation with the appropriate polynucleotide (poly A, poly C,or poly U)

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does not occur. Nevertheless, double- (or multi-) stranded secondary structures may be possible; for example, poly NlMeA readily gives an “acid” form corresponding to that of poly A. This is in accord with the postulated structure for poly A a t acidic pH involving the 6-amino group and N7 for hydrogen bond formation. Alkylation of the 6-amino group in adenosine produces an effect that depends on the alkyl group. Thus polyA treated with formaldehyde to give essentially the 6-hydroxymethyl derivative does not give an “acid” form or complex with poly U (219), nor does a copolymer of adenylate and 6-methylaminoadenylate (some 70% of the latter) ( 7 ) . However, the homopolymer of “6-methyladenylate” (6 MeA) apparently interacts with poly U (but only to give the 1:1 complex) ; the thermal transition of this complex is very low (near 15°C) (220). I n the case of poly 6-hydroxyethyladenylate,no interaction occurs with either poly U or poly BrU (use of the latter would be expected to increase stability of a possible complex by some 20°C) (83). Poly 6-hydroxyethyl adenylate (HEA) does not give a doublestranded “acid” form and, in accord with the lack of complex formation with poly U, no interaction occurs with poly I. Stacking of the bases in poly HEA a t neutral p H is nevertheless demonstrated by dichroic studies; hence a rather powerful steric effect of the alkyl group must be postulated to explain the lack of interaction with polyU and poly I. Nevertheless it may be noted that a very stable complex is formed with polyxanthylate (221). Here the geometry must be such that steric hindrance by the hydroxyethyl group is eliminated. [Similarly, formaldehyde treatment of polyA does not prevent hydrogen bond formation with poly X (221).] Methylation a t N7 in purine nucleosides (guanosine, inosine) also reduces structural stability (7). Thus no secondary structure can be observed in poly N7-methylinosinate a t pH 7 (at this p H the base is zwitterionic) ; the thermal stability of secondary structure in poly N7methylguanylate acid is markedly less than that of poly G, though this is partly the effect of liberation of the free purine on heating. Similarly, the double helical structures formed by these polymers with polyC (or poly BrC) are dissociated under milder conditions than those containing the nonmethylated polymers. It may be noted that in these complexes (and also in structured poly 7-methylguanylate) the purine base is protonated a t pH 7 and hence carries a positive charge. The complexes are thus similar to poly G * poly C or poly G a t pH 2.5, where a reduction of stability also occurs despite possible interactions with the negative phosphate groups. In addition, the alkaline titration of poly 7MeI * poly C is noncooperative, suggesting that formation of the zwitterion of

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SYNTHETIC POLYNUCLEOTIDES

7-methylinosine on removal of a proton does not lead to a disruptive influence on the rest of the hydrogen-bonded base pairs (7). Methylation a t C5 in pyrimidine polynucleotides leads to increased stability in both the homopolynucleotide secondary structure and in the heterologous complex (85, 222). For example, poly 5-methyluridylate (poly ribothymidylate) has a T , some 30°C higher than that of poly U, and the complex with poly A is also more stable than poly A poly U by about 20°C (85, 218). A wide variety of analogs of polyuridylate have now been described (85, 223). These include homopolynucleotides containing 5-fluoro-, 5-chloro, 5-bromo-, s-iodo-, and 5-hydroxyuracil. With respect to the secondary structure of homopolynucleotide a t low temperatures, stability is increased by substitution by bromine or iodine and decreased by fluorine, chlorine, and hydroxyl. However, when the analogous complexes with poly A are considered, although decreased stability occurs with poly 5-hydroxyuridylate, a marked increase in stability is found not only with poly 5-bromo- (or iodo-) uridylate but also with poly 5-chlorouridylate (Fig. 13).The case of poly 5-fluorouridylate is somewhat special since the pK of the polymer is 8.1 (in 0.15 M NaCl). At pH 7 poly A 2 poly FU is about 10°C less stable than poly A 2 poly U, but at p H 6 the Tmlsare virtually identical (224). Polypseudouridylate may also be regarded as an analog of polyuridylate (225).Here the substituent a t C5 is the sugar phosphate chain itself. P o l y q has a remarkably stable secondary structure with a T , some 55°C higher than that of poly U. Both double- and triple-helical complexes are obtained with poly A. Absorbance-temperature profiles of poly A 2 poly q are biphasic, the second dissociation occurring a t a higher temperature than poly A * poly U in the absence of magnesium ions. Since magnesium ions have no effect on the T, of poly A poly @, the thermal stabilities of poly A * poly U and poly A * poly Q are identical in solvents containing magnesium ions (1W2M ) . An anomalous alkaline titration behavior is observed with both poly A * 2 poly q and the structured form of poly q acid. Considerable noncooperative loss of protons can occur before there is a cooperative rupture of secondary structure (85, 225). Halogen-substituted poly C’s have also been studied. Like poly C, both poly BrC and poly iodoC show no indication of a rigid secondary structure a t pH 7 (226). Contrary to an earlier report ( 8 5 ) , polyBrC readily forms an “acid” structure, comparable with th a t of poly C, but is much less stable. Similar decrease in stability of the “acid form” has been reported previously for poly dBrC compared with poly dC, both of which are nevertheless more stable than the corresponding polyribo-

-

-

-

-

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nucleotides. Poly iodoC shows rather more complex behavior since thermal dissociation of the acid form is biphasic. Spectral properties indicate two ordered “acid” structures, both of which are protonated. (226). Introduction of the bromine (or iodine) atom again increases the stability of hetero complexes. Thus both poly I poly BrC and poly I poly I C possess Tm’s some 30°C higher than that of polyI * poly C

-

-2

-I

0

log “a*]

FIG.13. Variation of T, with ionic strength of triple-stranded complexes of A and T, U, ClU, BrU, and IU. Symbol

0-0 0-0 V-V

v-v x-x

+-+

A- A

Complex

References

rA . rU rU rA rBrU. rBrU rA rClU .rClU dA . rU .rU dA * dT * dT dA rU . d T rA rIU .rIU

41, 48, 86

+

86, 818 86, 818 266 266 266 86

(85). Similar increase in stability is also seen in the corresponding complexes with poly G (29). As is generally the case, increase in thermal stability is accompanied by an increase in the pH necessary t o cause dissociation of such complexes. This apparent inversion of the effect of substitution a t C5 by bromine depending on whether a homologous or a heterologous complex is involved is readily explained. The acid form of poly C requires addition of a proton for each pair of bases. Since substitution a t C5 by bromine (or iodine) reduces the basicity of the pyrimi-

129

SYNTHETIC POLYNUCLEOTIDES

dine (as shown by pK values of the nucleosides), uptake of protons is rendered less facile and hence leads to an apparent decrease in stability at slightly acidic pH. At neutral pH, the normal stabilizing effect of increased dipole interactions is apparent. That other factors are involved is shown by the fact that a similar inversion occurs if ribo- and deoxyribopolycytidylates are compared. Here the “acid” homologous complex formed by poly C is less stable than that of poly dC, whereas poly I * poly C is more stable than poly I poly dC. The stabilizing effect of substitution by bromine a t C5 of pyrimidines has been fully documented in the case of polydeoxyribonucleotides. Poly (dA-dBrU) has a T, 8°C higher than that of poly (dA-dU) (198) whereas the difference between poly A * 2 poly U and the corresponding polyA 2 poly BrU is some 36”C, that is 18°C per substitution since two strands of the polyU analog are involved (85). However, poly dI poly dBrC has a T, some 26°C higher than poly dI * poly dC (199), that is, the same difference as that between poly I poly C and poly I poly BrC in the ribose series. It thus appears that vertical interactions play a marked role in the stability of helical complexes. This is also seen in a comparison of the T,’s of poly (A-U) and poly A poly U since it may reasonably be assumed that the strengths of the hydrogen bonds (in both cases between A and U ) are the same. It has been suggested that subsitution by bromine increases hydrogen bond strength, but this seems insufficient as an explanation, since the effect should be independent of sequence, and this is clearly not the case. The explanation quoted by Inman and Baldwin (199) for an alleged differential effect of bromine substitution a t C5 in cytosine or in uracil cannot be accepted. First, there is no validity in the statement that bromine is ortho to the donor group in one case and meta in the other since both systems are similar (Br-C-C-N-H). Second, the direct comparison of an alternating copolymer with a complex formed from two homopolymers is not valid, as we have seen above. Third, if homologous structures are considered, poly BrU is more stable than poly U while poly BrC (acid form) is less stable than poly C. Another nucleotide analog that bears a resemblance to pseudouridylic acid is 3-isoadenylic acid ( 4 ) , in which the glycosyl linkage is a t N3 instead of N7. Polyisoadenylate readily interacts with poly U or polyI to give 1:l complexes, which are extremely stable in comparison with the corresponding structures containing poly A. Unlike poly A, however, poly isoA does not give a double-helical structured “acid form.” The hypochromicity of the polymer and the effect of increase in temperature a t p H 7 suggest that, like poly A, the bases are stacked to a high degree, thus giving a single-stranded helical conformation. While the greater

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basicity of 3-isoadenosine (pK = 5.5) compared with adenosine (pK = 3.45) may promote stronger hydrogen bonding, it is more likely that the increased stability of the complexes is a result of increased interplanar interactions, with perhaps an increase in organization of solvent molecules also playing a role ( 4 ) . In contrast with poly A * poly U, the complex poly isoA poly U shows no hyperchromic effect in the region of 290 mp. This is perhaps another demonstration that the assignment of particular transitions ( T + T” or n+ x ” ) to specific wave bands is a little premature a t present. Like poly I and poly G, the homopolymer of xanthylic acid possesses a helical secondary structure (221). The structural stability of poly X is somewhat greater than that of poly I, but markedly less than that of poly G. In contrast with polyI and poly G, polyX does not interact with poly C (or poly BrC) under a wide variety of experimental conditions. However complexes are readily formed with poly A, polyI, and poly U, and also with a wide variety of analogs of poly A, including poly 3-isoadenylate7 poly N1-methylA, and poly N6-methyl (or hydroxymethyl, or hydroxyethyl) A. Certain peculiarities are shown by these complexes. Thus the stability of poly A * 2 poly X is increased on lowering the pH (removal of negative charges from ionized xanthine residues) whereas that of poly U poly X is decreased (2221). Several other extremely interesting polynucleotide analogs have been prepared, but details of their physical properties are not as yet available (227). e

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X. Theory and Practice of Helix-Coil Transitions Ever since the proposal of the DNA model by Crick and Watson (1954) (215), the topological, mechanical, and thermodynamical prob-

lems of this structure have interested physical chemists. Numerous suggestions have been presented (228-233) for the untwisting and or “unzippering” of a double-stranded helix of the type of DNA, particularly in view of mechanisms of replication of DNA. These investigations essentially show the mechanical feasibility of the sequential opening mechanism for a double helix. No studies of the considerably more complicated case of a triple helix have been reported. On the other hand, considerable energy has been invested, particularly in recent years, in studies of the energetical aspects of the helixrandom coil transition. Statistical mechanics was originally used to describe this transition in polypeptides (i.e., the single-stranded a! helix) (234, 2%). This treatment has been extended to the DNA double helix and to polynucleotides by Hill and others (236-939). Zimm (6373 also investigated the effect of chain ends on the stability and coopera-

SYNTHETIC YOLYNUCLEOTIDES

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tiveness of the transition. He introduced a parameter, uo (in later papers 1 , ’ ~ related )~ to the free energy, C, of stacking the base pairs on top of each other: UO = exp (-e/kT) Zimm demonstrated that the sharpness of the transition depended on the value of the stacking parameter, u0. Comparing experimental values for the melting of DNA (in 1959) which indicated a breadth of the melting transition of about 2”C, Zimm estimated 0.1 < uo < 0.01 and therefore 3 kcal < L < 1.5 kcal. Since the melting curves become steeper (more cooperative j with more homogeneous and longer DNA chains, he was inclined to prefer the higher value. In practice, it has been found that this value still seems too small. I n a more refined treatment of poly (dA-dT) and poly dI * poly dC, Crothers and Zimm (239), using the partition function of Lifson and Zimm (S40), evaluated u0 between le3 and for these two cases. They used in their studies the measured value of AH = 7 keal/mole of Rawitscher et at. (163) for the stacking energy in poly A. This gives a breadth of the thermal transition of less than l0C, quite in agreement with the best recent observations on polynucleotides and DNA. This means that the transition of a completely and perfectly aligned long helix is much more cooperative than Zimm had evaluated from the earlier DNA data. Similarly, statistical thermodynamical treatments have been used by Applequist and Damle (941) to investigate the effect of chain length on polynucleotide transitions and by Magee et al. (242, 243) to study homo- and copolymeric binding. These calculations all predict a decrease of T,,, with decreasing degree of polymerization and a broadening of the melting profile (244-246a). This broadening is not always seen (246). Ionic strength has a very marked effect upon the stability of the helix. Schildkraut and Lifson (247) have studied this in detail, and developed a generalized theory of electrostatic repulsion based on the Debye-Huckel treatment.

XI. Factors Governing Structure All homopolynucleotides derived from natural, or almost natural, nucleotides form multistranded secondary structures under appropriate conditions. In the case of the “basic” (or 6-amino) nucleosides (A and C) , slightly acidic conditions are necessary, whereas the “acidic” (or 6-keto) polymers (G, U, I, X ) all give defined secondary structures a t pH 7, albeit with vastly different stabilities. Disregarding complexes containing substituted bases where substitution merely increases or decreases the stability without change in specificity, the family of inter-

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actions so far experimentally observed with homopolynucleotides can be summarized as follows: A A

+

c

-

C (H+)

+

U

G

+

-

-

+

(H+)

+

I +

+

+

-

-

-

X

+

u

+

-

G

-

+

-

I

+

+

-

-

+

+

x

+

-

+

-

+

+

+

-

-

While certain interactions cannot be demonstrated (for example, of A with G ) , this may be simply a consequence of the greater stability of secondary structure in poly G compared with poly A poly G ; that is, G-G interactions (in a total sense) are greater than those for A-G, whereas, in the case of poly I, the 1-1interactions are weaker than those present in poly A poly I. However, if interaction between G and G is excluded by some means such as spatial distribution, the possibility arises that A-G base pair formation could occur. This argument, also applicable to poly U poly G (compare with the known poly U poly X) cannot be extended to the absence of poly U poly C, and one can only assume that the stability of such a structure is extremely low. General rules governing stability of helical complexes have usually been analeptic in character. Despite the wealth of information now available, prediction is still hazardous. For example, although the alternating poly (A-U) is some 10°C more stable than polyA poly U, it cannot be assumed that alternating poly (G-C), when available, will likewise be more stable than poly G poly C. This involves virtually the simplest comparison possible:

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-

It is quite likely that vertical G-G and C-C interactions are in total stronger than G-C interactions (248),whereas A-U (or T) interactions are greater than those of A-A plus U-U (or T-T), thus leading to an inversion of the relative stabilities. The fact that the same enzyme produces poly dC poly dG but alternating poly (dA-dT) suggests that this might be the case, if one assumes that the enzyme catalyzes formation of the most stable possibility under admittedly abnormal conditions.

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SYNTHETIC POLYNUCLEOTIDES

133

However, several major factors govcrning structural stability of polynucleotide complexes can be defined. Clearly, direct interplanar interactions (London, van der Waals forces) (%’&a) play an important role and can be modified by substitution of the bases. I n addition “hydrophobic” forces, defined as organization of solvent molecules in the vicinity of the helix (249),exert a considerable control, and can be affected by base substitutions (including methyl) and also by the nature of the sugar. Finally, hydrogen bonds certainly control specificity of interactions and contribute t o stability. This contribution is by no means overwhelming and the concept that the three hydrogen bonds possible in poly G poly C are responsible for the greater stability than that of poly A poly U, containing two hydrogen bonds, is unwarranted. Despite the failure of efforts to obtain interactions involving only one hydrogen bond (e.g., poly A poly N1-methyl-5-bromouridylate), such a possibility cannot be excluded, since steric or disruptive effects on solvation by the N1-methyl group may be important. One hydrogen bond may well be insufficient to create the planarity induced by two or more hydrogen bonds, but one can nevertheless visualize possible structures. The presence of considerable stacking of bases in single-stranded polynucleotides and even oligonucleotides indicated by earlier studies (1) has now been extensively confirmed (70, 82, 169, 161). I n this connection, mention must be made of the definitive ORD studies on oligonucleotides by Tinoco and his co-workers (69, 70, 260, 251). The effect of chain length (251a) has also been studied by various techniques and it is now clear that interplanar interactions contribute significantly to single-stranded secondary and tertiary structures. However, this concept has been much misunderstood as leading to virtually rigid single-helical structures which is certainly not the case, either in polyA or in tRNA. The contradiction between hydrodynamic and optical results can be resolved readily if account is taken of the time of stacking, and the fluctuation of this stacking along a chain. Again, stacking of bases in a single strand represents not the lack of freedom as in a DNA double helix, but rather a restriction of rotation or separation of the bases, which are nevertheless in movement even when stacked. Nevertheless, this kind of stacking interaction is undoubtedly important with polymerizing enzymes such as DNA polymerase and RNA polymerase, where hydrogen bonds provide specificity, but interplanar interactions between the terminal nucleotide in the chain and the approaching nucleoside triphosphate provide some of the binding energy. Among other approaches indicating base stacking can be mentioned the transfer of excitation energy between the neighboring nucleotide bases determined by studies of the luminescence of various oligonucleotides. Here again, the order of

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+

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sequence, that is, direction of the chain, plays an important role (252, 253). The question of flexibility of double helices has received little attention recently, apart from studies of the polarization of fluorescence of acriflavine conjugates of polyribonucleotides (254). Some loss of rigidity precedes ultraviolet absorption effects accompanying thermal dissociation of double- and triple-stranded structures. Flexibility in a double helix is possibly quite important for the formation of certain triple helices. For example poly G and poly C give only a double helix whereas degraded polymer (either G or C) gives rise to 2 p o l y G poly C; that is, a third strand can be added only if a certain number of breaks are present in one strand of the double helix. Similar reasoning may explain why polyG (probably double helix) is much less viscous than oligo G, which is multistranded and can apparently aggregate to the point of precipitation. Yet another indication of the fact that hydrogen bonds alone play a relatively slight role in the stability of double helices is the observation that the Tm’sof complexes of a series of oligouridylates with polyA are markedly less than those of an analogous series of oligo A’s complexed with poly U. It is probably too naive to attempt explanations of the biological effects of various nucleotide analogs in terms of the physical behavior of homopolynucleotide analogs, despite certain marked effects. Explanations of the mutational effect of incorporation of bromouracil as a result of increased tautomerization are equally suspect despite the beauty of simplicity. It may well be that such an analog modifies the behavior of an adjacent base by increase in the interplanar interaction. I n this respect it may be noted that the observed energy transfer is much more marked in A +-BrU than in A + U. This may well lead to a different electron distribution giving increased tautomerization and even a change in chemical reactivity of the neighboring base in addition to the modified behavior of the analog compared with the natural base. Another overlooked possible cause of biological effect may be the slowing down of reading, thus increasing the possibility of error due to misreading of a tautomer. If replication, transcription, and translation use somewhat different time scales (even if very different from the time of tautomeriaation), one could visualize an analog (e.g., FU) behaving abnormally in one case and normally in another.

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93 (1961); J. R. Fresco and D. B. Strauss, Am. Scientist 50, 158 (1962); B. H. Zimm and N. R. Kallenbach, Ann. Re v . Phys. Chem. 13, 171 (1962); I. Tinoco and D. N. Holcomb, ibid. 15, 371 (1964). 2. M. Grunberg-Manago, This series, 1, 93 (1963). 3. A. M. Michelson, J. Dondon, and M . Grunberg-Manago, Biochim. Biophys. Acta 55, 529 (1962). 4. A. M. Michelson, C. Monny, R. A. Laursen, and N. J. Leonard, Biochim. Biophys. Acta 119,258 (1966). 5. M. N. Thang, M. Graffe, and M. Grunberg-Manago, Biochim. Biophys. Acta 108, 125 (1965). 6. J. Weill, N. Befort, B. Rether, and J. P. Ebel, Biochem. Biophys. Res. Commun. 15, 447 (1964). 7. A. M. Michelson and F. Pochon, Biochim. Biophys. Acta 114, 469 (1966). 8. L. Grossmann, Proc. Natl. Acad. Sci. U S . 50, 657 (1963). 9. J. H. Philips, D. M. Brown, R. Admann, and L. Grossmann, J . Mol. Biol. 12, 816 (1965).

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47. T. V. Venkstern and A. A. Bayev, “Absorption Spectra of Nucleotides and Oligonucleotides.” Nauka Press, Moscow, 1965. S. E. Kerr and K. Seraidarian, J . Biol. Chem. 159, 211 (1945). 49. R. F. Steiner, J. Biol. Chem. 236, 3037 (1961). 60. J. C. Reid and A. W. Pratt, Biochem. Biophys. Res. Commun. 3, 377 (1960). 61. S. K. Vadenko, S. G. Komzolova, and D. G. Knorre, Biokhimyia 27, 142 @.

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62. S. Lee, D. McMullen, G. L. Brown, and A. R. Stokes, Biochem. J . 94, 314 ( 1965).

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63.

1831 (1964). 66.

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New York, 1957. 66. J. B. Biot, Mem. Acad. Sci., Toulouse 2, 41 (1817). 67. P. Drude, “Lehrbuch der Optik.” Hirzel, Leipzig, 1906. 68. C. Djerassi, “Optical Rotatory Dispersion.” McGraw-Hill, New York, 1960. 69. W. Moffitt, J . Chem. Phys. 25, 467 (1956). 69a. W. Moffitt, Proc. Natl. Acad. Sci. U S . 42, 736 (1956). 60. W. Moffitt and J. T. Yang, Proc. Natl. Acad. Sci. U S . 42, 596 (1956). 61. W. Moffit, D. D. Fitts, and J. G. Kirkwood, Proc. Natl. Acad. Sci. US.43, 723 (1957). 62. S. F. Mason, Nature 199, 139 (1963). 63. I. Tinoco, J. Am. Chem. Soc. 82, 4785 (1960). 64. I. Tinoco, J. Chem. Ph.ys. 34, 1067 (1961). 66. J. R. Fresco, Tetrahedron 13, 185 (1961). 66. J. T. Yang and T. Samejima, J . Am. Chem. Soc. 85, 4039 (1963). 67. T. Samejima and J. T. Yang, Biochemistry 3, 613 (1964). 68. T. Samejima and J. T. Yang, J. Biol. Chem. 240,2094 (1965). 69. D. N. Holcomb and I. Tinoco, Biopolymers 3, 121 (1965). 70. C. R. Cantor and I. Tinoco, J. MoZ. B i d . 13, 65 (1965).

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The DNA of Chloroplasts. Mitochondria. and Centrioles

I

S. GRANICKAND AHARONGIBOR Rockefeller 1Jniversity. New Yolk. New York

I . Introduction . . . . . . . . . . . . . . I1. Chloroplast DNA . . . . . . . . . . . . . A . Evidence for the Presence of DNA in Chloroplasts . . H . Hereditary Factors of Chloroplasts . . . . . . . C . Genetic Significance of Chloroplast DNA . . . . . D . The Problem of Labile or “Metabolic” DNA . . . . I11. Mitochondrial DNA . . . . . . . . . . . . A . Evidence for the Presence of DNA in Mitochondria . . B . Hereditary Factors of Mitochondria . . . . . . . C . Genetic Significance of Mitochondrial DNA . . . . I V. Centriole DNA . . . . . . . . . . . . . . A . The Function of the Procentriole . . . . . . . B . Differentiation of the Procentriole . . . . . . . C. Evidence for DNA in Procentrioles . . . . . . . D . Genetic Aspects of the Procentriole . . . . . . . V . The Role of Cytoplasmic Nucleoids in Inheritance . . . A . Redundancy and Genetic Analysis . . . . . . . B . Constitution of a Cytoplasmic Nucleoid . . . . . C . Redundancy, Organelle Survival, and Organelle Mutation D . Interactions of Nuclear and Cytoplasmic Genes . . .

VI . Summary . . . . . . . . . . . . . . . References .

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

. . 143 . . 146

. .

146 151 157 160 161 161 165 168 172 172 . . 174 . . 176 . . 177 . . 179 . . 179 . . 180 . . 180 . . 181 . . 182 . . 183

. . . . . . . . .

. . . . . . . . .

.

1 Introduction Until recently all of the DNA of a eucaryote cell was considered t o be contained in the nucleus. a body separated from the cytoplasm by a double membrane . I n the last few years evidence has accumulated that small amounts of DNA (a few per cent of the total) are also present in the cytoplasm. localized in three kinds of organelles : the mitochondria. the centrioles. and. in plant cells. the plastids. A number of general properties of these cytoplasmic organelles pertinent to our discussion may be noted here . 143

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The mitochondria and the plastids in a eucaryote cell are surrounded by double membranes but the centriole lies free in the cytoplasm. All three organelles possess both immature and mature stages; that is, they undergo differentiation and maturation. To simplify terminology, we designate the immature stage of the mitochondrion as the promitochondrion, the immature stage of the centriole as the procentriole, and the immature stage of the plastid as the proplastid.

DNA

TABLE I SOMENUCLEOIDS

OF

Origin Mitochondrion Mitochondrion Mitochondrion Mitochondrion Mitochondrion Mitochondrion Chloroplast Chloroplast Chloroplast Chloroplast Chloroplast

of of of of of of

yeast Tetrahymena Neurospora beef heart Phaseolus Brassica, Ipomaea

of Acetabularia of Acetabularia of broad bean of tobacco of Euglena

DNA per nucleoid ( X 10l6gm) 1

3.7 0.2-1.8 0.5

5 5 1-10 3-4 150 80

References 6 7 8

9 10 10 11 12 13

110

14 15

Centriole or basal body of Tetrahymena

2

16

Pox virus T-even bacteriophages Polyoma virus

2 2 0.05

17

Escherichia coli Yeast nucleus (haploid) Neurospora nucleus (haploid)

50 250 900

17 17 18 19 20

The amounts of DNA in all three organelles are summarized in Table I. Depending on the species, the DNA of a mitochondrion varies from 0.5 to 5 X gm and the DNA of a chloroplast varies from 1 to 150 x gm. If we consider the lowest values of DNA reported for the three organelles then the DNA per organelle is remarkably similar, i.e., about 1 to 5 X gm, and in the range of values for the DNA of certain viruses (Table I ) . It is interesting to note that E . coli contains only 10 to 50 times this amount of DNA, yet the E . coli DNA can code

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145

for all the enzymes that make protoplasm out of inorganic salts and glucose. This is not surprising when it is considered that an amount of DNA of 1 X gin is sufficient to code for several hundred different kinds of protein molecules. A common feature of the cytoplasmic organelles is that usually only the organelles of the cytoplasm of the female gamete are inherited. The cytoplasmic organelles of the male gamete are often discarded a t fertilization or shortly thereafter. This “maternal” inheritance has made analysis possible of some of their genetic properties. The genetics of individual organelles cannot be studied by Mendelian analysis because of the large numbers of organelles per cell. In most plant and animal cells the mitochondria may number 20 to 1000 and the chloroplasts may number from several to 100. I n the higher animal cell at interphase there may be only two centrioles, each accompanied by a tiny companion procentriole. However, in the protozoan ciliates, the surface coat or pellicle contains numerous cilia; each cilium arises from a centriolelike body and in Tetrahymena there may be over 1000 of these bodies. The large number of organelles of the same kind that exist in a cell represents a high degree of genetic redundancy. For example there may be several hundred different kinds of genes per organelle, present as a DNA unit or nucleoid, and several hundred such nucleoids in a single cell. If a mutation occurred in one gene of a nucleoid it would not be possible to recognize that event. Owing to this redundancy, genetic studies have been largely unsuccessful in demonstrating cytoplasmic unit factors of inheritance, Only recently, because of the accumulating evidence for the presence, replication, and functioning of DNA in cytoplasmic organelles, has i t become generally accepted that cytoplasmic unit hereditary factors, i.e., genes or cistrons, exist. In this chapter we present evidence for the following conclusions: (1) the cytoplasmic organelles contain small amounts of DNA (Table I) ; (2) each type of organelle contains its own characteristic DNA; (3) the DNA of the organelles is double-stranded, and replicates in a semiconservative fashion; (4) the DNA codes for specific RNAs, and the RNAs are translated into proteins. For completeness, mention also should be made of other heritable units that may come to reside in the eucaryotc cell. These are infectious agents such as bacteria, rickettsias, and bacteroids, which contain both DNA and RNA, and viruses that contain DNA or RNA. The viruses may span the spectrum from those that are foreign agents, to those that are part of or may have originated from chromosomal genes. I n some cases, a virus may infect and kill the host cell, or live in a steady-

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state relation in the cytoplasm of the host cell as a complete or incomplete noninfectious virus. I n other cases, the DNA of a virus may become incorporated into the host cell nuclear DNA as an “episome” and replicate along with the host DNA; a t a later time the episomal DNA may detach from the host DNA, then replicate freely and assemble specific coat proteins around itself to form an infectious virus or an incomplete noninfectious virus. An incomplete virus may become infectious by using the coat protein of a closely related infectious virus. Whether a viral DNA as an episome may attach to the DNA of a cytoplasmic organelle is unknown, but theoretically possible. The origin of some viruses from chromosomal genes is suggested by the studies of Sonneborn (1). He found that a nuclear gene in Paramecium may give rise to an mRNA that can infect a strain of Didinium (another protozoan) ; this mRNA will then multiply in Didinium as if it were an RNA virus. That an episome may in some cases be a normally functioning component of a cell is suggested by the finding by Douthit and Halvorson ( l a ) . I n B. cereus the DNA has a density 1.696. When spores are germinated a satellite peak arises of density 1.725, which later disappears. The authors suggest that sporulation may be controlled by an episomal element present in variable amounts during a life cycle. Excellent discussions of various features of cytoplasmic inheritance for further reference are the books by J. L. Jinks ( 2 ) and D. Wilkie ( S ) , and the papers in the recent symposium of the Society of American Naturalists ( 4 ) and the International Symposium of Genetics ( 5 ) .

II. Chloroplast DNA A. Evidence for the Presence of DNA in Chloroplasts The presence of DNA in chloroplasts has been established by a number of procedures: by direct chemical determination and physical characterization of the DNA from isolated chloroplasts ; by electron microscopic visualization of DNA in the chloroplasts ; by incorporation of labeled precursors of DNA into chloroplasts ; by radioautography ; and by cytochemical staining. We consider in the following some of the evidence obtained by these procedures. 1. DIRECTDETERMINATION OF DNA

ISOLATED CHLOROPLASTS Analysis of isolated chloroplasts from different plants indicates that appreciable quantities of DNA are present in them (21). For example, Pollard (22) estimated that spinach chloroplasts, prepared by different isolation and purification techniques, contain about 3-4 pg DNA per mg protein. FROM

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These analyses were subject to the criticism that the chloroplast fractions isolated were probably contaminated with nuclear DNA. The extent of contamination became apparent when qualitative distinctions were found between nuclear DNA and chloroplast DNA. For example, Fig. 1, from thc work of Chun e t al. ( 2 3 ) ,is ii densitoiiietcr tracing of a U.V. photograph of DNA from beet leaves, sedimented in a CsCl density gradient. Curve a represents DNA extracted from whole leaves. A major band (a) is present, representing nuclear DNA. Curve b represents the

Density (g/cni3)

FIG. 1. Densitometer tracing of DNA's from beet leaves. [From Chun e t d. (28.1 ( a ) Whole leaf DNA; the major band is a ; M represents a density marker DNA. ( b ) DNA of isolated chloroplasts; one satellite /3 is apparent. ( c ) The /3 band has been further purified by repeated centrifugation in CsCI; another band in addition to the p band is seen as a shoulder of the a band.

DNA isolated from a so-called purified chloroplast preparation. There is, in addition to the LY band, a heavier band (@) that is scarcely detectable in curve a . This indicates that the @ band is concentrated in the chloroplasts. It is apparent, however, that even in this purified chloroplast preparation the major quantity of DNA is still of the a! band type identified as nuclear DNA and present as a contaminant of the chloroplast preparation. Better evidence that the chloroplasts themselves contain DNA was obtained by the use of enucleated Acetabularia as the source material from which chloroplasts were isolated. Gibor and Ieawa (11) determined the DNA in such preparations and estimated that :i single chloro-

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plast contains a t least 1 X 10-lGgni DNA. Similarly, Baltus and Brachet (12) found that DNA is present in chloroplasts derived from enucleated cells of Acetabularia. The uniform density of the DNA derived from the chloroplasts of Acetabularia is apparent in Fig. 2 (24) which represents the densitometer tracing of a CsCl gradient in which such DNA was sedimented. A single, symmetrical band is apparent in this preparation. The chloroplast DNA of Acetabularia differs from the nuclear DNA. This is apparent from Fig. 3, which represents the densitometer tracing of DNA isolated from young cysts of Acetabularia (24). Young cysts contain numerous chloroplasts and a single nucleus ( 2 5 ) . The prominent 1.695

1.742

I FIQ.2. Densitometer tracing of DNA from chloroplasts isolated from enucleated Acetabularia cells. [From Gibor (ag).l The heavy band (1.742) is an added marlier DNA.

DNA band is that of the chloroplasts. Which of the additional bands represents nuclear DNA is not yet definitely established. Another type of evidence indicating that chloroplasts contain their own DNA relies on the qualitative base differences between nuclear DNA and the DNA associated with isolated chloroplasts. Kirk (IS) found that nuclear DNA of bean plants may be extracted in 30 minutes a t 70°C with 0.5 N perchloric acid. On the other hand, 2 N perchloric acid is required to extract the DNA of the chloroplasts a t the same time and temperature. The ratio of adenine to guanine in the nuclear DNA was found to be 1.54 while that in chloroplast DNA was found to be 1.67. In many cases, chloroplast DNA differs from nuclear DNA in buoyant density and melting point temperature. A higher GC ratio is di-

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rectly correlated with an increased density and an elevated melting point (26, 27). As mentioned above, DNA extracted from chloroplast preparations and analyzed by sedimentation in a CsCl gradient is rich in a satellite type DNA, significantly different from the major band in buoyant density. Brawerman and Eisenstadt (15) succeeded in isolating this satellite DNA in a pure form by the following procedure. Isolated chloroplast preparations from Euglena were partially lysed in a solution of 1%

PIG.3. Densitometer tracing of DNA froin cysts of Acetabularia. Chloroplast DNA (band 1.697) is still the major component; the two other bands probably represent nuclear and mitochondrial DNA. [From Gibor (94).I

deoxycholate containing 5 mM MgCl,. A green pellet was obtained by centrifuging the suspension a t 23,000 g for 30 minutes. DNA extracted from this green pellet was practically all of the satellite type. This established clearly that the satellite DNA is associated with the chloroplasts. The distinguishing marks of this chloroplast DNA are: (1) absence of the base 5-methylcytosine1 present in the nuclear DNA of Euglena; (2) the chloroplast DNA is heterogeneous with respect to buoyant density with peaks a t 1.684 and 1.692 gm/ml, as compared to 1.708 gm/ml for the nuclear DNA; the 1.692 band might represent the mitochondrial DNA (88) ; (3) the melting temperature of this chloroplast DNA is 78230°C as compared to 89-91°C for the nuclear DNA; (4) in who!e cell DNA, the ratio of A to G is close to 1; it is 2.8 for chloroplast DNA.

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DNA from a variety of other plants was similarly analyzed by centrifugation in CsCl gradient. Distinguishable DNA satellites were recognized in all cases and these satellites were found to be associated with the plastids. Data from such analyses have been summarized by Iwamura (29) and by Schiff and Epstein (28). Additional data that became available more recently are presented in Table 11. 2. CYTOLOGICAL EVIDENCE FOR CHLOROPLAST DNA

Under this heading are included studies on the visualization of chloroplast DNA by electron microscopy or other staining techniques, TABLE I1 BUOYANT DENSITYIN CSCL OF DNA SPECIESFROM PLANTS Source

Major (nuclear)

Satellite (plastids)

Tobacco Polytoma Swiss chard Wheat Acetabuhriaa Mung bean Turnip Sweet potato Onion

1.690 1.725 1.689 1.707 (1.724)(1.717) 1.691 1.692 1.692 1.688

1.703 1.711 1.700 1.716 1.695 1.706 1.700, 1.706 1.706 1.706, 1.718

Reference 30 31 32 33 24

10 10 10 10

The 1.695 band is DNA from chloroplasts; the other bands seen in DNA from cysts represent probably nuclear and mitochondria1 DNA. 0

and studies on the incorporation of labeled DNA precursors into chloroplasts. Stained or labeled structures were identified as DNA by their susceptibility to hydrolysis by DNase. I n such studies, dyes specific for DNA were used, and the cells treated with DNase were compared to cells not so treated. This double specificity was thus utilized for establishing the presence of DNA. I n the chloroplast of Chlamydomonas ( 3 4 ) , acridine orange staining revealed a yellow-green fluorescence characteristic for double-stranded DNA. DNase-treated cells did not show such fluorescing bodies. Similarly, Feulgen-staining bodies, which indicated the presence of DNA, were found in the chloroplast but were absent from cells treated with DNase. Electron microscopy studies of the chloroplast indicated the presence of fibrils, 25-30 A thick, which were identified as DNA because of their resemblance to the fibrils found in the nucleoid region of bacteria and also because such fibrils were absent from DNase-treated cells. Young leaves of Swiss chard were similarly studied by Kislev et al. (36).Their criteria for identifying DNA in the

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chloroplasts included: ( 1 ) Fcuigcn staining and its abscnce after DNase digestion; (2) the finding of uranyl acetate-stained fibrils, 30 B thick, also susceptible to DNase digestion; (3) the finding that, during growth, tritiated thymidine is incorporated into the chloroplast region and that the label is removed by treatment with DNase. Similar cytological data from studies on various plant materials are summarized in Table 111. TABLE I11 CYTOLOGICAL EVIDENCE FOR DNA Material

CHLOROPLASTS

IN

Results

Chlamydomonas

Fibrillar material observed by electron microscopy, removed after DNase treatment. Feulgen test and acridine orange test positive before DNase treatment (34) Swiss chard Fibrillar material removable by DNase. Positive Feulgen test. Incorporation of tritiated thymidine, label removable bv DNase (3.9) Euglena Incorporation of tritiated guanine and tritiated adenine. Part of incorporated label found resistant to RNase but removable by DNase (36) Dictyota, Padina, Bryopsis Thymidine incorporation found in chloroplasts of these marine algae. Dictyota and Padina incorporated thymidine at some stages of their development cycle only. DNase or hot 5 % trichloracetic acid removed label (36) Allium DNA fibrils observed in several regions of the organelle (37) Tobacco leaves Incorporation of thymidine in young but not in old leaves. Label removed by DNase or 1.6 N HC10, at 70°C (38) ~I

B.

Hereditary Factors of Chloroplasts The finding of DNA in the chloroplasts provides independent evidence for the hypothesis that these organelles possess their own hereditary factors, i.e., genes or cistrons. This hypothesis had been formulated in the early 1900’s on the basis of some of the following kinds of observations.

1. PHYSICAL CONTINUITY OF THE PLASTIDS

The plastids are self-duplicating structures that arise by division of pre-existing plastids ( 2 1 ) .This physical continuity of the plastids is best observed in algal cells that have one or two plastids. These plastids divide and separate into the daughter cells a t the time of cell division. An unequivocal example of dividing chloroplasts is provided by the

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cinematographic studies of Green (39) on growing cells of Nitella. The division time for these chloroplasts was estimated to be about 22 hours. 2. PLASTID CLONES

There are a few observations that, within the same cell, different types of plastids may be present, and that these plastid clones maintain themselves in subsequent cell divisions. Such observations are important because they indicate that plastid differences are maintained in the presence of the identical cytoplasm and nucleus. One example is pro-

FIQ. 4. A “mixrd” cell of Antirrhinum mnjus that contains large green plastids and small white plastids in the center. LFrom Hagemann (4l).l (magnified approx. x 1200).

vided by the presence of two different kinds of chloroplasts in SpiroNgyra cells (40) ; one kind of chloroplast lacked pyrenoids, so starch granules were scattered throughout the plastid instead of being localized on the pyrenoids as in the normal plastid. Hagemann (41) has recently summarized the evidence for the presence of different types of plastids within the same cells of higher plants. Such cells are usually found in border regions between green and white areas of variegated leaves. The presence of green chloroplasts and small undeveloped white plastids side by side in the same cell (Fig. 4) is proof of intrinsic differences residing in the organelles themselves. Furthermore, the mutation affecting the development of such white plastids apparently cannot be cured by simple diffusible metabolites from the adjacent green plastids.

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3. MATERNAL INHERITANCE

The results of reciprocal crosses of many variegated higher plants also support the hypothesis that hereditary factors reside in plastids. Many plastid anomalies are inherited maternally, i.e., via the egg cell, with only occasional or no introduction of the paternal-type plastids via the pollen tube. Analyses of inter-species crosses in the genus Oenothem lead to the conclusion that a species contains its own distinct plastid type. The different types of plastids are distinguishable by their phenotypic expressions in the presence of different chromosomal complements. Certain plastid types or clones that become green in the presence of their own chromosomal complement remain as undeveloped proplastids when introduced into a plant with an incompatible genotype, i.e., they cannot differentiate to chloroplasts, whereas other plastid types become green under the influence of this same genotype. I n the case of variegated plants, such as Mirabilis j a l a p , the colorless plastid cannot be made to become green no matter in what genotype it is tested. 4. GENESOF PLASTIDS

Studies on the development of chloroplasts of Euglena provide evidence for the presence of a large number of genes in a chloroplast. A variety of types of bleached clones of Euglena was isolated from a population of cells irradiated with U.V. light (or bleached by streptomycin or treated by heat. These clones varied in their ability t o synthesize carotenoids and porphyrins ( 4 2 ) . Such findings indicate that a number of genes in the Euglena plastids are sensitive to U.V. light. Two types of plastid genes were recognized: (1) genes that function in the differentiation of the proplastid to chloroplast-they appear to be highly sensitive to U.V. light because the plastids were readily “bleached” and only developed as tiny proplastids; (2) genes that function in the multiplication of the plastids and are relatively insensitive to U.V. light and to other “bleaching” treatments-this result was inferred from the fact that the proplastids were never completely lost from “bleached” cells. I n the latter cells, our electron microscope studies revealed bodies of 1-2 p diameter, which we considered to be proplastids. These bodies had outer double membranes but lacked internal lamellae. To follow the changes that occurred in chloroplasts after U.V. irradiation, the progeny cells of a single irradiated green cell were observed with the fluorescence microscope. All these cells, which had multiplied in red light on an organic medium, were seen to contain a well-developed vesicular system in which the chlorophyll, derived from the original green cell, could be detected by fluorescence (Fig. 5 ) . Evidently these

FIG.5. A colony of Euglena cells grown from a single green Euglena cell that had received ultraviolet, irradiation to cause all the progeny t o become colorless. At the left is the colony seen in white light; the granules are paramylum and the nuclei appear as colorless spheres in the center of the cells. At the right is the same colony seen in fluorescent light. The red fluorescence appears white and indicates that there is chlorophyll in a vesicular system throughout the cells (approx. ~ 6 0 0 ) .

0 56

56

* 3 * F

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vesicles represented the growing proplastid system in which chlorophyll was being diluted by multiplication of these proplastids. Normal darkgrown Euglena cells that had been exposed to visible light for a short time were also studied; such cells developed a small amount of chlorophyll. I n tlic fluoresccncc microscope, the chlorophyll was seen to be localized in a vcsicular system that resembled the vesicles in the colorless “bleached” progeny derived from U.V. irradiated green cells.

FIG.6. An electron micrograph of a U.V.-induced “bleached” Euglena cell. [From Schiff and Epstein (S).l M , mitochondria; PE, pellicle; PM, paramylum; E R , endoplasmic reticulum; G, Golgi body. The question marks on the photograph we interpret to represent mutated vesicular proplastids.

Schiff arid Epstein (28) made careful electron microscope studies of bleached Euglena strains. Some of their excellent photographs are presented in Figs. 6 and 7. Schiff and Epstein (28) conclude that plastids can be completely lost from bleached Euglena cells. We, however, consider that this has not been unequivocally demonstrated. We interpret as proplastids the structures that Schiff et al. label with a question mark and refer to as “amyloplasts.” The vesicular nature of these plastids, as seen in the fluorescence microscope (Fig. 5 ) , can also be seen in

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their electron microscopc pictuiw (Fig. 6 ) in which tlic plastids contain double membrane structures. I n higher plants, starch is formed within proplastids or chloroplasts. I n Euglena another polysaccharide, paramylum, is the carbohydrate storage form that appears to be synthesized adjacent to the pyrenoid of the plastid but on the outside of the plastid, and may become detached from the plastid. If paramylum synthesis is a function of plastids, then the presence of paramylum may be indicative

FIG.7. Electron microscope picture of a heat-induced “bleached” Ezcglena cell (symbols are the same as those of Fig. 6). [From Schiff and Epstein (28).1

of the presence of plastids. All of the bleached Euglena mutants contain paramylum. If Euglena cells cannot be found without remnants of plastids, it may mean that some plastid function, e.g., polysaccharide synthesis, is essential for the life of the cell. The studies of Stubbe (43) also lead to the inference that there are many genes in a plastid. He studied fourteen wild species of the subgenus Euoenothera and demonstrated the existence of five different types of normal plastids. These plastid types gave full green pigmentation in combination with nuclei of their own species. However, they reacted

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differently when combined with different, i.e., alien, nuclei of the other species, the plastids appearing then as light green or yellow or white, i.e., in various degrees of chlorophyll deficiency, depending on the particular combination of plastid and alien nucleus. By crosses with alien nuclei i t has been shown that plastid genes not only cause recognizable phenotypic differences in a plastid, but also affect a number of other morphological features of the plant, such as the rate of plastid multiplication, the shape of starch grains in proplastids of pollen, differences in dentation of the leaves, and vigor of stem growth, and perhaps are responsible for some cases of pollen sterility ( 4 1 ) . Plastid development is directly dependent on the activity of nuclear genes. Different nuclear genes are active in different kinds of cells of the same organism. The effect of these different nuclear gene activities is clearly seen in epidermal cells of a leaf or in root cells where proplastids are present that do not differentiate to chloroplasts, even when exposed to light.

C. Genetic Significance of Chloroplast DNA The evidence that DNA is present in chloroplasts is sufficient to implicate this DNA as the bearer of genetic information because of the established role of this substance in other biological systems. Furthermore, the following experimental evidence indicates that this DNA codes for the differentiation of the proplastid into the chloroplast. 1. STUDIESON BLEACHING OF Euglena

Mutations of the plastids of Euglena can be induced by irradiation with U.V. light. Lyman et al. (44) found that the U.V. action spectrum for this effect has a maximum a t 260 mp and that blue light reactivates the damaged cells. Such properties are characteristic of a DNA target. Gibor and Granick ( 4 5 ) showed that plastid mutations in Euglena result from irradiation of the cytoplasm. They were not induced by irradiating only the nucleus. The DNA targets for U.V. mutation of Euglena plastids are therefore in the cytoplasm. Studies by Edelman et nl. (46) and by Ray and Hanawalt (47) of U.V. bleaching indicate that U.V.-bleached Euglena cells “lose” a t least 90% of the chloroplast satellite DNA of normal green cells and that this “deletion” of DNA is directly correlated with the loss of the ability of the proplastids to develop into chloroplasts. However, another interpretation of their data may be that the mutated plastids are minute in size and few in number per cell; i.e., their DNA replicates infrequently as compared t o large normal green chloroplasts (see Figs. 6 and 7).

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PROTEIN SYNTHESIS Another type of evidence that indicates the biological activity of the DNA of the chloroplasts is provided by studies on biosynthetic reactions of isolated chloroplasts in vitro and the effect of antibiotics and DNase treatment on these reactions. RNA synthesis, which requires a DNA template, is inhibited by actinomycin D. Therefore, an inhibition of the incorporation of ribonucleotides into chloroplasts by this antibiotic is taken to indicate a DNA-dependent RNA synthesis occurring normally in chloroplasts. Furthermore, since protein synthesis is directed by messenger RNA, the inhibition of amino acid incorporation by this drug also leads to the conclusion that a DNA-dependent RNA synthesis is involved in this reaction. Iwamura (29) has summarized the experiments of various investigators that provide evidence for amino acid incorporation by isolated chloroplasts and by ribosomes from such chloroplasts. Evidence for the incorporation of RNA precursors into isolated chloroplasts and the inhibition of incorporation by actinomycin D was also summarized by Iwamura. It is clear, from all the incorporation studies and from inhibition studies of these reactions by DNase or actinomycin D, that the classical scheme of DNA to RNA to protein is functioning in the isolated chloroplasts. The fact that chloroplasts contain ribosomes that differ from cytoplasmic ribosomes is also suggestive of the semiautonomous nature of these organelles. The sedimentation of plastid ribosomes indicates that they are smaller ( 4 8 ) , and this was confirmed by electron microscopy ( 4 9 ) . Also, the base composition of the plastid ribosomes has a higher A U/G C ratio than cytoplasmic ribosomes (15). The DNA of C ratio than nuclear DNA. chloroplasts also has a higher A T/G

2. ROLEOF CHLOROPLAST DNA

+

+

3. THEDNA

IN

RNA

+

OF THE

AND

+

CHLOROPLAST Is NOT DERIVED FROM

THE

NUCLEUS

Indirect evidence for this idea has been deduced from the experiments on U.V. bleaching of Euglena (45).As was mentioned above, when the nucleus is shielded from irradiation and only the cytoplasm is irradiated, bleaching occurs; conversely, if only the nucleus is irradiated, no bleaching occurs. Because a healthy nucleus does not cure the irradiated cytoplasm and an irradiated nucleus does not bleach the plastids, it is concluded that the DNA of the rhloroplasts is not derived from the nucleus. Direct evidence that chloroplast DNA can he synthesized in the absence of the nuclcus comes from experiiiients on the alga Acetabularia

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( 2 4 ) . Bacteria-free enucleated cells were exposed to C1400,for 7 days. From chloroplasts removed from these enucleated cells, it was possible to isolate and purify C1*-labeled DNA. Enzymatic hydrolysis of this DNA yielded radioactive deoxynucleosides. It is thus apparent that enzymes for the synthesis of deoxynucleosides and for their phosphorylation and polymerization are present in Acetabularia cytoplasm. Whether any of these enzymes are derived from the nucleus directly or via mRNA, or from plastids or mitochondria, is not known. Another report suggesting that the DNA of the chloroplasts is synthesized independently of the nucleus is that of Chiang et al. (50) on the replication of DNA of Chlamydomonas. N15-Labeled zygotes were germinated in an N14 environment. Both nuclear and chloroplast (satellite) DNA were found to replicate in a semiconservative fashion, as determined by CsCl gradient analysis. However, the chloroplast DNA did not replicate in synchrony with the nuclear DNA. 4. AMOUNTOF DNA

IN A

PLASTID

The lower content of DNA reported for Acetabularia chloroplasts (Table I) and the tenfold or larger amount of DNA reported for chloroplasts of other species ( 1 X 10-14 gm) require explanation. The volume of a chloroplast of Acetabula~ais 1/20 to 1/50 of that of chloroplasts of higher plant species. If there is a relatively constant ratio of DNA to organelle volume as there is of nuclear DNA to volume of cytoplasm, one may expect that either DNA polyploidy or polyteny may have occurred in the larger chloroplasts. I n higher plants, the number of proplastids per meristematic cell of a leaf is approximately ten to twenty. By the time the cell has expanded fully, there are about 100 plastids, suggesting that each proplastid divides about three times in a palisade cell. Studies by Shipp et al. (SO) indicate that plastid DNA continues to replicate as the plastids multiply in the enlarging cells. Labeled phosphate is actively incorporated into the satellite DNA (presumably chloroplast DNA) of young expanding and greening leaves, whereas low incorporation is observed in nuclear DNA. Fully expanded mature leaves incorporate much less labeled phosphate into satellite DNA. These facts suggest that plastid 'DNA continues to replicate in the young plastids as they multiply and enlarge Init little or no 'DNA is formed in the mature plastid. If each proplastid has a unit nucleoid, will there be only one in the fully enlarged chloroplast, or more than one? That there may be several nucleoids per chloroplast is suggested by the finding of Yotsuyanagi (37) of several zones of DNA fibrils within R single plastid of Allium.

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5 . STABILITY OF PLASTID DNA

Is there any turnover of plastid DNA in mature cells? Perkins and Roberts (51) looked for turnover of chlorophyll in grasses and found no evidence for it, a result suggesting that chloroplasts in these plants remain unchanged during the lifetime of the leaf. However, Shlyk et al. (52) found that broad-leaf plants of angiosperms may have a chlorophyll turnover of as much as 1% per day. This turnover may indicate either replacement in the same chloroplast or a slow replacement of the chloroplasts themselves. It is not yet established whether mature chloroplasts can give rise to proplastids in cases where new cells may arise from cortical tissues, or whether proplastids form only by division of other proplastids. An interesting and perhaps related observation was reported by Higuchi et al. ( 5 3 ) . They studied the induction of bacteriochlorophyll synthesis in Rhodopseudomonas spheroides when these bacteria were transferred from dark-02 to light-H, conditions. Though no cell multiplication occurred, DNA synthesis was demonstrated by incorporation of C14-uracil into DNA. Inhibitors of DNA synthesis, such as mitomycin, phleomycin, and acriflavin, stopped this incorporation and blocked chlorophyll synthesis. The authors concluded that new DNA synthesis is required for chromatophore formation. This fact suggests that it may be worthwhile to seek for a chromatophore DNA that is unique and distinct from the rest of the DNA. The studies of Iwamura (29) on phased cultures of Chlorella ellipsoidia suggest that, a t the beginning of the interphaee when the cell and its cup-shaped chloroplast start to enlarge, satellite DNA of the plastid begins to increase, but nuclear DNA increases later.

D. The Problem of Labile or “Metabolic” DNA Several reports suggest that there may be a form of DNA in plants that is metabolically labile. However, the experimental data offered in support of this thesis are open to other interpretations. Iwamura (29) studied Chlorella during phased culture, and pulselabeled the cells with P3,0,. Athough no net DNA synthesis was observed, he reported that P32was incorporated rapidly and turned over rapidly in DNA components of the cell assumed to be derived from cytoplasmic organelles, such as the chloroplasts. The “DNA fractions” he isolated after phenol and chloroform-isoamyl alcohol treatment were subjected to CsCl gradient sedimentation, and the optical density a t 260 mp was determined i n the different fractions isolated from thc

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gradient. He found sixteen different, peaks in the region of the gradient between densities of 1.77 to 1.645 gm/ml. These peaks were characterized by the presence of P32and or U.V. absorption at 260 mp. At different phases of the cell’s life cycle, different peaks were found to contain different amounts of P I 2 . However, P-containing compounds, such as phosphoproteins or polyphosphates, may have contaminated the DNA preparations and these might be present in different quantities during the life cycle of a cell. A higher rate of P3? incorporation into DNA (satellite DNA), as compared to nuclear DNA, of cytoplasmic organelles in roots of higher plants has been reported by Sampson e t al. (54) and by Hotta e t al. ( 3 3 ) . A 4-hour exposure of the roots to P W , medium followed by 48 hours in a nonradioactive medium indicated that the nuclear DNA retains its spccific activity while the specific activity of satellite DNA decreases markedly. The total DNA of the growing root tissue increased 5% during the 48-hour “chase” period. Because satellite DNA accounts only for several per cent of the total DNA, if most of the net increase represents synthesis of new, nonradioactive satellite DNA, i t may account for the apparent reduction in specific activity of P32that was noted. The available data do not justify the claim for the presence of DNA that takes part in “metabolism.” Labeling studies with H3 nucleotides (Table 111) and with NI5 ( 5 0 ) suggest that the chloroplasts contain a stable DNA compatible with its genetic role. Where conditions occur in which cytoplasmic organelles may break down or turn over, as in Euglena placed in the dark for a week and then brought back into light, one may expect changes in both mitochondria1 DNA and plastid DNA.

111. Mitochondria1 DNA A. Evidence for the Presence of DNA in Mitochondria The presence of DNA in mitochondria has been demonstrated by procedures analogous to those considered for chloroplasts. We therefore categorize the evidence in a similar fashion. 1. DIRECT DETERMINATION OF DNA

IN

ISOLATED MITOCHONDRIA

The major problem in evaluating data obtained by analysis of isolated mitochondria is the question of the purity of the preparations. Contamination by fragments of DNA from disintegrated nuclei is difficult to eliminate. Experimental approaches used to minimize this contamination include the following:

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S. GRANICK AND AHARON GIBOR

( a ) Repeated washing of the isolated initochondrial preparations. For

example, Nass e t al. (55) washed mitochondrial preparations up to four times and found that the amount of DNA in the mitochondria remained practically constant. ( b ) Treatment of isolated intact niitocliondria with DNasc prior to isolation of DNA from them. This DNase treatment removed extraneous DNA, but did not hydrolyze the intramitochondrial DNA (8, 1 0 ) .

Convincing evidence that the mitochondria contain their own DNA is the demonstration that this DNA is qualitatively unique and disTABLE IV BUOYANT DENSITIESOF MAJOR(NUCLEAR) A N D SATELLITE (MITOCHONDRIAL) DNA's Source

Nuclear

Yeast Yeast Neurospora Leishmania Leishmania Tetrahymena Rat liver Guinea pig Sheep heart Chicken liver Chicken embryo liver Chicken embryo heart

1.698 1.700 1.713 1.721 1.716 1.685 1.703 1.700 1.704 1.701 1.698 1.698

Mitochondria1 Reference 1 .684a 1.685n 1.701 1 ,699 1.703 1.671 1 . 701n 1 . 700a 1.714= 1.707" 1.707a 1.707"

7 9 8 66

67 68 69 9 9 9 60 60

Double-strandedness of this DNA was shown by hyperchromicity and density changes after heat denaturation.

tinguishable from nuclear DNA. An example is the isolation of mitochondrial DNA from Neurospovu crassa by Luck and Reich (8). Most of the nuclear DNA was removed by treating purified mitochondrial preparations with DNase, following which the DNA was isolated from the mitochondria and analyzed by sedimentation in a CsCl density gradient. The buoyant density of the mitochondrial DNA was 1.701 gmJ ml while that of the nuclear DNA was 1.712. Proof that the material banding a t a density of 1.701 was DNA came from its U.V. absorption, its ability to complex with actinomycin D, and its digestion by DNase. The DNA from mitochondria of different organisms was studied by similar techniques of isolation and analysis. The data from such studies are summarized in Table IV.

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Mitochondria1 DNA from some mammalian tissues, namely rat liver, beef heart, beef liver, and mouse liver (9),is identical in density with the nuclear DNA from the same tissue. The DNA of mitochondria and nuclei of all guinea pig tissues is identical in density. However, the riiitochondrial DNA of the above species differs from nuclear DNA in its renaturation after heat denaturation. Corneo et al. (9) suggested that the ease of renaturation arises from the greater homogeneity of the mitochondria1 DNA as compared to the nuclear DNA. They suggested that this attribute can be used as a distinguishing characteristic to identify mitochondria1 DNA in those cases where its density is the same as that of nuclear DNA.

EVIDENCE FOR hf ITOCHONDRIAL DNA Most mitochondria do not stain in the Feulgen reaction or with similar cytological staining reactions for DNA. However, some specialized mitochondria do so. Chevremont (61) found that if chick fibroblasts are grown for 1 day a t 16°C or are treated with DNase, their mitochondria accumulate large amounts of DNA. Such mitochondria give positive Feulgen staining reactions; furthermore, treatment of the fixed preparations with DNase causes the removal of the reactive material. Similarly, Steinert (62) found that the kinetoplast of hemoflagellate protozoa gives a positive Feulgen reaction. DNase treatment removes the stainable substances. Electron microscope studies by Steinert on trypanosomes (62) and by Rudzinska et nl. (63) on Leishmania revealed that the kinetoplast is a specialized mitochondrion. This organelle differentiates in conjunction with the metamorphosis of the protozoan from its intracellular parasitic form to the free-living aerobic form. The kinetoplast develops during the metamorphosis into a single large convoluted mitochondrion with typical cristae. With its relatively high DNA content, it probably represents onc especially large mitochondrion per cell in which the DNA is polytenic, as contrasted to the usual case of cells, with numerous mitochondria each with its own small DNA complement. The same regions of the kinetoplasts that stain chemically for DNA contain fibrillar material 25 k in diameter. These fibrils are absent after DNase digestion (63). Nass et al. (64) studied the mitochondria from over forty different sources and found similar fibrils in all of them. Because of their similarity to the fibrils found in the nucleoplasm of Ijactcria and because of their sensitivity to DNase, thew fibrils are considered to hr DNA. Tritium-labeled DNA precursors, fed to intact cells, are incorporated not only into the nucleus but also into mitochondria. That the in2.

CYTOLOGICAL

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corporated label is associated with newly synthesized DNA was demonstrated by showing that the label is not removed by cold acid or by RNase treatment. The labeled material removed by DNase is considered t o be DNA synthesized after the addition of the labeled precursor. Table V summarizes several such labeling st,udies. TABLE V INCORPORATION OF LABELED THYMIDINE INTO DNA Organism

OF

MITOCHONDRIA

Results

Labeled thymidine is incorporated into the kinetoplast; the label is sensitive to DNase (66) Leishmania Labeled thymidine is incorporated into the kinetoplast (67) Tetrahymena Labeled thymidine is incorporated into mitochondria; the label is sensitive to DNase or hot acid extraction (66). Incorporation of thymidine into mitochondria is independent of DNA synthesis in the nucleus. The labeled DNA is stable for at least four generations (68). Continuous mitochondrial DNA synthesis is independent of nuclear DNA synthesis (67).Localization of label in the mitochondria confirmed b y electron microscopy radioautography. Mitochondria1 DNA is stable as no loss of activity was found in subsequent cell divisions (68) Chick embryo Thymidine incorporation into mitochondria is active in cells pretreated with DNase or low temperature. The incorporated label is sensitive to DNase digestion (61) Swiss chard leaf Thymidine is incorporated into mitochondria as well as into plastids. It is DNase-sensitive (32) Physarum polycephalum Thymidine incorporation is independent of nuclear DNA synthesis (70) Rat liver Thymidine is incorporated at a higher specific activity in mitochondria than in nuclear DNA. Mitochondrial DNA synthesis is independent of the mitotic activity of the tissue. DNase removes the incorporated label (71) Trypanosovne

The labeling studies on synchronized Tetrahyrnena cells (58, 67, 68) indicated not only that DNA is present in the mitochondria but also that the synthesis of this DNA proceeds independently of the synthesis of nuclear DNA. Studies of pulse-labeling with tritiated thymidine followed by growth in nonradioactive medium revealed that the tritiumlabeled DNA of the mitochondria was halved with each cell division. This indicates that the mitochondrial DNA is stable in the growing cells. The experiments of Reich and Luck ( 7 l a ) on Neurospora have shown by the Meselson technique (72) that mitochondrial DNA replica-

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tion is semiconservative, as with bacterial DNA and nuclear DNA of higher plants and animals; i.e., one half of the original DNA strands are passed on to each of the two nucleoids.

B. Hereditary Factors of Mitochondria Mitochondria, like plastids, are thought to be endowed with a degree of genetic autonomy. It is therefore essential to establish whether their DNA functions as an information carrier. The hypothesis of mitochondrial genetic autonomy is based on the following genetic and cytological observations. 1. PHYSICAL CONTINUITY

Evidence that mitochondria arise by the growth and division of preexisting mitochondria was shown in electron microscopy studies of a small unicellular alga, Micromonas (73). This organism contains one nucleus, one chloroplast, and one mitochondrion. The three divide in synchrony a t the time of cell division. This is proof of the continuity of these organelles from one cell generation to the next. Experiments of Luck (74) on a choline-deficient strain of Neurospora also led to the conclusion that mitochondria arise from pre-existing mitochondria and not de novo. The mitochondria1 membranes were labeled during the early logarithmic phase of growth. B y pulse-feeding of radioactive choline, the newly formed labeled lecithin was found to be bound tightly to the membranes and it did not turn over in subsequent divisions. When the mitochondria increased in number as the cells divided, all of them were found to contain radioactive label. Therefore, the new mitochondria did not arise de novo from nonradioactive precursors but as a result of growth and division of pre-existing labeled mitochondria. I n other experiments, Luck (75) showed that the mitochondria of a choline-deficient Neurospora strain could be made to vary in density. The mitochondria of cells grown on a medium low in choline were heavier than those derived from cells grown on a high-choline medium. When a growing culture was transferred from low to high choline, a gradual decrease in the density of all the mitochondria was observed. This result is compatible with the notion of growth and division of the original organelles. If mitochondria arose de novo, two bands of different densities would have been expected. 2. NON-MENDELIAN INHERITANCE

Ephrussi and co-workers in their classic studies of yeast genetics (76) showed, by analysis of crosses between various yeast strains, that one type of “respiratory deficiency” results from mutations of cyto-

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plasmic factors. These mutations caused the production of poorly developed, nonrespiring mitochondria. Respiratory deficiencies were induced by U.V. light with an action spectrum maximum a t 260 mp; blue light photoreactivated the U.V. damage. Various basic drugs known to interact with DNA were also effective in inducing the respiratory deficiency syndrome (77). 3. MITOCHONDRIAL CLONES Several reports indicate that mitochondria within a single cell may differ from each other in certain properties. Avers et al. (81) found that certain clones in acriflavin-induced respiratory variants of yeast contain a t least two types of mitochondria, some of which are cytochrome oxidase positive and others cytochrome oxidase negative. I n a study of rat heart tissue, Ogawa and Barrnett (82) found that different mitochondria varied in their rate of reduction of tetrazolium salts; adjacent mitochondria were frequently found, one of which contained a heavy deposit of formazan over all its cristae while the other had no such deposit. I n both these studies, electron microscopy was used to localize the metabolic end products in specific mitochondria. Two cases of human pathology have been reported in which a muscular disorder was associated with the presence of morphologically abnormal mitochondria only in skeletal muscles and in no other tissues (83, 84). I n both cases normal mitochondria were also present in the afflicted tissues. Whether these abnormal mitochondria are genetically different or represent a stage in degeneration is not known. The buoyant density determinations of mitochondrial DNA also suggest the existence of clones, but it has not yet been shown that these density differences are the result of changes in base composition rather than in the presence of unknown components attached to the DNA. Reich and Luck (71a) found two mitochondrial DNA populations with densities 1.698 and 1.702 in Neurospora crassa whereas in N . sitophila the densities were 1.692 and 1.702. They demonstrated that the DNA of the mitochondria was inherited exclusively through the maternal parent by determining the mitochondrial DNA densities from appropriate crosses. Similarly with yeast, clones of mitochondrial DNA and maternal inheritance of this DNA were found by Mounolou et al. (85).A normal strain of yeast had a mitochondrial DNA density of 1.687; a cytoplasmically inherited yeast strain lacking cytochromes a, a3 and b had a mitochondrial DNA density of 1.683, and another cytoplasmically inherited yeast strain, “suppressive,” had a mitochondrial DNA density of 1.695.,I n yeast, the mitochondrial DNA constitutes about 10% of the total cell DNA.

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A demonstration of the transmission of biocheinical properties by mitochondria was achieved by Diacumakos et al. (7’8). They injected mitochondria obtained from a cytochrome-deficient, slow-growing Neurospora mutant into the hypha of a strain normal with respect to growth rate and cytochromes. The mycelium that developed from the injected hypha was found to be slow-growing and cytochrome-deficient. The transmission of these characteristics was not achieved by injection of nuclei from this slow-growing mutant. It is apparent, therefore, that the injected mitochondria transmitted their properties to the host cell. The details of the genetic interaction between the injected mitochondria and the original host mitochondria are not known. The facts suggest that the defective organelles either outgrow the normal organelles or suppress their multiplication. Variations among different respiratory-deficient strains have been observed in yeast. Ephrussi et al. (‘79)described two types of respiratory-deficient cells, suppressive and neutral. When neutral respiratorydeficient cells were crossed with cells containing normal mitochondria, all the progeny had normal respiration, as if the normal mitochondria had replaced or perhaps “cured” the defective mitochondria. Progeny from crosses of suppressive respiratory-deficient cells and normal cells were all respiratory-deficient. There are strains that are intermediate in their suppressiveness. A cross of such an intermediate strain to a normal strain results in a fraction of the progeny being deficient. From single-cell cloning experiments, Ephrussi and Grandchamp (80) concluded that this “intermediate” property is due to intrinsic factors in the cells and not to a mixed population of suppressive and neutral cells in the population. It is possible that a mixed mitochondria1 population is responsible for the intermediate state. Besides the genes that control the respiratory capacity of the mitochondria, other genes (either in the mitochondria or nucleus or both) controlling the continued multiplication of the defective mitochondria in the respiratory-deficient cells, are postulated. 4. GENESOF MITOCHONDRIA

Woodward and Munkres (86) have obtained evidence for specific gene mutations in the mitochondria of “poky” mutants of Neurospora. They isolated the mitochondria1 structural protein (MSP) of Criddle (23,000 in01 wt) froiii Neurospora initochondria and dctcrmined its amino acid composition. Similarly, they isolated this protein from two poky mutants. The mutant mi-1 was found to contain one less tryptophan and one more cysteine residue per mole of MSP than the wild type. The mutant mi-3 had onc less tryptophan. The authors conclude that these

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changes in MSP result from mutated genes of mitochondria1 DNA. This experiment provides the first evidence for a single gene change in a cytoplasmic organelle that is expressed as an inheritable phenotypic change in a specific protein, if it is indeed one and not a mixture of proteins.

C. Genetic Significance of Mitochondria1 DNA That the DNA of the mitochondria is the carrier of its genetic information is indicated by the following experiments. 1. MITOCHONDRIAL DEFECTS

As mentioned above, mitochondrial defects are induced in yeast by agents such as U.V. light and acridine dyes, which are known to react with DNA. By sedimentation in CsCl, Corneo e t al. (9) analyzed DNA from a normal yeast strain and a respiratory-deficient strain. The DNA of the respiratory-deficient strain appeared to lack a prominent satellite band. It appears therefore that the loss by the mitochondria of their respiratory potential is correlated with a deletion of much of the DNA of these organelles. However, the quantitative reduction in the amount of satellite DNA might be due to the decrease in the number of promitochondria per cell. More recently, Mounolou et al. (85) have found that respiratorydeficient yeast strains do contain a satellite DNA. This DNA differs in density from the normal satellite DNA (see above). 2. SEMICONSERVATIVE REPLICATION

By labeling with heavy isotopes, mitochondrial DNA is found to be replicated in a semiconservative fashion, according to Reich and Luck ( 7 l a ) . This mode of replication is characteristic for the genetic DNA of bacteria and is compatible with the function of this DNA as a stable genetic substance. The stability of the mitochondrial DNA was indicated by studies on Tetrahymena in which it was shown that Hathymidine incorporated into the cytoplasmic organelles is not lost from the cells in subsequent cell divisions (58, 68). 3. DNA

AND THE

SYNTHESISOF RNA

AND

PROTEIN

Biological activity of this DNA is also shown by the synthesis of RNA and protein by isolated mitochonclria. Actinomycin D or DNase treatment blocks these biosynthetic reactions. Actinomycin D is known to interfere with a DNA-directed RNA synthesis; thus inhibition of ribonucleotide incorporation by this antibiotic is taken to indicate that a DNA-dependent RNA polymerase reaction is involved. Similarly, inhibition of ribonucleotide incorporation after DNase treatment impli-

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cates the DNA in coding for the polymerization reactions. Results of such studies are presented in Table VI. The experimental results listed in Table VI are compatible with the assignment of a genetic role to the DNA of the mitochondria. A serious objection to studies on the incorporation of labeled precursors into isolated organelles has been the possibility that contaminating microorganisms may have been present in the incubation mix-

ROLE OF DNA

IN

TABLE V I BIOSYNTHETIC REACTIONS O F ISOLATED MITOCHONDRIA

Organism

Results ~~

~~

~

Neurospora

Actinomycin D inhibits G T P incorporation. DNase and RNase do not inhibit incorporation in intact mitochondria. All four ribonucleoside triphosphates are required for incorporation into RNA. (8) Yeast ATP incorporation is inhibited by actinomycin D. All four triphosphates are required for incorporation into RNA. DNme and RNase do not inhibit intact mitochondria. Amino acid incorporation is sensitive t o oligomycin, antimycin A, acriflavin, and chloramphenicol. There is some inhibition of amino acid incorporation by actinomycin D (7). Leucine and phenylalanine incorporation is inhibited by actinomycin and by acriflavin (87). Uridine incorporation is inhibited by actinomycin and acriflavin. Three different labeled high molecular weight RNAs were isolated after uridine incorporation (88) Lamb heart ATP incorporation is inhibited by actinomycin D. Amino acid incorporation is inhibited b y actinomycin D. RNase and DNase inhibit the incorporation of amino acids into sonicated mitochondria but not into intact mitochondria (89) R a t tissues Nucleoside triphosphates are incorporated into RNA. All four nucand leotides are required. Actinomycin D, acriflavin, and DNase inhibit Pigeon tissues the incorporation (90)

tures. According to Van der Dehn (91) et al., no amino acid incorporation is found in sonicated mitochondria1 preparations prefiltered to remove bacterial contaminants. However, the kinetics of several of the incorporation experiments we have cited rule out the possibility that bacterial growth was responsible for the results obtained. I n these experiments ( 8 ) , the incorporation reaction slowed down after a short time (less than 30 minutes) or stopped altogether. I n other experiments, complex requirements were necessary for incorporation to occur. For example, all four ribonucleotides were necessary to demonstrate GTP incorporation by Neurospora mitochondria (8). These results suggest that experiments on the conversion by organelles of precursors into high

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rnolecular weight substances cannot all bc disniissed as resulting from contamination due to bacterial growth. Complementation experiments between mitochondrial RNA and nuclear DNA of the mouse have led Humm and Humm (92) to conclude that thcre are two types of mitochondrial RNA. One typc forms cornplementary hydrogen-bonded strands with nuclear DNA ; the other does not. Pretreatment of the mitochondria with RNase ruled out contamination with cytoplasmic RNA. If this result is confirmed, i t indicates that mitochondria may obtain coding information for protein synthesis not only from mRKA of the mitochondria but also from mRNA of the nucleus. 4. REPLICATION OF MITOCHONDRIAL VERSUS NUCLEAR DNA

Under some conditions mitochondrial DNA, like the chloroplast DNA, incorporates labeled precursors a t a much higher rate than the nuclear DNA. Schneider and Kuff (93) found that incorporation of thymidine into mitochondrial DNA of rat liver cells is a t least ten times faster than the incorporation into nuclear DNA. In Tetrahymena ( B Y ) , thymidine incorporation into mitochondrial DNA can be detected throughout the life cycle of the cell while nuclear DNA synthesis is restricted to a short period of this cycle. The replication of mitochondrial DNA in Neurospora is metabolically independent of nuclear DNA because precursors for synthesis of the two kinds of DNA are drawn from different pools. The precursor pools differ in their rate of turnover and dilution by exogenous nitrogen sources (Yla). These observations indicate that the replication of mitochondrial DNA is independent of nuclear DNA replication and that different control mechanisms operate for the two DNA systems of the cell. 5. TURNOVER OF MITOCHONDRIAL DNA

It may be expected that mitochondria may turn over in certain types of cells. For example, in Euglena cells grown in the dark and then transferred to light, the electron microscope pictures revealed a marked diminution in the volume occupied by the mitochondria a t the same time that the chloroplasts enlarged. Brandes et al. (94) noted the breakdown of mitochondria in digestive vacuoles of such cells. Similarly, Elliott and Bak (96) studied the fate of the mitochondria in Tetrahymena. During the logarithmic growth phase of the culture, the organelles were located toward the outer surface layer. When cell multiplication ceased, a large number of mitochondria concentrated in

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the central part of the cell, Later many of these mitochondria were found inside vacuoles in different stages of breakdown. Figure 8, from the work of Elliott and Bak, is an example of such disintegrating mitochondria in Tetrahymena. Fletcher and Sanadi (96)reported that r a t liver mitochondria have a half-life of about 10 days as estimated by the rate of disappearance of three simultaneously incorporated pulse-labeled isotopes. Similarly,

FIG.8. Electron microscope picture of a Telrnhymena cell. In the center is a vacuole containing mitochondria a t different stages of disintegration. [From Elliott and Bak (961.1

Neubert et al. (71) estimated a half-life of 8-9 days for pulse-labeled rat liver mitochondria1 DNA. The half-life of nuclear DNA of the same tissues was about 20 days. Therefore, the life of liver mitochondria is considerably shorter than the turnover of the liver cells themselves. The loss of mitochondria of erythrocytes as the cells become mature has been known for a long time. No turnover of mitochondria was observed in healthy growing hreurospora cells ( 7 4 ) . What may be the significance of this turnover of mitochondria and their DNA in mature cells of higher organisms is not a t all clear. Nor is

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S. GRANICK A N D AHARON GIBOR

it known whether this turnover will be found to be of general occurrence in these cells.

IV. Centriole DNA Centrioles are cytoplasmic bodies that are implicated in the formation of new centrioles, basal bodies, astral fibers, spindle fibers, cilia, flagellae, and retinal rods and cones (96a, 96%). Suggestive evidence is beginning to appear that information for the synthesis of these cytoplasmic components is derived from a tiny DNAgm doublecontaining unit. This unit may contain about 2 X stranded DNA (16), it lacks a membrane, and may be 700 A or less in diameter when organized into a compact granule. We designate this unit as a “procentriole.” Other synonyms that have been used for this body are procentriolar granule and basal granule. There is much work in progress concerning this unit. The following interpretation of the available data is based on the tentative hypothesis that the procentriole gives rise to a differentiated centriole (Fig. 9). The terms, cilia and flagella, are best defined in terms of their characteristics in the Ciliata and Flagellata, two of the major classes of Protozoa. In the Ciliata, the cilia are usually small, numerous, have a coordinated beat and arise from the pellicular surface in rows or kineties. The centriole a t the base of each is called a basal body or kinetosome. It does not appear to be derived directly from a centriole associated with the nucleus. In the Flagellata, flagella are usually few in number, longer and thicker, and have a more complex sheath. The centriole a t the base of each is derived from a centriole associated with the nucleus. I n protozoa as well as in higher plants and animals where cilia and flagella occur, the basic elongated motile structure has the same fundamental pattern of nine double microtubules on the periphery and two microtubules in the center. At the base of the ,motile structure is a centriole, which shows considerable variation in its organization among various species (96c). Because of the essentially similar structures we use the term “cilium” to designate cilium of flagellum; and the general term “centriole” to designate centriole or basal body or kinetosome.

A. The Function of the Procentriole This organelle is suggested to give rise to all cytoplasmic microtubules (Fig. 9). All the microtubules of a cell are constructed on the same basic plan. They are 230-270 A in diameter with a lumen 100 A wide. The dense 70 A wall of the tubule consists of ten to thirteen microfilaments each 35 A in diameter, arranged cylindrically and parallel to

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173

the length of the niicrotubulc. The microtubule may bc short, or many microns long, and appears to be straight and rather stiff (97, 98). Microtubules, variously arranged, may be found in different parts of a cell. ( a ) The microtubules may be organized in the shafts of cilia in thc charactcristic concentric arrangement of nine pairs and two central ones. ( b ) They may be organized into the astral and spindle structures observed during cell division. (c) They may be present during interphase, lying free in the cytoplasm usually adjacent to the protoplast membrane. It is conjectured that the microtubules take part in the for-230-270

A-

centriole

'.I

I, I,

I'

!!

I

;

,

,

*

Microtubule with

10-13 microfilaments

FIG.9. Left, structure of microtubule; right, cell in interphase with two centrioles and their companion procentrioles.

mation and maintenance of cell form as wcll as in cytoplasmic streaming or movement in both plant and animal cells. Auber (99) has observed microtubules in differentiating muscle ; they are incorporated probably as thick filaments into the developing A band. Ledbetter and Porter (97) and Pease (98) have suggested that all muscular movement and nonmuscular movement, whether in cilia, in the spindle, or in cytoplasmic streaming, occurs in connection with these microtubules. The origin of the microtubules of the cilia is from a centriole or basal body derived from a procentriole. The origin of spindle and astral fibers is from centrioles or a region surrounding the centriole, the centrosphere. The origin of microtubules or cytotubules in the cytoplasm is unknown but one hypothesis (98) is that they represent microtubules of the spindle that become dispersed during the last stages of

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cell division. Possible support for this idca is that protein, serologically similar to that derived from the isolated spindle, appears to be present in embryonic cells during interphase (99a).

B. Differentiation of the Procentriole A sequence of procentriole changes may be inferred from various current studies (2, 98, 96c, 100-103). The following description should be regarded as a tentative interpretation of the available data. A procentriole may divide to form a daughter procentriole. Or, the procentriole may differentiate t o form a cylindrical ring, about 1500 A wide by 1500 A long, called a basal body or centriole (Fig. l o ) , one end of which is closed and the other open. The wall of the centriole may contain nine triplets of microtubules arranged cylindrically in a pinwheel pattern; the outermost microtubule of the triplet does not extend further than the basal plate. Within the cylindrical centriole is a dense matrix (101). The centriole presumably contains the DNA of the procentriole. More recent work (Fawcett, 104) indicates that the centriole and the region surrounding it are highly structured. The centriole appears to be surrounded by 3 4 concentric rings of dense materials; a number of “satellite” granules, 500-700 A diameter, have been seen from which microtubules arise. In connection with the centriole there may arise (a)a new procentriole, ( b ) astral and spindle microtubules, ( c ) a cilium, or ( d ) a modified cilium of the outer segment of a retinal cell (Fig. 10). At the end of cell division each new daughter cell has two cylindrical centrioles that lie in a characteristic perpendicular position to each other close to the nuclear membrane. During interphase, a stainable procentriole appears adjacent to each centriole, presumably having been formed in connection with the nucleoid of the centriole (Fig. 9 ) . I n preparation for mitosis, one centriole and its companion procentriole migrate to the opposite pole of the cell. At metaphase, the astral and spindle microtubules arise in the region of the two centrioles a t either pole. Simultaneously, each procentriole differentiates into a centriole. Thus a t telophase, when the cell begins to divide, there are two centrioles a t each pole, i.e., two tiny cylindrical units lying perpendicular to each other. One is derived from the mother cell and the other is newly differentiated from the procentriole. I n cells that develop a cilium, the centriole that comes to lie closest to the cell membrane is the one that will form the cilium. This cylindrical centriole first increases in length to about 5000 A. It is probable that its DNA codes for certain substances that, together with other substances from the cytoplasm and cytoplasmic membrane, are used in cilium formation.

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When the elongating centriole touclics the cell nicmbrane a basal plate develops a t that end (Fig. 10). The cell mcmbrane bulges as a bud over the basal plate (10%’).This bud enlarges into a fingerlike extension of the cell membrane. The two inner microtubules of each triplet now grow through the basal plate. The two ccntral microtubules develop only above the basal plate. Vesicles, originating by pinocytosis of

FIG.10. DNA of the procentriole (1) provides information for the differentiation of the centriole (2). The centriole differentiates further into a cilium (3) and (4) or into an outer retinal rod segment ( 5 ) .

the cell membrane, supply some materials for the enlarging cilium. It is not known whether most active growth and organization occurs in the region of the centriole and basal plate, or a t the tip of the cilium. I n the development of the outer rod segment of the retinal cell (Fig. l o ) , the protruding fingerlike extension from the centriole enlarges. Then numerous pinocytotic vesicles pinch off from this extension. The vesicles fuse to form flattened discs that layer together, resulting in the photoactive elements of the outer segments of rods and cones. A modified

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S. G R A N I C K A N D A H A R O N GIBOR

flagellar structure, originating from the ccntriole in the main body of the cell (i.e., the inner segment), presumably serves for transmission of impulses from the photoactive discs. The cilia in Paramecium do not arise from the nuclear centriole but rather from centrioles that are localized in the pellicle. However, in the flagellate Barbulanympha, the nuclear centriole not only gives rise to the astral and spindle microtubules but also to a procentriole that in turn forms the ciliary apparatus of this organism (103). More precisely, Barbulanympha has a long, spindle-shaped centriole 30 p long by 5 p wide. At cell division the internal end towards the nucleus forms the astral and spindle microtubules; and the outer end buds off a procentriole. This procentriole, near the cytoplasmic membrane, serves as a reproducing unit. It buds off a series of procentrioles from which subsequently arise an organized flagellar apparatus and it also enlarges into a spindle-shaped body that projects inwards, and becomes identical with the original long centriole. The internal end of this new centriole a t cell division also develops astral and spindle microtubules.

C. Evidence for DNA in Procentrioles Randall and Disbrey (16) have established the presence of DNA units in the pellicle of the ciliate, Tetrahymena pyriformis. In this organism, each cilium arises from a centriole or basal body which in turn is derived from a procentriole. The centrioles, i.e. kinetosomes, are aligned in characteristic rows (kineties) in the pellicle and it is assumed that one centriole in a row arises indirectly from an adjacent centriole. The pellicle coat of the organism may be removed, stained with acridine orange, and examined with a fluorescence microscope. When the pellicle from an organism in a stage just prior to cell division was examined in this way it was found that its centrioles fluoresced green. This is evidence that the centrioles contain double-stranded DNA or RNA. After digestion with DNase, but not RNase, the green fluorescence disappeared. Confirmation of the presence of DNA in the regions of the basal bodies was obtained by autoradiographic studies with tritium-labeled thymidine. It was estimated that the DNA per centriole was about 2 X gm and that the over-all DNA in all the centrioles of this organism was about 1%of the DNA of the macronucleus. The greenish fluorescence of the centrioles stained with acridine orange was not observed just after cell division, but during cell enlargement it gradually increased in intensity and became maximal shortly before cell division. A reasonable interpretation of these results, offered by Randall and Disbrey, is that after cell division, when the young cells begin to enlarge, the procentriole DNA is unfolded and dis-

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persed. I n this form it is active in replication of DNA and in coding for ciliary material. However, in this unfolded form it is too diffuse to be seen by the fluorescence technique. As the cell nears maximal growth the procentriole DNA condenses into a tight packet, detectable by its fluorescence in acridine orange. Although analytic methods had suggested that DNA might be present in the pellicle, the possibility of contamination with nuclear DNA dissuaded acceptance of this idea. More recently, Hoffman (105) analyzed fractions of pellicles containing centrioles. He estimated that they contained 1.2-2.6% RNA by weight but because of the possibility of nuclear DNA contamination he estimated that the DNA content of the centrioles was as high as 1%or as low as 0% by weight. At present there exists no direct evidence that the centrioles associated with the nucleus contain DNA. However, it is assumed that, because centrioles of the pellicle of ciliates contain DNA, homologous centrioles adjacent to nuclei will also contain it. I n animal cells and flagella-producing cells of certain plants, a deeply staining centriole granule and a more diffusely staining region around the centriole (the centrosphere) are seen by light microscopy; such cells produce both astral and spindle microtubules; the latter microtubules connect up with the attachment regions (kinetochores) of the chromosomes. However, all other plant cells apparently lack a procentriole granule and its existence is doubtful in many animal cells. In plant cells during cell division, spindle tubules are formed that issue from a diffusely staining polar region. Perhaps in plants and many animal cells the procentriole DNA is more diffuse, and cannot be detected by conventional staining methods. Another but less likely explanation is that coding for spindle microtubules in plants may reside in nuclear DNA. A search for filamentous DNA by electron microscopy in the region of the centriole from which the spindle microtubules extend may help to establish the presence or absence of procentriole DNA.

D. Genetic Aspects of the Procentriole The procentriole and its differentiated form, the centriole or basal body, is a semiautonomous organelle. Only a procentriole (or centriole) has the ability to reproduce a centriole, a hypothesis first proposed by Lwoff (106). Reproduction of a procentriole is independent of the nucleus and its DNA is not derived from the nucleus. This conclusion may be inferred from an experiment by Moore (107)on enucleated eggs of Rana pipiens. I n this enucleated egg, even in the absence of a nucleus, cell division can occur many times during the cleavage period. At each division centrioles multiply and carry out their customary functions.

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The centriole, like the plastid and mitochondrion, is a self-reproducing cytoplasmic organelle that undergoes differentiation. Unlike the plastid and mitochondrion, the centriole is not separated from the cytoplasm by a membrane, and the centriole appears to differentiate into a number of different kinds of cytoplasmic structures, all of which contain tubules of a similar structure, i.e., microtubules (Fig. 9). Ciliates have a surface coat or pellicle that consists of rows of centrioles or basal bodies from each of which a cilium arises. Because of the large number of centrioles and their arrangement, the ciliates have been extensively studied by Sonneborn (108) and others with regard to inheritance and perpetuation of pellicle organization. It has been found, for example, that when a patch of pellicle is removed by microsurgery and rotated and implanted back into the pellicle, the cilia beat in the direction opposite to the other cilia, and this artificial organization is inherited by the daughter cells. Many experiments have established the replication of ciliary precursors and ciliary patterns from one generation to the next. Pellicle pattern is determined by the pre-existing state of pellicle organization. Organization of ciliary bodies takes information from pre-existing ones. Attempts have been made to find out whether procentrioles from different parts of the pellicle are genetically different. Pellicles of the basal region have been transplanted to the oral region, or removed completely. It is concluded from these studies that the procentrioles, however different in function and regardless of the patterns in which they are organized, are essentially equipotent. The various cortical patterns can arise in the absence of a pre-existing pattern that, although useful, is not essential (109). On this basis we may assume that all the procentrioles of a pellicle contain nucleoids of DNA with identical genes that code for the architecture of the individual cilium, but other information may be required to establish ciliary patterns of the pellicle. Because of the large number of centrioles in a ciliate pellicle, one may expect mutations to arise that will result in the production of different clones of centrioles that may possess modified cilia. For example, it may be possible to find a row of nonmotile cilia or a row of procentrioles that cannot make flagella. As yet, no reports have appeared to indicate that such changes occur, but these may be worth looking for. I n Barbulanympha, after sexual fusion, the flagellar apparatus is destroyed and a new one, with all of its procentrioles, is derived from a single internal centriole derived from the female (104). I n this case, only one clone of procentrioles is to be expected. Mutations of procentrioles also may be sought in those cells that have one or two cilia and can form a zygote. Paternal inheritance is

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usually the rule in centriolc inheritance, but this depends on the species ( 9 6 b ) . I n this regard, the following experiment is of interest. Randall et al. (110) found a nonmotile mutant of Chlamydomonas reinhardi that lacks the two central niicrotubules in each of the two flagella. Tetrad analysis of crosses between this iiiutant and the wild type gave a 1:l segregation. The authors considered this result to indicate that the nonmotile flagella were caused by a nuclear gene mutation. However, in C. rknhardi the fusion in zygote formation is between isogametes; if the centrioles of both gametes were retained then a n apparent 1:1 segregation might result that might be indicative of a mutation in centriole DNA rather than in nuclear DNA. Defective flagellar movement caused by nuclear gene mutations has been reviewed by Ebersold ( 1 1 0 ~ ) .

V. The Role of Cytoplasmic Nucleoids in Inheritance As noted above, there are many cytoplasmic organelles of the same kind in a cell. Each organelle has its own nucleoid. Each nucleoid contains enough DNA to code for several hundred genes. This means that there are many copies of the same kinds of cytoplasmic genes in a cell. This gene redundancy has several implications.

A. Redundancy and Genetic Analysis One implication, which we have already discussed, is that genetic analysis of cytoplasmic genes is made difficult. For such studies, methods of analysis different from those that depend on Mendelian segregation must be used. Mention has been made of the recognition of phenotypic differences among plastids of the same cell by observation of their size, pigmentation, starch content, and distribution. Phenotypic differences of mitochondria in the same cell have been recognized by their reducing or oxidizing abilities in which insoluble materials that become recognizable on electron microscope examination have been deposited within them. The possibility of seeking for differences in lines of cilia in protozoan ciliates has also been suggested. Another method to study differences of genes between organelles is to transfer an organelle into an environment of an alien nucleus, as was mentioned in the case of Oenothera. This permits the recognition that plastids of closely related species are different genetically. I n variegated plants, segregation of clones of plastids occurs spontaneously to give areas of green and white tissues, facilitating recognition of such clonal differences. Likewise, in various fungi, segregations of particles t h a t occur in the tips of the hyphae can be recognized because they produce

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phenotypically different mycelia. Scgregatioii rates and selectivity of clones may be different in different parts of a life cycle. More definite experimental results may be expected if organelles can bc transferred into cells at will by rnicromanipulation, as referred to in the scction on mitochondria. Complementation studies, it is hoped, will soon demonstrate the relation of plastid mRNA to plastid DNA, and of mitochondrial mRNA to mitochondrial DNA. This powerful tool may eventually be developed to analyze for specific genes perhaps by coupling i t with electron microscope observations. A more direct approach to the study of cytoplasmic organelles may be possible if they can be cultured in vitro. In this way one may learn what their genes are doing and what interactions occur with nuclear products. This approach appears to be less unreasonable now than formerly because genes and protein-synthesizing mechanisms have been found to be localized in these organelles.

B. Constitution of a Cytoplasmic Nucleoid The nucleoids, each consisting of several hundred genes, may be expected not to differ greatly in structure from viral DNA. Each nucleoid gm) . The chromoprobably contains a single chromosome (1-100 x some may be circular and it is known to be double-stranded. Doublestrandedness was deduced from base ratios, hyperchromicity, and increased density after melting, and by acridine orange staining. One may therefore expect to find that all genes of an organelle belong to one linkage group only. Where there is only one nucleoid per organelle, and each organelle in the case of plastids and mitochondria has a double membrane, exchange of genes between the nucleoids is not to be expected unless a particular infection mechanism has been developed. However, if the fusion of mitochondria as seen in the light microscope is real, then transfer of mitochondrial nucleoids from one mitochondrion to another should occur and mitochondrial gene exchange should be possible. Because the centrioles are naked, there is more chance that its nucleoids may incorporate or exchange genes, even genes of some viral DNA. The relatively high values reported for DNA per plastid (Table I) of certain plants may indicate either that there is more than one nucleoid per plastid or that there is a single polytenic nucleoid that has arisen by repeated duplication.

C. Redundancy, 'Organelle Survival, and Organelle Mutation Besides the effect of gene redundancy in prohibiting the application of Mendelian analysis to mutations in mitochondria and plastids, two other consequences of redundancy may be noted.

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A mutation in one organelle may cause its elimination if it results in a decrease in rate of multiplication, as compared to a nonmutant. There is, however, no danger of losing all the organelles by gene mutation because of the large number of organelles per cell. This means that the stability of the cytoplasm is assured with regard to mitochondria and plastids. The other consequence of redundancy is that clones of organelles can develop in the cytoplasm of a cell, each somewhat different in its genetic makeup. For example, because the organelles are numerous, certain mutations may accumulate in them if they do not decrease the rate of multiplication of the organelle. This should lead to the presence in the same cytoplasm of a mixture of clones. Such a mixture may provide the cell with greater physiological versatility. A practical implication of these considerations is that, by crossbreeding, i t may be possible to select clones of chloroplasts that will function best under different conditions of light and temperature. I n the case of mitochondria, selection for clones t o function under different conditions of temperature and 0, supply may be considered. Such crosses and back-crosses should be made with due regard to the fact that the desired organelles will be inherited maternally. It will be of interest to examine whether some agricultural plants such as corn or wheat, which are derived from interspecies crosses, already contain selected clones of plastids and selected clones of mitochondria. I n seeking among plants for clones of plastids or mitochondria, one may consider that a species that grows under extremes of temperature and light may be favored by clones that can multiply more rapidly a t low or high temperatures, or by clones that multiply more rapidly under conditions of low or high light intensity. May it be that the same plant species, growing a t the base of a mountain, contains a population of plastids and mitochondria in its cells that differ from the population of organelles in the same plant species growing near the top of a mountain? I n animals, may a species be artificially changed in its habitat tolerance by incorporating a clone of mitochondria from a closely related species of a different habitat? Assuming that the respiratory rate of different mitochondria are controlled by temperature, may it be possible, for example, to transfer mitochondria from egg cells of a fish that lives in arctic waters to a fertilized egg cell of a closely related species that lives in tropical waters and get the developing fish to survive cold waters?

D. Interactions of Nuclear and Cytoplasmic Genes The probability of survival of cytoplasmic organelles, when transferred from one species to another, even closely related, may be very

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low. Even the organelles of a male cell often do not survive in a female cell a t fertilization, but occasionally they do. There must be factors of compatibility that are controlled not only by nuclear genes but by cytoplasmic genes as well. Evolution has resulted from mutations in both nuclear and cytoplasmic genes. Survival requires compatibility, which must mean that coadaptation among all of these genes must take place. For example, one plastid mutation has led to pollen sterility, which has been overcome by mutation of a nuclear “suppressor” gene. Coadaptation of nuclear and cytoplasmic genes to a particular environment has led to divergent races of Oenothera, Culex, etc., and has ultimately led to speciation ( 2 ) . Clausen (111) found that the most widely distributed trees are the most tolerant to low temperatures and high altitudes. They belong to species complexes that are known for their ability to hybridize. Because of studies on cytoplasmic inheritance, one must now consider that the ability of an organism to cope with difficult environments may depend not only on nuclear gene pools but also on pools of clones of cytoplasmic organelles. In the development and differentiation of an organism, the numbers of organelles per cell and their differentiations to mature organelles are the result of the interaction of nuclear and cytoplasmic genes of a particular tissue. For example, liver mitochondria are small in size and have relatively few crystae in contrast to the mitochondria of kidney tubules, which are elongated and have numerous crystae. It is known that nuclear genes affect the differentiation of organelles. For example, the growth of a proplastid to chloroplast may be controlled by a number of nuclear genes. A nuclear gene has also been found that induces an acceleration of mutations in plastids (112). But cytoplasmic genes also have an effect on nuclear genes. For example, genetic studies in Oenothera have shown that cytoplasmic genes may affect the stability of the nuclear chromosomal complement itself. Certain cancers are thought to be caused by the attachment of viral DNA as episome to nuclear DNA. The presence of DNA in cytoplasmic organelles leads to the question whether other types of cancer might be caused by viral DNA carried on the DNA of cytoplasmic organelles and inherited.

VI. Summary Evidence is presented for the presence of DNA in three kinds of cytoplasmic organelles: the chloroplasts, mitochondria, and centrioles. The DNA in each organelle is double stranded. The lowest values for DNA in each of these organelles is about 1 X gm, or enough to code for a maximum of a hundred different kinds of protein molecules. The DNA

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strand of a plastid or mitochondrion gives rise to a new DNA strand by semi-conservative replication as it does in bacterial DNA and in the nuclear DNA of plants and animals. I n the case of E u g l e m plastids, it is inferred that its DNA does not arise, even indirectly, from the nucleus. Labeling studies wit11 plastids and mitochondria sulqiort the hypothesis that DNA codes for mRNA, which is translated to protein. The proteins are required for the growth and differentiation of the proplastid to chloroplast, of the promitochondrion to mitochondrion, and of the procentriole to centriole. Products from the nucleus affect the multiplication and differentiation of the organelles. Concomitanly, products from the organelles affect nuclear DNA. The multiplicity of plastids and mitochondria in a cell suggests the hypothesis that there may be a number of clones of these bodies in a cell. Thus, the inheritance of an organism may be constituted not only of a nuclear gene pool but, in addition, of gene pools of clones of cytoplasmic organelles. The significance of cytoplasmic genes is discussed in terms of their phenotypic effects, and their relation to hybrid vigor, ecology, and cancer.

ACKNOWLEDGMENTS This review was supported in part by Grant GM-04922 from the National Institutes of Health, and Grant GB-2617 from the National Science Foundation. We also desire to express our gratitude to Dr. Maria Rudeinskn for her helpful criticisms of a portion of the manuscript.

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Behavior, Neural Function, and RNA H. H Y D ~ N Institute of Neurobiology, Medical Fucult y, University of Goteborg, Goteborg, Sweden

I. Introduction . . . . . . . . . . . . . . . . 11. Problems Discussed . . . . . . . . . . . . . . 111. Methods of Analysis. . . . . . . . . . . . . . IV. Biosynthesis of Rapidly Labeled RNA in Brain Cells . . . . V. Acidic Proteins Specific for the Brain . . . . . . . . . VI. Base Ratios of RNA during Physiological Stimulation . . . . VII. Base Ratios of RNA during Chemical Induction of RNA Synthesis VIII. The Emergence of RNA Rich in Adenine and Uracil during Learning Experiments . . . . . . . . . . . . . . . IX. The Possible Transfer of RNA between Glia and the Associated Neuron . . . . . . . . . . . . . . . . . X. The Synthesis of an Asymmetric, Adenine-Rich RNA in Parkinson’s Disease . . . . . . . . . . . . . . . . XI. A Working Hypothesis . . . . . . . . . . . . . References. . . . . . . . . . . . . . . . .

187 190 191 194 197 199 199 202 208

210 213 21 7

1. Introduction The discussion that follows is concerned with the question whether biochemical reactions involving macromolecules specific for learning occur in brain cells but do not occur during increased demands on neural function. This problem involves as a main point the level a t which such reactions may occur and the regulatory mechanism that might be utilized. Small molecules of short life can presumably be excluded if a molecular substrate that can serve long-term memory is considered for storage of information. Interest is instead focused on macromolecules, especially on RNA and proteins. When empirical data are analyzed and an animal experimental program carried out, learning means the capacity of a system to react in a new or modified way as a result of experience. Memory, in a more strict sense, can be defined as the capacity to store information that can be 187

188

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recalled again with high distinction to steer thc function correlated with the new information. The execution of an experimental program in this field is fraught with potential fallacies. More than in other biological fields, would-be discoveries may turn out to be delusions. It seems appropriate to quote Polanyi ( 1 ) in discussing the content of empirical statements. Polanyi has pointed out that such content relies on clues that are largely unspecifiable, integrates them by principles that are undefinable, and speaks of a reality that is inexhaustible. Nevertheless, the problems of learning, life-long memory, and problem seeking of man have been phrased and dealt with throughout the ages. In 375 BC, Plato published a dialogue in which Socrates discussed learning and memory (2). He said that, when we solve problems, we do it by remembering past incarnations. What we call learning is in fact remembering. To learn the truth, on the other hand, is impossible. If we know the solution to a problem, there is no problem. If we do not know the solution, we do not know what to look for and cannot expect to find anything. Today we would phrase such a suggestion by speaking of activation of a genetically stored memory. We can contrast these ideas of Socrates by the belief of John Locke that knowledge is obtained solely experientially. Only by experience, said Locke in 1686, can man get his immense and broadly varied knowledge (3). Both these ideas are easily recognized main points in modern psychology from a phenomenological point of view. Ethology has now broadened the scope of neurobiologists. The study of insect behavior for example, has given a detailed picture of the genetically programmed activity, instinct” behavior, triggered by key factors in the environment. It is easy to find examples of such behavior, governed by a wellprogrammed computer, as it were. The Capricorn beetle digs channels in the center of oak trees for 3 years during the larval period ( 4 ) . The adult beetle, however, cannot digest the hard oak tree and the question is how it escapes from the tree after metamorphosis. The answer is that the larva has everything planned in detail. It has dug a wide channel twice its own diameter on the cortical surface, which becomes paper-thin. The larva connects this wide channel with a chamber whose size is just sufficiently large to house the beetle, which still does not exist. Then the larva lies down in the chamber to change into a pupa, always in such a way that its head is towards the exit to the channel. Otherwise the adult beetle could not escape, since it cannot turn in the chamber. This type of behavior is not known to be modified by experience. Genetic studies have also revealed

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that certain recessive genes in homozygous form may govern various steps of complex motor behavior in insects. One might ask if it ever will become possible to discern genetically programmed activity from other types of activity in man. One such activity seems to be recognized in the human male as well as in the male of other primates ( 5 ) . It could be called the imperative of the territory and the status symbol. In contradistinction to this is the insight learning of man, the problem solving by the pursuit of deepening coherence, to quote Polanyi (11, and the new combination of happy ideas. But i t should not be forgotten that rats also utilize abstraction in maze solving (6). If we turn to biological concepts and technical tools of today, it may be asked whether there has a t last arisen the possibility of investigating experimentally a probable substrate for neural memory and the mechanism for learning and recall of data. This would require not only the technical means but new concepts that could mediate access to various ways to gain deepening coherence that leads to a discovery. Molecular aspects of biological processes have perhaps most to offer for the formation of new concepts of primary cellular events, especially genetic mechanisms, regulatory cell processes, and protein production. As has always been the case in the history of science, new ideas in important fields create a crisislike uncertainty among scientists until these concepts have been assimilated and can serve productive creation by those who have the capacity to reorganize older, and usually firmly rooted, concepts (7). Our concepts about genes have certainly changed greatly during the last few years. Until recently they were looked upon as more stable functional units, active a t meiosis and mitosis. Once activated in cells in a differentiated organ, they went on ticking like clocks for the rest of the life cycle. Now it is considered that genes can be switched on and off by external factors. Differentiation is reflected in the population of the so-called messenger RNA, which shows rapid turnover, heterogeneity in size, and “template” activity. During morphogenesis, RNA is transcribed from specific genetic loci. Only part of the genome is active at a given time in a cell. As much as 80% of the DNA may be inactive in the cells of higher organisms. We also have learned that some external factors have the capacity of penetrating to the genes and releasing activity, whereas others do not. One of the best cases deals with the effect of testosterone in increasing the RNA polymerase activity and level of RNA in the accessory sex glands of the male (8, 8 a ) . RNA has also been reported t o mimic the effect of a hormone if applied locally on cells. This has led to the view that some hormones, and even nutritional

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influences, may constitute factors able to depress regions of the genome. This leads to a production of more specific messenger RNA which in turn gives more specific protein molecules as end products. Such concepts have begun strongly to influence our way of reasoning by analogy, even if little is known about the extent to which protein synthesis in specialized mammalian cells is dependent on rapidly synthesized RNA or on messenger RNA with slow turnover and greater stability. Nevertheless, concepts from the field of molecular biology have begun to influence the neurosciences. Such concepts can be expected to stimulate problem seeking in neurochemistry before their magic diminishes and the position becomes static, when the next crisis brings about a new cycle of progressive events. II. Problems Discussed Neurons contain large amounts of RNA. I n learning experiments, a production of RNA and changes in base ratios occur in both neurons and glia (9a-c) . Physiological and chemical stimulation, on the other hand, increase the RNA in neurons and decrease the RNA in glia (lU, 11). Since such phenomena inevitably bring up the question of the turnover of the various types of RNA in the brain and in its cells, it is appropriate first to discuss rapidly labeled RNA in brain cells. I n most cases, RNA synthesis is followed by protein production. Each somatic cell can be expected to produce more than a thousand different proteins. Most of these are common to the protein population in all cells of an organism. Therefore, only a very limited number of protein end products may characterize a highly differentiated tissue, such as the central nervous system. An acidic protein has been described, apparently specific for nervous tissue (la), that contains a fraction with a very high turnover and a characteristic localization; some of its properties are worthy of discussion. There are also indications that soluble proteins are synthesized during learning experiments. The main features for discussion, however, are the following:

Do the biochemical responses of neurons and glia in learning differ from those following physiological or chemical stimulation? Does the RNA composition reflect nuclear and genetic activity when an animal is faced with a situation it has not previously encountered? ( b ) What mechanism underlies the inverse RNA changes between glia and neurons? In this connection we discuss the question of a transfer of RNA between glia and neuron in contradistinction to the situation in learning where the RNA changes more in the same direction in both neurons and glia.

(a)

BEHAVIOR, NEURAL FUNCTION, AND RNA

191

a genetic aspect will be discussed on the basis of information derived from RNA analyses, demonstrating that factors in the environment may lead to a stimulation of genetic material and a release of genome activities in brain cells. A biochemical glial error in a neurological disease may serve to demonstrate a case in which such a genetic activity is undesirable and functionally harmful.

(c) Furthermore,

Finally, a working hypothesis for memory will be suggested.

111. Methods of Analysis Sucrose density gradient studies were performed on nuclear and cytoplasmic RNA from the lateral vestibular nucleus in rabbits. The animals received 150 pc of H3-orotic acid through a cannula inserted into the fourth ventricle in pulses whose duration varied from 15 to 180 minutes ( I S ) . Each sample of tissue from the lateral vestibular nuclei had a wet weight of 0.4 gm. It was homogenized and the RNA was differentiated by centrifugation into nuclear and cytoplasmic fractions. The specific activity was determined as cpm/pg RNA. Some of the information referred to in the following pages has been obtained with microchemical techniques on isolated neurons or samples of glia (l4a-e, 1 5 ) . The reason is the complicated structure of the brain and the intimate structural relationship between glia and neurons (Fig. 1). In certain circumstances the biochemical changes in neurons may be the inverse of those in the surrounding glia. Therefore, even if only 1 pg of material is taken from the brain, the sample is probably 50% nerve cell material and 50% glia. Inverse biochemical changes in these two types of cells will most certainly cancel each other in a bulk analysis. Different biochemical responses may also escape detection if microgram amounts of brain material are subjected to analysis. Therefore, nerve cells including the first part of the dendrites, and neuronal glia, have been isolated from precipitated or fresh brain material. A de Fonbrune micromanipulator was used after free-hand dissection of the fresh material. In some cases only nerve cell nuclei were used for RNA analysis. For each analysis twenty-five to thirty nuclei were isolated by microdissection. The technique is briefly as follows. The isolated nerve cells, on a glass slide, were treated with cold phenolsaturated water for 15 minutes followed by cold absolute ethanol for 10 minutes and then covered by liquid paraffin. The effect of this treatment is to cause the nuclei to contract slightly but this is hardly noticeable a t a magnification of 600 times. The nucleus from each nerve cell could then easily be removed with the aid of the micromanipulator. Twentyfive nuclei were used for each RNA analysis.

192

€1. H Y D ~ N

Careful checks were made to ensure that the phenol treatment precipitates all RNA in nerve cells. For the quantitative and qualitative analysis of RNA, the micro methods deve!oped a t this laboratory were employed using the detailed technical microelectrophoretic procedures

FIG.1. Schematic picture of two nerve cells surrounded by glial processes. To the right is seen a capillary that is also ensheathed by glial membranes. The present paper includes a discussion of the biochemical processes occurring in nerve cells and in the glia immediately surrounding the nerve cell body, the so-called ncuronnl glia. The two types of cells composing the central nervous system consist of nerve cells with their processes, dendrites, and axon, and different types of glial cells. The thin, delicately folded and membranelike processes of the glia intertwine and ensheath the neurons except a t those places where the synaptic knobs take up part of the neuronal surface.

recently published (14e). For each analysis, 500 to 700 ppg of RNA were used. The random error in the determination of the RNA in single nerve cells was found to be 4%. The average coefficient of variation of the analytical results in the microelectrophoresis was 5% for nerve cell RNA and 7% for yeast RNA. I n one case in which i t was

BEHAVIOR, NEURAL FUNCTION, AND RNA

193

possible to conipare tlic result of the niici~oe1ectroi)lioreticseparation of hydrolyzed RNA from biological material with that obtained by conventional macrochemical electrophoretic separations, the analysis of nucleolar and ribosomal RNA of mature starfish oocytcs gave the same results with both methods ( 1 6 ) . In iiiotlcl cxperiiiicnts on purified saiiiples of RNA, the correspondence between macro- and microelectrophoresis is clear (14e). The advantage of microelectrophoresis over macroelectrophoresis is the possibility of analyzing samples a t the cellular level. This has proved to be a sine qua non for nervous tissue since its two cellular components, neurons and glia, differ in content and composition of RNA. The neuronal and glial RNA were determined on the same dry weight basis by quantitative X-ray microspectrography ( 1 7 ) , using a scanning and computing densitometer (16a,b). RNA metabolism a t the cellular level has been determined by a new micromethod (18, 1 9 ) . Total, labeled RNA extracted froin nerve cells or glia, or the individual bases obtained after electrophoresis on a microscopic cellulose strip, are combusted a t 650°C in a single step in glass capillaries containing KClO, and Zn particles so that the organically bound tritium is transformed into hydrogen gas. The activity is determined in a modified Geiger-Muller tube ( 1 8 ) . This method combines the high efficiency in tritium detection (>70%) and a low background ( < 5 cpm) and permits an accurate assay of low levels of radioactivity. Soluble proteins extracted from fresh, isolated neurons or glia were analyzed by disc electrophoresis on polyacrylamide gel in capillaries of diameter 200-300 p (20), the appropriate manipulations being carried out free-hand. The samples contained about gm of protein. Homogenization of the cells was carried out in a capillary with a twisted loop of steel wire of 28 p diameter driven a t 12,000 rpm. The current required for separation is 1.5 pa for 3-4 hours. The protein pattern was obtained by amido-black staining and scanning with a microdensitometer. The biosynthesis of proteins was studied on material from animals previously treated with H”1abeled amino acids. The individual bands in the gel were cut out under a microscope, combusted in capillaries, and counted as described above. For the localization of the acidic protein in the various types of brain cells, single-diffusion agar precipitation was performed in glass capillaries 300 p in diameter, using antiserum against the acidic protein. T o localize this protein a t the intracellular level, the multiple-layer method of Coons (21) was applied. This technique involves cryostat sections of the tissue and the antigens were identified by fluorescence methods.

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IV. Biosynthesis of Rapidly labeled RNA in Brain Cells The distribution of the various RNA species in brain and the changing density gradient profiles of RNA during the first 30 minutes of synthesis differ in detail from those found in other somatic cells of inammals. After a 15-minute exposure to H3-orotic acid administered by permanent cannulae in the fourth ventricle, the main finding in the nuclear fraction is the occurrence of heterogeneous RNA with sedimentation

b/3 *vtQ

200 250

I'

,

- Marker RNA I

5

I

'I0 Tube

I

20

I

25

I

"

50

- 10

!5

150

100

/

~

5

number

10 Tube

15

20

25

number

p I 0.300

0

N #

2

0.200 0.100

100

50 5

10

15

Tube number

20

25

50 5

10 Tube

15

20

25

number

FIG. 2. Sucrose density gradients of nuclear RNA species after injection of H3-orotic acid into the fourth ventricle; (a) after 15 minutes, (b) after 30 minutes, radioactivity; -0-, UV (c) after 60 minutes, and (d) after 180 minutes. -0-, absorbance. [From Egyhazi and HydCn (36).l

coefficients around 8 to 12 S (36) (Fig. 2 ) . This implies that the RNA synthesis in brain cells started with an initial formation of small RNA molecules, though of a messenger type. The final judgment of the messenger character must, however, await evidence of template activity. After 30- and 60-minute pulses, the profile of the synthesized RNA is changed, showing a shift of the radioactivity to regions between 16 and 30 S. New peaks also become evident. The specific activity increases during the period from 27 to 335 cpm/pg of nuclear RNA. It is interesting to note that, in the cytoplasmic RNA, heterogeneous RNA with a profile similar to that of the nuclear RNA after a 15-

195

BEHAVIOR, NEURAL FUNCTION, AND RNA

minute pulse appears after 30 minutes (Fig. 3 ) . The specific activity increases from 8 to 67 cpm/,ug of cytoplasmic RNA. This phase shift and correlation of types of density gradient profiles suggests a transport of RNA of a messenger type from the nucleus to the cytoplasm. If breakdown RNA products occurred in the cytoplasm, one would expect 4-8 S RNA. These observations agree with those of Jacob et al. (37) who found a high amount of radioactivity in the 10 to 45 S of the nuclear RNA from rat brain 30 minutes after intracisternal injections of nucleosides.

125

Marker

2

RNA

0.150

N

V

50

d

25

5 Tube

number

10 15 20 Tube number

25

n

N

150 100

5

10 15 20 Tube number

25

FIQ.3. Sucrose density gradients of cytoplmmic RNA species after injection of H3-orotic acid into the fourth ventricle: (a) after 15 minutes, (b) after 30 minutes, and (c) after 30 minutes and removal of soluble RNA. [From Egyhazi and HydCn (36).I

I n the following discussion, it is shown that an increase of RNA content of 30% can occur in neurons within 1 hour together with concomitant changes in base ratios. It would be simple to assume th a t this is due to an excessive biosynthesis of nuclear RNA flooding the cytoplasm, because there is no evidence that there exists a mechanism for RNA synthesis in the cytoplasm. This possibility cannot, however, be excluded. Microchemical fractionation of nerve cell RNA has shown the presU) ence of a substantial fraction of RNA with a low (G C ) /( A ratio (0.85), and hence a composition complementary to that of DNA

+

+

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( 3 8 ) .This may be a cytoplasmic messenger RNA. To look for structural correlations, isolated, fresh nerve cells were cut open by free-hand microsurgery and the cellular content was transferred to electron microscope grids for photography (39). A striking finding was the occurrence of

FIG. 4. Shadowed preparation from Deiters’ nerve cells showing: (a) 100 A ribosomes in a linear formation attached to a thin thread structure, ~ 1 3 0 , 0 0 0 ;(b) a giant polysomal structure, ~120,000.[From Ekholm and Hyddn (39).l

large polysomal structures, some of them containing more than a hundred ribosomes (Fig, 4). The RNA metabolism of nerve cells and glia has been compared by the use of micro methods after electrophoretic separation of the bases (19). The conversion relations between the RNA purine precursor pools

197

BEHAVIOR, NEURAL FUNCTION, AND RNA

and the pyrimidine precursor pools were the same in nerve cells and glia, although the labeled RNA in the two types of cells differed in composition. The synthesis of RNA was twice as rapid in the glia as in nerve cells. I n the following discussion, several examples are presented of RNA production effected by physiological stimulation or by chemical agencies. It should not be forgotten, however, that a cyclic change occurs during the lifetime of the RNA content of neurons. Table I gives some figures from a study of motor neurons in the spinal cord of man. The samples were taken from neurologically sound individuals from 3 to more than 80 years of age who died in traffic accidents (40). It can be seen that the RNA content increases up to the age of 40 when it levels out for the TABLE I TOTALRNA CONTENT I N MOTORNERVE CELLSFROM MAN (SPINAL CORD,Cat- NUCLEUSVENTRALIS LATERALIS)

0-20 21-40 41-60 61-80 Over 80 a

402 553 640 504 420

28 f 38 k 5.5 ?c 31 f 30

Fifth cervical segment.

next two decades. After the age of 60 the RNA content falls quite rapidly. This change in RNA during the life cycle is superimposed upon the short-lasting, reversible fluctuations in RNA content that may be a result of increased functional demands.

V. Acidic Proteins Specific for the Bruin From kinetic analysis of whole brain protein, it has been concluded that the life spans of the various brain proteins vary from a few minutes to many months [for a review, see Lajtha (4111. Moore e t al. (1.2) have recently isolated and characterized a n acidic protein, called S 100 protein. This was found to be specific for nervous tissue and to make up 0.5% soluble proteins of brain tissue. Antiserum to this protein showed a cross reaction with proteins from a number of vertebrate species, a finding that has given a new impetus to the study of brain proteins. We have investigated this protein by macro- and microelectrophoretic and immunological methods (42, 43) with the aim of correlating protein reactions with the extensive RNA synthetic processes that occur

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in brain cclls. The protein band containing the S 100 protein can be separated into a t least three components. Two of the three acidic proteins attain very high specific radioactivities within 30 minutes after injection of H”1eucinc. The specific activities for these two proteinsone of which consisted entircly of the S 100 protein-decline rapidly between 6 and 12 hours (Fig. 5 ) . This suggests that the newly synthesized proteins turn over rapidly. A third acidic protein belonging to the original S 100 protein band had a low specific activity. I

L-Leucine-4,5-T

30’2hr 6hr

12hr l8hr 24hr I w k Time

3wk

FIG.5. Specific radioactivity of bands Oa, Ob, and Oc separated on 11.2% polyacrylamide gels as a function of time between isotope injection and sacrifice. Radioactivity was determined by liquid scintillation counting, after combustion of slices of the polyacrylamide gels. Isotope: ~-leucine-4,5-T. [From McEwen and Hydkn (421.3

Rubin and Stenzel (44) have reported that 15% of the radioactive protein formed in a cell-free system from rabbit brain can be precipitated by a n t i s 100 serum. Their observations suggest that a considerable part of the protein synthetic capacity of the brain is devoted to the synthesis of this protein. Our own observations imply that this large synthetic capacity is necessary because of the high turnover of the acidic brain proteins. The detailed localization of the S 100 protein was determined by single-diffusion agar precipitation using antiserum and antigen extracted from isolated neurons and glia, and by fluorescent antibody techniques (43). The S 100 protein was localized mainly in the cell bodies and in the membranous system of oligodendrocytes, but was absent from their

BEHAVIOR, NEURAL FUNCTION, AND RNA

199

nuclei. It was present in the nuclei of the big neurons, but not in the cytoplasm. Pertinent questions relate to the site of synthesis of this protein, whether i t be in the glia or in the neuron or in both or whether i t is transferred after synthesis from one cell type to the other. There is also a cell regulatory question to consider. Histones seem to regulate the activity of genes. The acidic nature of the S 100 protein is such that i t might well react with the histones in the nuclei of the neurons and control gene activity. I n other words, the acidic proteins in the brain might control the controllers.

VI. Base Ratios of RNA during Physiological Stimulation To stimulate the neural function in experimental animals, rotatory, horizontal, and vertical vestibular stimulation through 120°C, with direction changing every 2 seconds, was used. It was found that the socalled Deiters’ nerve cells in the vestibular nucleus of the brain stem and nerve cells from the reticular formation increased their RNA content by 5 to 25% [Deiters’ nerve cells in the rat, from 680 t o 740 ppg (horizontal rotation), from 680 to 850 ppg (vertical rotation) ; nerve cells of nucleus gigantocellularis in the rat, from 540 to 590 p p g ] (9b, 11). But no significant change of the base ratios in the RNA was observed in any of these experiments. The composition remained typical of the cytoplasmic high-polymer ribosomal RNA, which is, for Deiters’ nerve cells in the rat: 20.5 A, 33.7 G, 27.4 C, 18.4 U. The nuclei of nerve cells are usually small in comparison with the amount of cytoplasmic material. I n Deiters’ nerve cells from rats, for example, they contain 30 p p g of RNA compared to the 650 ppg of cytoplasmic RNA. This nuclear RNA has the following base ratios: 21.4 A, 26.2 G, 31.9 C, 20.5 U (9a). The explanation is presumably the predominance of nuclear ribosomes and the nucleolar RNA, which is of ribosomal RNA type. This means that the newly formed RNA of the nerve cell during physiological stimulation has the same base composition as the original RNA of the ribosomal type.

VII. Base Ratios of RNA during Chemical Induction of RNA Synthesis Tricyano-amino propene (triap) adniinistercd intravenously to animals at 20 mg per kg causes an increasc within 1 hour in the amount of RNA per nerve cell, in the respiratory enzyme activities, and in the total dry weight including proteins. The base ratios of the total nerve cell RNA change significantly (10). The quantitative increase in RNA was of the order of 570 ppg in cclls containing an original amount of

200

13.

HYDBN

1500pp per nerve cell. Table I1 shows the RNA base ratios of the control nerve cells. The chemical agent changes the proportions of guanine and cytosine significantly. A more useful insight into the changes in RNA composition occurring after chemical stimulation is obtained if the base-ratio alterations are referred to the change of the amount of RNA (in the following called the “ARNA fraction” or “ARNA”), and not to the final amount of RNA. This means, for instance, in the case of an increase of the RNA content with a simultaneous change of the base ratios (as in the case TABLE I1 MICROELECTROPHORETIC ANALYSESOF THE COMPOSITION OF DEITERS’NERVECELLSFROM RABBITS( 1 O P Basc

Controls

Adenine Guanine Cytosinc Uracil

+

(G C)/@ A/U

19.7 33.5 28.8 18.0

+ U)

No. of animals: No. of nerve cells:

f 0.37

k 0.39 f 0.36 f 0.18

Trirtp 20.5 34.6 26.7 18.2

f 0.31

k 0.28 f 0.24 f 0.20

1.65 f 0.023 1 . 0 9 f 0.021

1 . 5 8 f 0.018 1 . 1 3 f 0.021

5 490

11 1201)

THE

RNA

IN

ARNA fraction 22.5 37.7 21.0 18.8

f 1.61

k 1.66 f 2.03 f 0.91

1.42 f 0.090 1 . 2 0 f 0.103

Rabbits were treated with 20 mg/kg of tricyano-amino propene and killed 1 hour later. Purine and pyrimidine bases in molar proportions in percentages of the sum. Total amount of RNA increased significantly from 1550 ( f 7 8 ) to 2120 ( f 1 0 6 ) rrdcell.

above after injection of tricyano-amino propene) that the chemical change of RNA is described as a synthesis of a newly formed amount of RNA ( A R N A ) with a base ratio differing from that of the original RNA. I n the same way, a decrease in the amount of RNA with a simultaneous change in base ratio can be considered as a loss of an RNA fraction ARNA) in such a way that the composition of the ARNA removed differs from that of the original RNA. Table I1 shows that an increase in the proportion of guanine and a decrease in the proportion of cytosine is characteristic of the final RNA. This is more pronounced if the changes are calculated on the basis of the newly formed ARNA fraction, as described above. The ratio (G C ) / (A U) in both cases (final RNA and ARNA) differs little from 1.7 in the control nerve cell, being 1.6 and 1.4, respectively. This means that the newly synthesized RNA is of the ribosomal type.

+

+

20 1

BEHAVIOR, NEURAL FUNCTION, AND R N A

As has already been pointed out, tlic amount of RNA of the nucleus in these big nerve cells is only a fraction of the RNA total, 55 ppg of 1550 p p g , i.e., 3.5%. Considering, however, the importance of the nucleus for protein synthesis in the cell, the nuclear RNA composition was analyzed in control nerve cells and in cells after induction of RNA synthesis by triap. This was done by microdissection whereby the nucleus was removed from each cell. The RNA collected from twenty-five nuclei was pooled for each electrophoretic analysis. Table 111 shows the observations made, which indicate an average loss of 16% of RNA from the nucleus. If this loss (ARNA, column 4, TABLE 111 MICROELECTROPHORETIC ANALYSES OF THE COMPOSITION OF THE NUCLEAR RNA IN DEITERS’NERVE CELLSFROM RABBITS( 9 ~ ) ” Base

Controls

Adenine Guanine Cytosine Uracil

+

((2 C)/(A A/U

No. of animals: No. of nuclei:

ARNA fraction

f 3.75 f 7.10 f 3.56 f 7.36

21.5 f 0 . 5 4 3 0 . 0 f 0.45 30.4 f 0.49 18.1 k 0 . 4 4

20.3 9.4 32.8 37.5

1.34 f 0.022 1 . 0 0 f 0.028

1.52 f 0.040 1.19 f 0.041

0 . 7 3 f 0.172 0 . 5 4 f 0.148

22 275

26 380

21.3 26.6 30.8 21.3

+ U)

Triap

f 0.42 f 0.27 f 0.38 f 0.43

a Rabbits were treated with 20 mg/kg of tricyano-amino propene and killed 1 hour later. Purine and pyrimidine bases in molar proportions in percentages of the sum. Total amount of RNA decreased significantly from 56 ( 5 2 . 7 ) to 47 ( f 1 . 9 ) ppg/ nucleus.

Table 111) is considered to represent an RNA constituent of the nucleus, it has an unusually high amount of cytosine and uracil. The ratio (G C ) / ( A U ) is 0.73 compared to 1.34 for the control nuclear RNA and 1.52 for the total nuclear RNA of the triap-treated nerve cells. Thus, the composition of the nuclear RNA after treatment with triap is of a more ribosomal character, since the guanine and cytosine values are increased. This fact may be explained as a consequence of the activity of the nucleus during the period of high RNA synthesis in the cytoplasm. It can accordingly be asked whether the base ratios of the lost RNA (20.3 A, 9.4 G, 32.8 C, 37.5 U ) represent a “messenger” RNA. Furthermore, since the nucleolus is the main site of the ribosomal RNA synthesis in the nucleus and has an RNA of ribosomal type, the finding of a more ribosomal character of the nuclei is not surprising.

+

+

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With phenylcyclopropylainine, a powerful inhibitor of monoamineoxidase, in doses of 0.3 mg/kg body weight, an increase of the neuronal RNA of 30% per neuron and a decrease of the glial RNA by almost 50% (9d) were obtained in 1 hour. Significant base ratio changes involving guanine and cytosine were found. Moreover, the cytochrome oxidase activity increased in the neurons by 250% and decreased in the glia by 30%. When actinomycin D was given in doses of 0.3 mg intravenously to rabbits and 3 to 7 hours before injection of phenylcyclopropylamine, not only did the actinomycin inhibit the RNA changes, but it also produced a loss of RNA by 100 ppg/cell. The base composition of this lost RNA fraction was 25.7 A, 43.3 G, 23.0 C, 8.0 U. The conclusion from these experiments was that the RNA production induced by chemical stimulation is immediately DNA-dependent. Moreover, this production actually involves a loss of RNA, the base composition of which is highly asymmetric through the influence of actinomycin. It can thus be said that the cytoplasmic RNA newly synthesized after both physiological stimulation and induction by a chemical agent has the base ratios of a ribosomal RNA.

VIII. The Emergence of RNA Rich in Adenine and Uracil during Learning Experiments Two types of learning experiments were performed on rats. Neurons in the cortex and in the brain stem involved in the learned behavior were analyzed. I n contrast to the result after physiological and chemical stimulation, the RNA formed in the neurons during learning had, in one case, high adenine and uracil values and was, in the other case, asymmetric with high adenine values. The RNA fraction rich in adenine and uracil has a low (G C)/ (A U) ratio of 0.70 to 0.90 and cannot be extracted by cold perchloric acid, acetic acid, or water. The same holds for the asymmetric, adeninerich RNA. It is, therefore, likely that this fraction consists of polymer RNA rather than of small molecules. I n mammalian cells, nuclear and cytoplasmic RNA fractions have been described that stimulate the incorporation of amino acids, and are rapidly labeled. They furthermore show the low (G C)/(A U) ratio of 0.9 to 1.0 and an asymmetric base ratio composition with adenine values higher than the uracil values (22, 23). The question of protein synthesis in mammalian cells and its dependence on messenger RNA with a slow turnover or on rapidly synthesized RNA in the nucleus or cytoplasm is, however, far from settled. Work in progress may elucidate possible template activity and other characteristics of the different RNA fractions of neurons and glia. I n the first experiment, right-handed rats were induced to use the left paw

+

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203

in retrieving food from a narrow glass tube ( 9 c ) . Training periods of 2 X 25 minutes per day were given. The neurons of both sides of the cortex were analyzed, especially from those areas whose destruction prohibited transfer of “handedness.” These control centers are situated bilaterally in the sensory motor cortex and comprise around 1 mm3 of the cortical material. Layers 5-6 are the most important. These neurons have a nucleus large in comparison with the cytoplasm and, consequently, the analytical results mainly reflect nuclear RNA. Eighty-eight rats were used for the analysis of 14,000 cortical neurons. The advantage with this learning experiment is that the control cells are derived from the same brain. Therefore, a paired t analysis can be performed on the results from the neurons of both sides. Other control experiments have also been performed. A significant increase of the amount of RNA per cell occurred already in the learning side of the cortex. I n an extension of the work already published ( S c ) , the amount of RNA was found to have increased from 220 to 310 ppg RNA per 10 nerve cells. When the base ratios of the neuronal RNA of the control side were compared with those from the RNA on the learning side, it was found that the ratio (G C)/(A U) had decreased significantly from 1.72 to 1.51. In this paper, a detailed analysis was performed to investigate whether the extent and type of RNA change can be correlated with the learning process in individual cases. For the original RNA base ratio data, the reader is referred to HydBn and Egyhazi ( 9 c ) . In Table IV, therefore, the results are divided in two groups. The cell material from the cortex of animals 1 and 2 was taken on the ascending part of the learning curve on the 3rd to the 5th day, i.e., during an early part of the learning period. The material from the other two rats was taken on the asymptotic part of the curve on the 9th to the 10th day. I n this case, the animals had reached the maximal number of successful performances per training period on the 6th to 7th day. The increase in the RNA content per neuron in the animals after 3 to 5 days treatment amounts t o 25 to 30%. Qualitatively, the RNA formed in the neurons, i.e., the ARNA, is characterized by a DNA-like base ratio composition with adenine and uracil values around 26. (Rat DNA has the following base composition: 28.6 A, 21.4 G, 21.5 C, 28.4 T.) The cytosine values are remarkably low. The results are statistically significant. These results cannot be obtained from the animals trained for 8 to 9 days that have performed with a maximal number of reaches, i.e., 70 to 80 reaches per period of 25 minutes. The RNA in such animals deviates both quantitatively and qualitatively from that in the first

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group. The relative RNA increase per neuron is 60 to 100% in the cortical cells involved in learning. The base ratio of the RNA formed (ARNA) is similar to that of ribosomal RNA. This is particularly obvious in animal number 4. An important point is that, when the nerve cells within the learning part of the cortex are taken during the early and acute part of the learning process, the relative RNA increase per neuron is small. The TABLE I\’ CHARACTERISTICS O F THE RNA FORMED PER NEURONDURING TRANSFER OF HANDEDNESS CORRELATED TO TRAINING PERIODS A N D PERFORMANCE OF THE ANIMALS(27c)

Animal

Training periods 2 X 25 min/day (days)

Number of successful reaches

Relative increase of total RNA per neuron (%)

ARNA composition A 25.5 G 36.1 c 9.7 U 28.7 A 24.5 G 35.7 C 11.7 U 28.1

3

107

33

5

163

23

8

625

63

A26.2

105

G 34.9 C 16.1 U 22.8 A 21.0 G 35.2 C 24.0 U 19.8

9

1041

nuclear RNA formed, however, has a DNA-like base composition. Thus, stimulation of the genome seems to occur early in a learning situation the animal has not previously encountered. By contrast, when the animals perform well, this RNA of high adenine-uracil values is replaced by an RNA with a ribosomal base composition. At this stage of learning, the relative increase of ARNA is greater than during the earlier phase of learning. Thus a differentiated formation of RNA occurs during a learning period in the neurons involved, and to judge by the character of the RNA formed the early phase seems to be characterized by genetic stimulation.

205

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These changes, quantitativc and cspecially the qualitative, in RNA in the neurons raise the pertinent question as to whether new species of proteins are also formed during the establishment of a new behavior pattern. Although the determinations of the biosynthesis of protein bilaterally in the control centers of the rats involved in change in handedness are still incomplete, some preliminary results may be of interest. Upon separation of proteins from the control cortical center for handedness on the learning side, four protein bands were found that showed no corresponding patterns on the control side. The final conclusion as to whether these reflect protein synthesis must, however, await the result of further incorporation studies. Flexner e t al. (46) give strong support to the view that inhibition of protein synthesis by puromycin injected into the temporal cortex and into the hippocampus destroys memory (maze learning) in mice. In analyzing the mechanism of this phenomenon, they used an antibiotic (heximide) that inhibits protein synthesis by preventing transfer of amino acids from tRNA to the polypeptide, but that does not interfere with the messenger RNA. Heximide was without effect on both short- and long-term memory, and, furthermore, protected memory against the destructive effect of puromycin. The authors therefore drew the conclusion, in agreement with the view expressed by us, that the initial niacromolecular change underlying maintenance of memory involves a change in the quantity of one or more species of messenger RNA. The experiments by Agranoff ( 4 7 ) also show a relationship between protein synthesis and memory. In the second learning experiment, young rats were forced to learn to balance on a thin steel wire, 1 m long, strung at 45" between the floor and a small feeding platform in order to obtain food (9u,b). Training periods occupied 45 minutes per day. Seventy-eight rats were used for the analysis of 12,000 nerve cells. Vestibular Deiters' nerve cells clearly involved in this balance experiment were analyzed. I n these cells, the cytoplasm is large in comparison with the small nucleus. The characteristics of nuclear RNA when whole cells are analyzed are therefore swamped in the bulk of the cytoplasmic RNA since the ratio of nuclear to cytoplasmic RNA is 1:50. No base ratio changes were detected during learning in the cytoplasmic RNA, although an increase from 680 to 750 ppg of RNA was found. However, when the nuclei were isolated and the base ratios of the nuclear RNA were investigated, a clear increase in the ratio A/U of the nuclear RNA was found (1.06 to 1.32), but no significant change in the ratio (G C ) / ( A U). Since no control neurons can be obtained from the same brain in such an experiment, four different types of control experiments were per-

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formed with physiological stimulation and stress involving the vestibular pathways. No significant changes of the A/U ratio were found in these controls although a significant increase of RNA per neuron was found. This signifies that the increase in adenine and decrease in uracil are specific for the learning experiment. The significance of the synthesis of such a nuclear RNA fraction with such high adenine values is more problematical. I n defined parts of chromosomes from Chironomus, RNA has been extracted and found to have an asymmetric composition with high adenine values ( 2 4 ) . This type of nuclear RNA stimulating amino acid incorporation has been found in Euglena ( 2 5 ) , and also in starfish oocytes (16). The conclusion that may be drawn, therefore, is that the nuclear RNA with high A/U ratio found in the second type of learning experiment in rats is chromosomal RNA. Table V presents results obtained on both nerve cell nuclei and glia in such a learning experiment. The increase of the total amount of nuclear RNA of the neurons was estimated to be around 2076, as was also the increase of glial RNA. It is seen that the ARNA fraction of the nerve cell nuclei has a very high adenine and a low uracil content. It may be noted that the ratio (G + C ) J ( A U) of 1.25 does not differ significantly from the corresponding control value of 1.38. The enormous increase of the ratio A/U from 1.06 to 5.9 is, however, very striking. It should be noted, however, that this A/U value of 5.9 is subject to a large error owing to the two small errors in the determination of U in control and learning. Since there is a significant change in A/U between control and learning animals (1.06 and 1.32, respectively) and a significant increase and decrease of adenine and uracil, respectively, in the ARNA fraction, the A/U value of ARNA must be high. I n order to obtain a more reliable value of this latter A/U value, a greater number of measurements are necessary. These are not available a t present, since the results used in this case were not originally prepared for use in an analysis of the ARNA fraction. This may be compared to the A/U ratio of the RNA produced in the Balbiani rings in chromosomes of Chironomous, found to lie around 2 ( 2 4 ) . Since controls to the rat learning experiments demonstrate the significance of the increased adenine and decreased uracil values of the newly formed ARNA in the nucleus, it seems justifiable to assume that this RNA is of chromosomal origin. The neuronal glia reacts in the following way during learning (Table V). The adenine content of the ARNA fraction increases by 70%, and the cytosine content is markedly decreased in comparison with con-

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0

m

TABLE V MICROELECTROPHORETIC ANALYSES OF THE COMPOSITION OF THE NUCLEAR RNA IN THE NEURONSA N D OF THE DEITERS' NUCLEUSIN A LEARNING EXPERIMENT I N RATS(9u,b)a Nucleus

Adenine Guanine Cytosine Uracil

+

(G C)/(A A/U

21.4 26.2 31.9 20.5

+ U)

No. of animals: No. of cells: No. glia samples: a

k 0.44 f 0.45 f 0.77 f 1.01

Learning 24.1 26.7 31.0 18.2

f 0.39 f 0.87 f 0.96 f 1.11

1.38 f 0.042 1.06 It 0.056

1.37 0.049 1.32 f 0.084

5 500

8 900

GLIALRNA

A RNA fraction

Control

38.1 f 3.25 28.8 & 5.75 2 6 . 7 f 6.95 6.4 k 8.08

25.3 29.0 26.5 19.2

1.25 f 0.218 5.9 7.5

f 0.16 f 0.24 f 0.43 f 0.27

Learning

3

?F

2 3

Glia

Base Control

THE

ARNA fraction

28.3 f 0.45 28.8 & 0.31 24.3 & 0.36 18.6 0.21

43.0 27.0 14.0 16.0

1.25 f 0.030 1.32 f 0.020

1.13 f 0.028 1.52 f 0.030

0.69 f 0.076 2.70 f 0.364

5

6

33

42

f 2.83 f 2.24 f 3.12 f 1.88

-2

' ?-

2

kd

Purine and pyrimidine bases in molar proportions in percentages of the sum.

N 0 -3

208

H. H Y D ~ N

trols. The glial ARNA is very similar to the adenine-rich, asymmetric RNA being formed in the neurons during learning and has a (G C)/ A U) ratio of 0.7. The high A/U glial ratio of 2.70 indicates that the ARNA being formed in the glia during learning is of chromosomal origin in analogy with the ARNA formed in the neuron. It may therefore be concluded that factors in the environment (such as the new learning situation not encountered before by the animal) elicit a stimulation of the genome of the glia and the neurons involved and that this response can be characterized in biochemical terms. The analysis of the RNA in two learning situations has thus shown that the response of neurons and glia engaged in the behavior to be established differs from the response to physiological and chemical stimulation in two important respects. First, the content of RNA increases during learning in both neurons and glia. I n physiological and chemical stimulation, the RNA changes are the inverse. Second, asymmetric RNA rich in adenine and uracil is formed in both glia and neurons during learning. After stimulation, the RNA formed has the base ratios characteristic of ribosomal RNA. It is therefore concluded that asymmetric RNA rich in adenine and uracil which is formed in learning is of a chromosomal type.

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IX. The Possible Transfer of RNA between Glia and the Associated Neuron The discussion of the results presented above has led to the conclusion that, during physiological and chemical stimulation, RNA of a ribosomal character is formed both in glia and in neurons. I n the two types of learning experiments, on the other hand, the RNA synthesized during the course of learning is of an asymmetric, adenine-rich type, probably a chromosomal RNA from its composition. This observation leads to the problem of the relationship between the neuron and its surrounding glia a t the molecular level. It is wellestablished that the morphological relationship between them is most intimate, for glial membranes can even invaginate the nerve cell cytoplasm ( 2 6 ) . Furthermore, all parts of the neuronal surface not covered by synapses are covered by glial membranes. From a biochemical point of view, kinetic observations of enzyme activities (27~14,28, 29) lead to the conclusion that the neuron and its glia are coupled energetically, and that they form a functional unit of the central nervous system. After physiological and chemical stimulation, the RNA and protein content and the respiratory enzyme activities increase significantly in the neuron, but decrease in the glia. A study of the kinetics of enzyme reactions indicates that the neuron is the demanding and dominating

BEHAVIOR, NEURAL FUNCTION, AND RNA

209

partner of the functional unit (27b).During increased function it rapidly doubles its rate of oxygen consumption. The glial cells do not change in this respect, but resort partly to anaerobic glycolysis to cover their energy demands. On the basis of these measurements, it seems reasonable to suppose the existence of a mechanism transferring macromolecules between the glia and the neuron. Some speculations on a transfer of RNA between neurons or between glia and neurons have recently been published (30, S1, 3 2 ) . In the following pages, the discussion of such a mechanism is based on the results obtained during induced RNA synthesis. We have chosen the results obtained with vestibular nerve and glial cells after the administration of triap to rabbits (10,9a). The animals were killed 1 hour after intravenous injection of 20 mg triap per kg body weight. Quantitatively, the existing 1500 ppg per nerve cell showed an increase of 570 ppg of RNA, whereas in the glia the control value of 123 ppg decreased to 55 ppg per glial sample. These are dramatic changes and from a quantitative point of view, sufficiently large to serve as a basis for discussion. It has already been concluded that the ARNA fraction synthesized is ribosomal in character. I n Table VI, the data from both the glia and the neurons are listed together with the calculated composition of the corresponding ARNA fractions. First, it can be noted that the A R N A fractions of the neurons and the glia are of the same ribosomal type. Second, the amount of glial RNA is about one tenth of that per nerve cell determined on the basis of the same volume and dry weight. The amount of ARNA fraction determined in the glia is also around one tenth that of the neuronal ARNA fraction. The volume relationship of glia and neurons within the lateral vestibular nucleus is not known with certainty, but the following considerations hold. The large Deiters’ cells used in our studies are situated some hundred microns apart. We are concerned with the so-called neuronal glia enclosing the perikaryon of each nerve cell. Taking the figures for the areas of individual nerve cells calculated by Sholl (33) and Schad6 e t al. ( S 4 ) , a volume of lo6 ps for the neuronal glia belonging to each Deiters’ nerve cell seems reasonable. On the basis of this figure, and assuming the volume of each nerve cell to be about lo5 ps, this would mean a volume ratio of nerve cell to glia of 1 to 10. It is not unreasonable, therefore, to assume that the loss of 55 X 10 ppg of glial RNA corresponds to the determined increase of 570 ppg of RNA in each neuron. These experimental values could reflect a transfer of RNA from glia to neurons. Third, support for this assumption is supplied by the calculated base composition of the glial and neuronal ARNA fractions

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H. H Y D ~ N

(Table VI) . The values are identical. However, experiments with labeled RNA are needed to prove decisively that transfer of RNA from glia to neurons takes place. At this point, the difference in response of neurons and glia in learning and during stimulation might again be stressed. The RNA content increases in both glia and neurons during learning whereas during stimulation it decreases. One comment is that the possible mechanism of RNA transfer between glia and neurons that expresses itself in inverse RNA changes is linked to and regulated by the level of neural function. I n learning, on the other hand, the establishment of the new functional response releases genomic activities that express themselves in their primary RNA products, which are formed in both neurons and glia.

X. The Synthesis of an Asymmetric, Adenine-Rich RNA in Parkinson’s Disease I n biopsy material from the globus pallidus of patients suffering from Parkinson’s disease, the big nerve cells and the surrounding glia have been analyzed for RNA ($6). At a very early stage of the disease, the glia contain a highly aberrant RNA. The adenine value is greatly increased and the values for guanine and uracil decreased; these changes persist. The neuronal RNA shows fewer changes. Later in the development of the disease, and from the time when the overt clinical symptoms emerge and progress, similar though not identical changes are observed in the neuronal RNA. The amount of RNA per nerve cell and in glial samples is significantly increased, averaging 145 ppg RNA/nerve cell (control-116) and 34 ppg RNAJglial sample (control-17). These striking changes in base ratios and in the quantitative amount of RNA were found in the biopsies whether the patient had been ill for 1 year or for as long as 20 years. Due regard was paid to the time relationship between clinical symptoms and biochemical changes in the neurons and glia in the two sides of the same brain. The conclusion derived from analyses was that a biochemical error involving the synthesis of polynucleotides arises in the glia a t a very early stage of Parkinson’s disease. The nerve cell RNA changes were probably secondary to their nature. Table VII records, in the second and third columns, the base changes in the glia during the course of the disease. The changes in the base ratios are calculated on the basis of the increased amount of RNA (ARNA, column 4, Table VII). Clearly an RNA of a highly asymmetric type has been formed in the glia. Especially striking is the unusually large increase in adenine. The ratio (G C)/(A U) is 0.79

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+

2 5

MICROELECTROPHORETIC ANALYSES OF

THE

TABLE V I COMPOSITION OF THE RNA I N NERVE CELLS NUCLEUSFROM RABBITS (Su, 10)s

Neuron

Adenine Guanine Cytosine Uracil

19.7 33.5 28.8 18.0

(G +C)/(A + U ) A/U No. of animals: No. of nerve cells: No. of glia samples:

f 0.37 f 0.39 f 0.36 f 0.18

GLIA O F

THE

DEITERS’

Glia

Base Control

AND I N

Triap

Control

Neuron ARNA fraction

Glia ARNA fraction

20.1 k 0.74 21.9 f 2.15 38.6 f 2.40 19.4 f 0.65

22.5 k 1.61 37.7 f 1.66 21.0 f 2.03 18.8 f 0.91

21.6 k 1.11 37.8 f 3.19 23.0 f 3.33 17.6 f 1.49

1.42 0.090 1.20 f 0.103

1.55 0.138 1.23 f 0.122

Triap

20.5 f 0.31 34.6 k 0.28 26.7 f 0.24 18.2 f 0.20

20.8 28.8 31.8 18.6

1.65 f 0.023 1.09 f 0.021

1.58 f 0.018 1.13 f 0.021

1.54 0.030 1.12 f 0.058

1.53 f 0.090 1.04 f 0.051

18 720

20 800

9

12

50

62

& 0.28 f 0.64 f 0.27 f 0.55

+

+

a Rabbits were treated with 20 mg/kg of tricyano-amino propene and killed 1 hour later. Purine and pyrimidine bases in molar proportions in percentages of the sum. Total amount of nerve cell RNA increased significantly from 1550 ( f78) to 2120 ( f106) ppg/cell. Total amount of glial RNA decreased significantly from 123 ( k6.2) to 68 (k3.4)ppg/sample.

2

C!

212

H. H Y D ~ N

TABLE V I I MICROELECTROPHORETIC ANALYSESOF THE COMPOSITION OF THE GLIALRNA I N GLOFJUS PALLIDUS O F SIX CASES O F PARKINSON'S DISEASE (56)" Base

Controls

Adenine Guanine Cytosine Uracil

+

(G C)/(A A/U

19.0 29.1 33.7 18.2

+ U)

No. of analyses: No. of cells:

f 0.78 f 0.15 f 0.72 f 0.36

Parkinson 30.8 20.3 33.2 15.7

f 0.70 f 0.90 f 1.70 f 0.60

1.69 f 0.044 1.05 f 0.048

1.15 f 0.047 1.96 f 0.087

23 4600

32 4800

ARNA fraction 42.4 2 3.19 1 1 . 5 k 2.75 32.8 f 3.45 13.3 f 1.25 0.79 f 0.093 3.18 0.384

~~

Purine and pyrimidine bases in molar proportions in percentages of the sum. The amount of RNA increased from 17 ( f1.7) ppglglial sample (controls) to 34 (f2.2) ppg/glial sample (Parkinson). TABLE V I I I MICROELECTROPHORETIC ANALYSESO F THE COMPOSITION O F THE RNA I N NERVECELLSIN GLOFJUS PALLIDUS OF SIX CASES OF PARKINSON'S DISEASE (5~56)~ Base

Controls

Adenine Guanine Cytosine Uracil

+

(G C)/@ A/U

18.3 30.5 35.3 15.9

+ U)

No. of analyses: No. of cells:

f 0.42 f 0.44 f 0.60 f 0.36

Parkinson 20.7 28.8 34.4 16.1

f 0.65 f 0.25

f 0.79 f 0.38

1.92 f 0.038 1 . 1 5 k 0.037

1.72 f 0.042 1.29 f 0.051

31 310

39 300

ARNA fraction 30.3 22.2 30.8 16.7

f 5.00 f 3.53 f 5.02 f 2.16

1.13 f 0.185 1.82 f 0.380

Purine and pyrimidine bases in molar proportions in percentages of the sum. The amount of RNA increased from 116 (f5.3) ppglnerve cell (controls) to 145 ( f6.3) ppg/nerve cell (Parkinson).

compared with 1.69 for the control cells in the glia. The AJU ratio is 3.18 compared with 1.05 of the control cells in the glia. I n the neurons of the globus pallidus, the RNA base ratio changes are less striking (Table VIII) but they involve a significant increase of adenine and decrease of guanine in the total RNA. If the change in composition is referred to the increased amount of RNA (ARNA, column 4, Table VIII), in analogy with the treatment of the results from

BEHAVIOR, NEURAL FUNCTION, AND RNA

213

glia mentioned above, this neuronal RNA is characterized by a decreased (G C)J(A U) ratio and a high adenine value, as was also the case with the glial RNA, although the neuronal changes are less conspicuous. All the results suggest that the glia represent the type of brain cell where in Parkinson’s disease the change in RNA formation begins, as well as where the most pronounced RNA changes occur. One explanation may be the observation that the glia have a lower RNA content than that of the neuron. Therefore, a change in the nuclear RNA formation is more easily traced in the glia. To judge by the base ratios and the high adenine content, the RNA in Parkinsonism resembles chromosomal RNA of the type found in Chiranomus and in the starfish. It may be that Parkinson’s disease represents a type of disorder where factors in the environment (e.g., infections) a t a crucial period of the life cycle initiate the release of undesirable genomic activities leading to the biochemical error in the glia. A more tenuous explanation, for which there is a t present no evidence, would be that the ARNA fraction reflects the presence of a virus infection.

+

+

XI. A Working Hypothesis During the last 5 years, an increasing number of speculations on memory mechanisms have been published. The paucity of experimental data on which they are based contrasts markedly with the speculatory nature of the papers. At present no consistent hypothesis exists covering the complex phenomenon of learning, long-term memory, and retrieval of data. I would like to stress that there exists no evidence for a mechanical taping of memory molecules to serve for the storage of neural information. Anyone who has tried to plan and carry out meaningful experiments involving animals and has used a chemical, physical, and morphological approach is immediately confronted with the complexity of the problems and the various levels a t which the phenomena occur. The process of remembering is for us not a recall of fixed and fragmentary details. It is a meaningful reconstruction where the past constantly is being remade by the present. We recognize and remember, with high distinction, complex patterns triggered by key factors. If the evaluating word “meaningful” is struck out we can, however, return to the cellular and molecular level and model experiments. To stimulate speculation the following working hypothesis, based on data discussed in this paper is presented. When neurons forming a pathway in the brain are activated, the frequency of the action potentials will increase. Millions of neurons are activated and integrated. I n learning, the activity is facilitated in certain pathways, but inhibited in others.

214

H. H Y D J ~ N

The hour immediately following the last training period is critical and the animal is sensitive to interfering factors. This is the stage of short-term memory. During this time, some form of fixation occurs so that a long-term memory is established. The long-term memory, once formed, can withstand electric shock, freezing, and coma. I n a suitable model system involving insects, Luco and Aranda (48) obtained evidence that, during the establishment of a new motor behavior, a previously closed pathway becomes open and the delay of the impulses through a ganglion becomes shortened. I n discussing a model for learning and memory, it is important to state that the mechanism should have easy access to the genome. I n highly differentiated cells, only a small part of the genome is active (20% in a cell like the neuron is presumably a high figure). On the other hand, a high specificity would result from an additional activation of only a small fraction of the genes in a cell. Seen from an executive point of view, specificity in neurons is generally placed a t the synapses. I n the hypothesis outlined below, the specificity is placed a t a step before the transmitter of the synapse. In this connection i t is important to point out that only two substances are known to serve as true excitatory transmitters in the central nervous system, namely acetyl choline and glutamic acid. (1) All perception organs generate nerve impulses. When this occurs, an integrative response of the whole central nervous system arises. I n defined brain regions, responsive because of phylogenetic reasons, the activity of certain gene areas will be released as a response to factors in the environment. It is assumed that this is effected by a change of the electrical field strength that releases RNA synthesis by way of conformational changes. From the results on RNA presented in this paper, it seems that the RNA synthesized in both cortical neurons and glia during learning is of the messenger type [Hyd&nand Lange (27d)l. There is a gap in our knowledge of the reaction between bioelectrical potentials and the mechanism governing synthesis of proteins. A pertinent question is whether electrical field strength can initiate synthesis of macromolecules like RNA, in this case primary gene RNA synthesized by gene activation. Katchalsky and Oplatka (49) have recently discussed the possibility that RNA in cells is changed by conformational changes. They found that RNA in solution exhibits hysteresis phenomena with respect to changes in pH. Hysteresis cycles are independent of time but dependent on the history of the system. The changes in free energy are low during phase transit,ions of nucleotides, which is a prerequisite for useful transformations in cells. On the other hand, the energy values were sufficiently high to give an effective signal differing

BEHAVIOR, NEURAL FUNCTION, ANI) RNA

21 5

from the thermal noise. It seems probable, therefore, that genc activation with a synthesis of RNA could be initiated by electrical field strength, since 58 mV corresponds to 2 p H units. (2) This RNA synthesis is supposed to initiate a production of protein, which facilitates the transfer from one neuron to next of the temporal pattern of those electrical frequencies that, a t the onset of this process, activated the gene areas and led to the synthesis of protein. In this way, neuron after neuron can be specified and facilitated permanently in the most complicated chains and pattern. Thus, the specific protein produced and the electrical pattern are both integral parts of a self-organizing system. Changes in electrical field strength may therefore both initiate synthesis of RNA and give conformational changes. A synthetic process seems necessary in a memory mechanism and a question may be whether large RNA molecules are necessary. It does not seem so, since ten sequences each with four nucleotides would allow around 108 permutations. Smaller RNA molecules would have the advantage of being able to move faster than larger molecules in the axoplasm, e.g., down to the synapse. The question is important since RNA but not ribosomes have been observed in the axoplasm of the Mauthner nerve cells in fish [Edstrom (60)1. I n the mechanism suggested here, it is proposed that the specific protein produced triggers the excitatory transmitter. When not acting, the protein combines with a complementary molecule. If the signal arrives that once had led to the formation of the protein by release of RNA synthesis, then the complex dissociates and the transmitter is activated. I n this way, the specificity of the neuron resides in the step before the transmitter. Only a few transmitter substances would be needed. The mechanism visualized has certain traits in common with the antigen-antibody reaction. It may be remembered that plasma cells can produce antibodies against antigens that do not occur in nature but are produced synthetically. At this point it is pertinent to return to Socrates and Locke. Socrates argued that learning is remembering. In modern terminology it would be phrased as activation of a genetically stored memory. By contrast, according to Locke, we are born with a blank mind. Only by experience is knowledge obtained. The tabula rma cerebri of Locke does not exist, and, on the other hand, strict determinism cannot be defended. There exists the new combination of happy ideas, and the solving of problems by pursuing gradients of deepening coherence. I n speculation, there exist other alternatives. Factors in the environment lead to gene activation in brain cells, i.e., synthesis of RNA by

216

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small arcas of the genome. But this RNA can be moclulatcd, perhaps by hysteresis cycles. The protein formed as an end product would gain from this increased specificity. An increased rate of synthesis is a mechanism adding another parameter. It is unlikely, on the other hand, that a neuron should synthesize a random RNA that would then undergo permanent and specific permutations and store information as does a magnetic tape. It does not seem necessary that extensive synthesis of messenger RNA is needed for a protracted period a t the establishment of a new behavior. As is seen from the results quoted, the DNA-like RNA is found during the beginning of the learning process when the animals strive hard a t the new task. This may represent the short-term memory period vulnerable to interfering factors during which specific proteins are first synthesized. When the animals perform according to some criterion and their performance is good, this DNA-like type of RNA can no longer be determined in the neurons. This does not mean that it has ceased to be synthesized and a steady-state synthesis of such specified RNA molecules is assumed to occur. It only implies that our methods of analysis may be insufIicient when the amount of RNA decreases below a certain level and, furthermore, the specific base ratios are not measurable because of the bulk of the ribosomal RNA. It is evident from the quantitative determinations that this type of RNA increases considerably. The existence in the neuron of a cytoplasmic RNA which can undergo permutations, e.g., by methylation, and sustain protein synthesis for a memory mechanism is still a question of interest (40). (3) The reason for this differentiation in RNA response and for the increase of the ribosomal-like RNA during the late stages of learning is worthy of comment. It may be recalled that several examples in the present paper show that increased motor or sensory function is reflected in increased synthesis of RNA. The cells studied were motor neurons or brain stem cells. The RNA produced had a ribosomal base composition, in contradistinction to the RNA rich in adenine and uracil formed in the early stages of learning. It may be concluded that the RNA response of the neurons engaged when a new behavior pattern is established differs only in the beginning of the learning period from the response to other types of physiological stimulation. Only at the beginning is there a formation of RNA rich in adenine and uracil. The specific proteins then synthesized determine and facilitate the activity of neurons in a certain pattern required for the new behavior. I n addition to this specific, new response a t the macromolecular level, there is believed to occur an increased motor and

BEHAVIOR, NEURAL FUNCTION, AND RNA

217

sensory activity in all neurons engaged. This is reflected in the considerable formation of ribosomal RNA in the late stages of learning.

ACKNOWLEDGMENTS These studies have been supported by the Swedish Medical Research Council and the United States Air Force under Grant No. EOAR 63-28 through the European Office, Office of Aerospace Research.

REFERENCES 1. M. Polanyi, in “Proceedings of the Analogy Symposium,” Held at Bellaggio,

1966. To be published. (F. A. Von Hayek, ed.) 2. Plato, “Meno” (R. S. Bluck, ed.), Cambridge Univ. Press, London and New

York, 1961. 3. J. Locke, “Essay Concerning the Understanding, Knowledge, Opinion and Assent”

(B. Rand, ed.), Harvard Univ. Press, Cambridge, Massachusetts, 1931.

4. J. H. Fabre, “Les merveilles de l’instinct chez les insectes.” Paris, 1913. 6. R. Ardrey, “African Genesis,” Dell, New York, 1961. 6. I. Krechevsky, Psychol. Rev. 39, 516 (1932). 7. T. S. Kuhn, “The Structure of Scientific Revolutions.” Univ. of Chicago Press, Chicago, Illinois, 1962. 8. H. G. Williams-Ashman, Biochem. J. 97,22P (1965). 8u. J. R. Tata, This Series, Vol. 5. 9. H. Hyd6n and E. Egyhazi, Proc. Nutl. Acud. Sci. US. (a) 48, 1366 (1962); (b) 49, 618 (1963); (c) 52, 1030 (1964); (d) to be published. 10. E. Egyhazi and H. Hydkn, J. Biophys. Biochern. Cytol. 10, 403 (1961). 11. H. Hyd6n and A. Pigon, J. Neurochem. 6,57 (1960). 18. B. W.Moore and D. McGregor, J. Biol. Chem. 240, 1647 (1965). 13. E. Koenig and G. B. Koelle, Federation Proc. 20,344 (1961). 14. J.-E. Edstrom: (a) Biochim. Biophys. Acta 12, 361 (1953); (b) Microchem. J. 2, 71 (1958); ( c ) J . Biophys. Biochem. Cytol. 8, 39 (1960); (d) J . Biophys. Biochem. Cytol. 8, 47 (1980); (e) in “Methods in Cell Physiology” (D. M. Prescott, ed.), Vol. I. Academic Press, New York, 1964. 16. H. Hydkn, Nature 184, 433 (1959). 16. J.-E. Edstrom, W. Grampp, and N. Schor, J. Biophys. Biochern. Cytol. 11, 549

(1961). 17. S.-0. Brattggrd and H. HydCn, Acta Radiol., Suppl. 94 (1952). 18. E. Koenig and S.-0. BrattgLrd, Anal. Biochem. 6, 424 (1963). 19. S.-0. BrattgCd and B. Daneholt, J. Neurochem. 13,913 (1966). 20. H. Hydkn, K. Bjurstam, and B. S. McEwen, Anal. Biochem. 17, 1 (1966). 81. A. H.Coons, Ann. N.Y. Acad. Sci. 69, 658 (1957). 22. G. Brawerman, L. Gold, and J. Eisenstadt, Proc. Natl. Acud. Sci. U.S. 50, 630 (1963). 23. B. H. Hoyer, B. J. McCarthy, and E. T. Bolton, Science 140, 1408 (1963). 24. J.-E. Edstrom and W. Beermann, J. Cell Biol. 14, 371 (1962). 26. G.Brawerman, Biochim. Biophys. Acta 76, 322 (1963). 26. A. Hem, J. Biophys. Biochem. Cytol. 4, 731 (1958). W . H. Hydkn and P. W. Lange: (a) 4th Intern. Neurochem. Symp. Pergamon Press, Oxford, 1960; (b) J. Cell Biol. 13, 233 (1962); ( c ) Proc. Natl. Acad. Sci. U S . 53, 946 (1965); (d) Naturwissenschuften 53,64 (1966).

218

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98. A. Hamberger, Acta Physiol. Scand, Suppl. 203 (1963). 99. A. Hamberger and H. Hydhn, J . Cell Bwl. 16,521 (1963). 30. T.K.Landauer, Psychol. Rev. 71,167 (1964). 31. R. B. Roberts, Neurosci. Res. Progr. Bull. 2, 39 (1964). 3.2. F. 0.Schmitt, Neurosci. Res. Progr. Bull. 2, 43 (1964). 33. D.A. Sholl, in “Structure and Function of the Cerebral Cortex” (D. B. Tower and J. P. SchadB, eds.), p. 21. Elsevier, Amsterdam, 1960. 34. J. P. Schad6, H. van Backer, and E. Colon, in “Progress in Brain Research” (D. Purpura and J. P. SchadB, eds.), Vol. 4 Elsevier, Amsterdam, 1964. 36. G. Gomirato and H. Hydbn, Brain 86,773 (1963). 36. E. Egyhazi and H. HydBn, Life Sci. in press. 37. M. Jacob, J. Stevenin, R. Jund, C. Judes, and P. Mandel, J. Neurochem., 13, 619 (1966). 38. E. Egyhazi, Biochim. Biophys. Acta 114,516 (1966). 39. R. Ekholm and H. HydBn, J. Ultrastruct. Res. 13,269 (1965). 40. H. Hydkn, in “The Cell’’ (J. Brachet and A. E. Mirsky, eds.), Vol. IV. Academic Press, New York, 1960. 41. A. Lajtha, Intern. Rev. Neurobiol. 6 , l (1964). 49. B. 5.McEwen and H. HydBn, J . Neurochem. 13,823 (1966). 43. H. Hydkn and B. S. McEwen, Proc. Natl. Acad. Sci. U S . 55, 354 (1966). 4.4. A. L. Rubin and K. H. Stenzel, Proc. Natl. Acad. Sci. US. 53, 963 (1965). 46. H. Hydkn and S. Larsson: (a) J . Neurochem. 1, 134 (1956); (b) Proc. 9nd Intern. Symp. X-ray Microscopy and X-ray Microanalysis, Stockholm, 1960, p. 51. Elsevier, Amsterdam, 1960. 46. L. B. Flewer and J. B. Flewer, Proc. Natl. Acad. Sci. US.55, 369 (1966). 47. B. W. Agranoff and P. D. Klinger, Science 146, 952 (1964). 48. J. V.Luco and L. C. Aranda, Nature Wl, 1330 (1964). 49. A. Katchalsky and A. Oplatka, Neurosci. Res. Progr. Bull. 3, 15 (1965). 60. J.-E. Edstrijm, D. Eichner, and A. Edstrom, Biochim. Biophys. Acta 61, 178 (1962).

The Nucleolus and the Synthesis of Ribosomes ROBERTP. PERRY Department of Molecular Biology, The Institute for Cancer Research, Philadelphia, Pennsylvania

I. Characterization of the Genes Coding for Ribosomal RNA A. Relation to Nucleolar Organizer . . . . . . . B. Properties of the rRNA Cistrons. . . . . . . C. Regulation of the Transcription of rRNA Genes . . 11. Synthesis of the Precursor of Ribosomal RNA. . . . A. The Nucleolus as the Site of Synthesis . . . . . B. Evidence That the Precursor is a 45 S Component . C. Properties of the 45 S Component . . . . . . D. Association of Precursor with Other Cell Constituents 111. Subsequent Events in the Formation of rRNA . . . A. Cleavage of the Precursor. . . . . . . . . B. Synthesis of Ribosomal Protein . . . . . . . C. Relation to Nucleolar Ultrastructure . . . . . IV. Appearance of Ribosomes in the Cytoplasm . . . . A. Subunits. . . . . . . . . . . . . . B. Distinctive Properties of New Particles. . . . . C. Involvement with Polyribosomes. . , . . . . D. Turnover of Ribosomes . . . . . . . . . V. Synopsis. . . . . . . . . . . . . . . Addendum . . . . . . . . . . . . . . References . . . . . . . . . . . . . .

.

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.

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. . . .

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220 220 221 224 228 228 229 231 232 233 233 235 238 239 239 243 245 249 249 251 253

One of the most conspicuous structures in cells of higher organisms is the nucleolus. Because of its prominence and ubiquity, this organelle has long been an intriguing subject of study for those interested in biological structure-function relationships (1). Over the years, advances in our knowledge of the nucleolus have occurred in recognizable stages that can be correlated with methodological developments in genetics, cytology, and biochemistry ( 2 ) [see also reviews by Vincent (S), Sirlin ( 4 ) , and Busch et al. ( b ) ] . During the recent upsurge of molecular biology, sufficient evidence has accumulated to attribute to the nucleolus a key role in the biosynthesis of ribosomes. Thus, much of the research that 219

220

ROBERT P. PERRY

previously centered on nucleolar function has now turned to the study of ribosome synthesis in higher organisms. In this paper, I review briefly some of the various lines of evidence that have implicated the nucleolus with ribosome synthesis, trying, where possible, to indicate the liiiiitations of current concepts as well as the mechanisms that appear to be firmly established. Some attention is given to other metabolic activities that might be associated with the nucleolus, but in these areas no attempt a t completeness is made. In addition, some aspects of ribosome formation are discussed that, a t the moment, do not seem to involve the nucleolus, but that will surely be necessary for a full understanding of the functional role of the ribosome in the cell.

1. Characterization of the Genes Coding for Ribosomal RNA A. Relation to Nucleolar Organizer Direct evidence linking the genetic material responsible for nucleolus formation with that responsible for the synthesis of ribosomal RNA was provided by the finding of Brown and Gurdon (6) that embryos of the mutant of Xenopus Zaevis that lacked the ability to form nucleoli were also incapable of synthesizing rRNA. Homozygotes of this mutant die a t the tail-bud stage, possibly because the maternal ribosomes are no longer able to support the embryo’s demands for protein synthesis; heterozygotes develop normally. Further elucidation of the relationship between the DNA that codes for rRNA and the DNA associated with the nucleolus or belonging to the nucleolar organizing regions of the genome has been achieved using the technique of DNA-RNA hybridization (7). I n these experiments, isotopically labeled rRNA is allowed to anneal with DNA from various sources. Determination of the maximal amount of rRNA that can form specific duplexes with a given quantity of DNA allows an estimate of the number of cistrons or DNA complements coding for rRNA in the particular DNA sample. Two methods for distinguishing between nucleolar and nonnucleolar DNA have been used. One method, the first to be tried, involves cellular fractionation to separate nucleoli from the rest of the nuclear material and then extraction of DNA from each fraction (8, 9 ) . This technique requires that one obtain pure nucleoli still attached to their chromosomal organizer segments and uncontaminated by other chromatin. Since this is an extremely difficult requirement to meet, it is not surprising that the results have been contradictory and not decisive. With HeLa cell DNA, the nucleolar fraction was found to be enriched in ribosomal cistrons

T H E NUCLEOLUS AND T H E SYNTHESIS OF RIBOSOMES

22 1

fivefold over total nuclear DNA ( 9 ) , whereas in pea embryos no difference between nucleolar and total nuclear DNA was observed (8). Even in the case of the HeLa DNA, the enrichment factor was much less than expected and the possibility had to be considered that, although the rRNA cistrons were concentrated in the nucleolar DNA, there might also be some nonnucleolar genes capable of, but not normally functional in, producing rRNA (9, 10). These uncertainties were resolved by the second method, in which genetic means were used to obtain variants of the same species carrying different dosages of the nucleolar organizer. Ritossa and Spiegelman (11) determined the number of DNA complements of rRNA in stocks of Drosophila melanogaster containing X chromosomes either deficient or duplicated for a short piece of heterochromatin containing the nucleolar organizer. Appropriate choice of material supplied genomes possessing one, two, three, and four organizers. Birnstiel e t al. ( 1 2 ) made similar measurements on wild-type Xenopus embryos and on homozygotes and heterozygotes carrying the anucleolate mutation. As seen in Fig. 1, the saturation plateaus obtained with these organisms are directly proportional to the number of their nucleolar organizers, thus providing very convincing evidence for the assertion that the DNA complementary to rRNA is confined to a region of the genome encompassing the organizer. I n Drosophila it appears that the “bobbed” locus, which maps within the aforementioned heterochromatic region of the X chromosome, is the site of the organizer (13). Examination of the DNA from a series of “bobbed” mutants has revealed a varying number of rRNA cistrons (Fig. 2), all less than that characteristic of the wild-type genome. These mutants are characterized by abnormally short bristles. Since the bristles are produced rapidly and require an intense protein synthesis, it is not difficult to imagine how the abnormality might be attributable to a ribosome deficiency.

B. Properties of the rRNA Cistrons An actual isolation of the rRNA cistrons was achieved with the Xenopus system by banding the DNA in CsCl density gradients ( 1 2 ) . A satellite band of buoyant density about 0.025 gm/cm3 greater than the major band and comprising roughly 0 . 1 5 4 2 % of the total DNA was demonstrable (Fig. 3 ) . This satcllite, which was not found in the DNA of the anucleolatc mutant, was many times more active in forming hybrids with rRNA than the major DNA component. The practicability of isolating the satellite is itself indicative that the rRNA cistrons exist in extended clusters. From molecular weight estimates it is apparent that these clusters can be isolated in groups of three or more, thus pro-

222

ROBERT P. PERRY

viding convincing evidence that the cistrons occur in a repetitive linear sequence within the genome. An analogous arrangement of the rRNA cistrons has been found in bacterial systems (14-16'). The percentage of the genome devoted to rRNA production is relatively constant for the various types of organisms studied, although compared to the microorganisms the absolute number of cistrons is greater by a t least an order of magnitude (Table I). In almost all cases where the two rRNA components were individually studied, the cistrons complementary to the large rRNA component are distinct from those responsible for the small rRNA component; no evidence of competition is found and hybridizations with both components are completely additive (13,14, 18, 19). The question naturally arises: are the multicistrons for each rRNA component completely redundant or is there some degree of uniqueness within each population? Although an unequivocal answer cannot be given a t the moment, the experiments with the a-d 0-? I

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223

THE NUCLEOLUS AND THE SYNTHESIS OF RIBOSOMES

“bobbed” Drosophila mutants (IS) and with a “split nucleolar organizer” in Chironornous (2.2, 2s) suggest that the degree of redundancy is probably high. This follows from the fact that in these instances the deletion of a major portion of organizer DNA, directly shown to be equivalent to a deletion of rRNA cistrons in Drosophila, does not necessarily constitute a lethal event. Moreover, there is no measurable difference in the RNA of nucleoli formed by organizers located on different chromosomes (24). The data also suggest a rather high degree of interspersion of the two ribosomal components; they do not seem to be transcribed from blocks of DNA on opposite sides of the organizer. Whether the cistrons for the two components are in an exactly alternating sequence has not been definitely established. Such an arrangement would predict an equal number of cistrons for the two components, and, although some of the data in Table I do not seem to support this prediction, the estimates of cistron number are not sufficiently precise to allow one to draw strong conclusions from them ( 2 5 ) . Indeed, from the nature of the 45 S precursor and the coordinated synthesis of the two rRNA components dur-

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FIQ. 1. Saturation curves for annealing of radioactively labeled ribosomal RNA to DNA’s from organisms with varying dosages of nucleolar organizer (n). (a) Drosophila system [from Ritossa and Spiegelman ( 1 1 ) l . Note that the saturation level of RNA binding for wild type (2) is the same (- 0.27%) for males, where there is one organizer on the X chromosome and one on the Y , as it is for females, where both organizers are on the X. (b) Xenopus system [from Wallace and Birnstiel (1811. In both systems, the number of DNA stretches complementary to ribosomal RNA is directly proportional to the number of nucleolar organizers, suggesting that the ribosomal cistrons are contained within the organizer region.

224

ROBERT P. PERRY .20

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FIQ.2. Rank order distribution of the percentage of DNA forming hybrid with ribosomal RNA where DNA is obtained from various bobbed stocks of Drosophilu mehogaster. The values are referred t o a single nucleolar organizer so that wildtype stocks (closed symbol) average about 0.14%. Note that the bobbed stocks (open symbols) average about one-half of this. [From F. Ritoasa. See (IS) for experimental details.]

ing embryogenesis (see below) the arrangement of cistrons in an alternating sequence appears to be a likely possibility. Perhaps the stretches coding for the 5 S RNA, which is associated with the large subunit (cf. Section 111, A ) , act as spacers for each cistron or pair of cistrons.

C. Regulation of the Transcription of rRNA Genes The precise molecular mechanisms regulating the expression of the rRNA genes are unknown. Current notions regarding the general control

THE NUCLEOLUS AND THE SYNTHESIS OF RIBOSOMES

225

mechanisms of genic transcription involve the interaction of the DNA with regulator protein (26, 27), and/or modification of chromosomal structure, e.g., compaction (heterochromatiaation) vs. extension (puffing) (28, 29) mediated by interactions of the DNA with basic proteins, polysmincs, specific RNA's, or RNA-protein coniplcxcs (30-34). Sincc the regulation of ribosoinal transcription may entail the siinultaneous 111

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FIQ.3. Tracings of photographic records obtained from analytical centrifugation of Xenopus DNA samples. The CsCl gradient (at a mean density p = 1.720 gm cm-') contained the following amounts of DNA (MW 1MO x 10') : homozygous mutant, 87 &g; phenotypically normal sibs of the same matings, 69 pg; wild type (without marker), 102 pg; wild type with denatured P. aeruginosa as a density marker ( p = 1.740 gm cm-'1, 95 pg. As an internal standard for the determination of the satellite mass, a 500-fold dilution of the wild-type DNA (without marker) was used in parallel experiments. The major DNA band (1.698 gm cm-') shows a considerable spread owing to the fact that it is present in 100-fold excess. The tracings have been displaced vertically t o facilitate comparison. Similarly, the internal standard has been shifted to a region of high density. A satellite at 1.723 gm cm-' is present in all preparations at high DNA input, with the exception of the homorygous mutant DNA. In the wild type this satellite represents some 0.15-0.2% of the total DNA. [From Birnstiel et al. ( l a ) . ]

226

ROBERT P . PERRY

TABLE I SOMEREPRESENTATIVE DATAO N HYBRIDIZATION OF VARIOUSDNA's HOMOLOGOUS RIBOSOMAL RNA's

Organism

Per cent of total cell DNA comprising rRNA cistrons (j)" ~

Total number of ribosomal cistrons per haploid genome (both components)*

WITH

Ratio of number of cistrons: large rRNA component Refersmall rRNA component ences

~~

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0.005-0.02

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0.27

260-1300

0.06-0.11

1200-2000

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0.32 0.38 0.42-0.65

160 6-14 8-22

0.5-0.7

0.9

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a j = 100 (weight of rRNA cistrons, single strand)/(weight of total cell DNA, both strands) = ratio a t saturation plateau of pg RNA hybridized to pg DNA in incubation mixture, X100. Either the value obtained when both ribosomal components were used together in the hybridization mixture or the sum of the two saturation values when the ribosomal components were measured individually. f2N/A2 for the two components, 1 and 2, bcalculated as the sum: jlN/A1 measured individually, where N is the total molecular weight of DNA per haploid genome and A is the molecular weight of the rRNA component. When the components A2), where j is are not measured individually, this figure is estimated as 2fN/(A1 the hybridization value for the mixture. The two figures would be the same only if the number of cistrons for each component is equal. Corrected to human haploid, assuming a modal ploidy for HeLa of 3.5.

+

+

repression or derepression of hundreds or thousands of genes, one is attracted to models that can accomodate this feat with comparative simplicity, such as those specifying structural changes. In this regard we note that the nucleolus formed by the Y chromosome of Drosophila appears to contain some markedly different loop structures than those formed by the X (28), suggesting a possible difference in the degree of compaction between two comparable sets of genetic material. Furthermore, the virtual absence of two classes of histones in anucleolate X e q u s embryos could imply that the state of compaction is itself regulated by specific interactions with basic proteins (36). It is perhaps worthwhile here to enumerate a few examples in which the regulation of the activity of the ribosomal genes is readily apparent.

THE NUCLEOLUS AND THE SYNTHESIS OF RIBOSOMES

227

(i) Systems with Variable Amounts of Nucleolar Organizer. In the heterozygous (1-nu) embryos of Xenopus, the wild-type genes produce twice as much rRNA as do the same genes when present in homozygous wild-type individuals (6). Similarly, the presence of an extra sex chromosome (X or Y) , and hence an extra nucleolar organizer, in certain stocks of Drosophila does not cause an increase in the amount of rRNA per organism (28, 3 6 ) . I n the case of X!enopus, there is apparently an augmentation of the transcription activity, whereas with Drosophila we have an example of a suppression. A common element in these two systems is that the deletion or addition involves a unitary block of rRNA genes comprising one member of a set of alleles. It is well-known that the expression of alleles may be mutually exclusive (S7, 38) and it is conceivable that the control of rRNA transcription belongs in this general class of regulatory phenomena. Moreover, this control can be modified by the specific chromosomal arrangement of the organizer [cf. ( 2 )I. (ii) Activation and Suppression of rRNA Cistrons during Development. I n unfertilized eggs the rRNA genes behave as the majority of their neighbors : they are quiescent. After fertilization they are reactivated during a particular developmental stage of the embryo that seems to vary according to the amount of rRNA available in the form of cytoplasmic and yolk reserves (39, 40). I n mammalian embryos, nucleolar development-and apparently rRNA synthesis-begins during an early cleavage stage (S9), whereas in amphibia (41), echinoderms (429, and others (40) it occurs a t the onset of gastrulation. I n these latter systems, the reserves are achieved by an intensive rRNA synthesis during maturation of the oocyte (43-46). It was demonstrated that the transplantation of nuclei from tissues actively engaged in rRNA synthesis into enucleated unfertilized eggs leads to cessation of the transcriptive activity of the rRNA genes and disappearance of nucleoli ( 4 6 ) . It is not yet clear whether this regulation arises from a specific interaction between the rRNA genes and cytoplasmic constituents or, more remotely, from the highly altered physical and metabolic state of the nucleus (increased volume, rapid division). During the reactivation phase both rRNA components are synthesized coordinately, suggesting that they are under common control. (iii) Variations of rRNA Synthesis during the Cell Cycle. The rRNA genes appear to react to the same set of regulatory stimuli as does the rest of the genome. During cell division, when the chromosomes are in a condensed state, there is a cessation of transcriptive activity that is usually accompanied by a dissolution of the nucleolar body (4760).I n some systems, the nucleoli remain active sites of RNA synthesis

228

ROBERT P. PERRY

during prophase after most of the other chromosomal activity has stopped (51). When cell division is completed, there is a rapid formation of nucleolar structure (52) and a resumption of nucleolar RNA synthesis ( 5 1 ) ,which we may presume marks the onset of rRNA formation. I n exponentially growing cells, the rate of nucleolar synthesis seems to be relatively constant throughout G1 and part of S phase [Perry (M), Vincent ( 2 6 ) , (53-55)], indicating that the rRNA genes are being transcribed a t a relatively constant rate. I n some cases, a depressed activity has been observed in S phase, possibly related to the period of replication of the nucleolar genes (54-56).I n the latter part of the cycle, there may be a burst of rRNA synthesis [Vincent ( 2 5 ) , Perry (261, (5.91, although the maximum rates observed are not always double the initial rates, leading one to suspect that, after duplication, some of the rRNA genes are in a partially repressed state.

II. Synthesis of the Precursor of Ribosomal RNA

A. The Nucleolus as the Site of Synthesis Realization that RNA synthesized in the nucleolar region of the nucleus might be a precursor to rRNA came initially as a result of studies designed to clarify the relationship between nuclear and cytoplasmic RNA [see (60) for refs.]. Once i t became evident that the bulk of the cytoplasmic RNA is ribosomal, then the demonstrations of nucleolar dependence of cytoplasmic RNA synthesis (57-59), or close similarity in base composition between nucleolar and cytoplasmic RNA (60), were readily interpretable in terms of a nucleolar origin for rRNA. Early attempts a t cellular fractionation by the phenol method (see article by Georgiev in this volume) and separation of the RNA into high and low molecular weight fractions (61) also pointed towards this conclusion. Strong confirmation for this hypothesis was the demonstration that low doses of actinomycin D, which selectively inhibit nucleolar RNA synthesis (66), also selectively inhibit the synthesis of the precursor to rRNA ( 6 3 ) . That the nucleolus contains the necessary enzymatic apparatus for the polymerization of RNA (64, 65) and its subsequent methylation (66) has also been shown. Whether the nucleolus is a site of synthesis of other types of RNA is still the subject of some controversy. From a correlated autoradiographic and biochemical study of the RNA synthesized in the presence of low doses of actinomycin, it was concluded that in mammalian cells messenger and transfer RNA are made predominantly in the extranucleolar or chromatin portion of the nucleus (10, 67,68). Analysis of the RNA produced by the anucleolate Xenopus embryos (6, 69) and differentially

THE NUCLEOLUS AND THE SYNTHESIS OF RIBOSOMES

229

extracted Vicia faba cells (70) also indicates that tRNA synthesis is extranucleolar. On the other hand, from results of a similar correlative study in Smittia utilizing substituted benzimidazoles, Sirlin and coworkers (71) concluded that the nucleolus is an active synthetic site of tRNA. Nucleolar involvement in tRNA synthesis has also been proposed for the nucleoli of starfish oocytes (72) and an aquatic fungus (73), although, in view of the finding that the submethylated 5 S RNA associated with nucleoli is not tRNA ( 7 4 ) ,some of these results should be re-evaluated. Nevertheless, it is possible that part of the disagreement is due to variation among organisms with respect to the proximity of rRNA and tRNA genes. For example, according to hybridization tests, the tRNA cistrons in Drosophila lie outside the sc8-sc4 region, which contains the nucleolar organizer ( I S ) , whereas in B. subtilis the rRNA and tRNA cistrons map very close to each other (76).

B. Evidence That the Precursor is a 45 S Component Characterization of the precursor in several cell types became possible as methods were developed for the extraction of rapidly labeled nuclear RNA’s in an undegraded form (76). When pulse-labeled RNA from a wide variety of sources, including cultured mammalian cells (63, 67, 76-79), amphibian oocytes and embryos (6, 80), developing sea urchins (4.29, rat liver (81, 82),tumor tissue (82, 83), and cotton seedlings (8 4) , is submitted to zonal centrifugation on sucrose gradients, the radioactivity profile is readily distinguishable from the 28 and 18 S rRNA components. Under favorable circumstances, the rapidly labeled RNA can be seen to consist of a 45 S component, some polydisperse components which appear to be the most rapidly synthesized of all, and a 4 S component (Fig. 4a). After slightly longer periods of labeling, a 35 S component is visible as well (Fig. 4b). Several lines of evidence combine to indicate that the 45 S component1 is the early precursor of rRNA. These are:

(i)Actinomycin-chase kinetics (63, 78, 79, 83, 86, 87). I n these experiments, cells were pulse-labeled with RNA precursor and then “chased” in the presence of sufficient actinomycin D to inhibit ‘ A s used here, the term “45 S component” refers to the relatively monodisperse RNA component that can be observed after appropriate labeling intervals in the 45 S region of a sucrose gradient (Fig. 4b). I t should not be confused with that portion of the heterogeneous RNA (Fig. 4a) sedimenting in the 45 S region. These two species can be readily distinguished under conditions where rRNA synthesis is selectively depressed (68) or b y further fractionation of nuclei prior to extraction of the RNA (86).

230

ROBERT P. PERRY

subsequent rRNA synthesis completely. During the chase, the 45 s component disappeared and labeled 28 and 18 S rRNA components appeared.2

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FRACTION NUMEtR (b) FIQ.4. Sedimentation diagrams of RNA from strain L cells pulse-labeled for various intervals with H3-cytidine. Solid curves: absorbance at 260 mp; dashed curves: radioactivity. RNA was extracted with 0.5% sodium dodecyl sulfate and phenol a t 60°C. Centrifugation for 3 hours at 39,000 rpm (SW 39) in 10 t o 40% (w/V) sucro8e gradients. (a) Very brief pulses. Radioactivity is plotted as percentage of total label to emphasize the progressive increase in the ratio of 45 S to heterogeneous components. (b) 30-minute pulse showing preponderance of 45 and 35 S components.

'The proportion of the original pulse label ultimately observed in ribosomal components has varied among different experiments from almost 100% (63, 79, 80) to intermediate values (83,86) t o almost zero (89). For the most part, this variation can be traced to the duration of the pulse, longer pulses giving higher yields than short pulses [see discussion6 in (78, 88, 90, 9 1 ) l . It has also been proposed that a secondary effect of actinomycin is to prevent entrance of newly formed rRNA into the cytoplasm (86,88,91).

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(ii) Selective inhibition of the 45 S component with low doses of actinomycin D (63, 6 7 ) . Subsequent chasing demonstrated complete inhibition of rRNA synthesis. Under these conditions the polydisperse messenger and 4 S components are still synthesized (68)* (iii) Base composition analyses (92), showing that the 45 S component resembles a composite of the 28 and 18 S components. (iv) Competitive hybridization between the 45 S and rRNA components, indicating a high degree of similarity in base sequences (68). (v) Diminished synthesis of 45 S RNA in the anucleolate mutants of Xenopus, which are incapable of rRNA synthesis ( 6 ) .Other rapidly labeled components, closely resembling those made in the presence of low doses of actinomycin, are synthesized. (vi) Proportionately reduced synthesis of 45 S RNA in early stages of embryogenesis that are also characterized by a lack of rRNA synthesis (42). The question of the universality of a large 45 S-like precursor to rRNA is a difficult one to answer. For those systems in which it has not been demonstrated, one can usually invoke the possibility of degradation during extraction and fractionation procedures. This is especially true for bacteria where there are appreciable differences in the profiles of rapidly labeled RNA, depending on whether one prepares extracts by alumina grinding (93, 94) or more gentle methods (96).

C. Properties of the 45 S Component With the advent of techniques for isolating pure nucleoli (86, 96) it is now possible to obtain 45 S RNA relatively free from contamination with other RNA components and in sufficient quantities for molecular weight determinations and other physical analyses. Although no molecular weight data have been published a t the time of this writing, we can expect them imminently. An estimation made on the basis of the sedimentation coefficient3 yields a molecular weight of 4.4 X loG,a value 'According to the theoretical formula for random coils (97) and an empirical formula for RNA from various sources (981, molecular weight is approximately proportional to the square of the sedimentation coefficient. With this dependency, the molecular weight of a 45 S component is calculated to be 4.4 x loa when the 28 S component is taken as 1.6 x 10" (99). I t should be emphasized that this is only an estimate. The sedimentation coefficient has not yet been obtained under the precisely defined conditions of the analytical ultracentrifuge, and also we cannot rule out the possibility that the 45 S component possesses an atypical configuration. The methylation of rRNA, which has been reported to occur at the 45 S level (86, loo), probably alters the sedimentation behavior of the molecule, but only slightly (101).

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consistent with the idea that the 45 S molecule contains two 28 S pieces of MW 1.6 X lo6 and two 18 S pieces of MW 0.6 )( lo6.Such a structure is also in agreement with analyses of 45 S base composition (92) and with the finding that no appreciable part of the 45 S molecule is lost during its transformation to the 28 and 18 S components (102) (Section III,A), although these data support equally well the notion that the 45 S component contains a single 28 and 18 S pair. It has not been possible to dissociate the 45 S component into smaller pieces under conditions favoring hydrogen bond breakage, e.g., heating in the presence of formaldehyde or treatment with high concentrations of urea or dimethylsulfoxide (87, 92, 103). Thus it would seem probable that the 45 S precursor is a covalently continuous complex of the two rRNA components. This would be compatible with the concept of alternating rRNA cistrons discussed in Section I,B if the 45 S component were derived from either one or two pairs of such cistrons. However, such a scheme is not without controversy. Busch e t al. (96) have suggested that there are two 45 S rRNA precursors, one for the 28 S component, which remains associated with the nucleolus during its isolation, and another for the 18 S component, which is lost during the isolation procedure. On the other hand Georgiev (104) has interpreted results with a thermal-phenol fractionation procedure to mean that the 45 S molecules are precursors only of the 18 S components. Further work is needed before a decisive answer to this question can be given.

D. Association of Precursor with Other Cell Constituents During the process of transcription, there is a transitory period when the 45 S RNA component must be associated with a t least part of its DNA complement. Therefore it might be expected that, under especially mild extraction conditions, some 45 S rRNA precursor could be isolated as part of a DNA-RNA complex. Yet, although isolation of complexes of DNA with rapidly labeled RNA has been achieved with certain types of animal cells (105, 106), until now no one has sucessfully demonstrated the presence of 45 S RNA in such complexes. Owing to the relatively low number of rRNA cistrons, i t may be that, in order for the newly synthesized 45 S RNA to reach a detectable level, each cistron must be transcribed several times. In this case, the proportion of 45 S RNA in the form of a hybi*id with DNA would be relatively low. Tamaoki and Mueller (86, 107) have reported that a combination of deoxycholate, DNase, and dextran sulfate can be used to isolate the 45 S component as part of a heterogeneous group of ribonucleoprotein com-

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plexes sediinenting between 60 and 100 S. These complexes have the same effective charge density as riboe~niesand contain some newly synthesized protein. It was proposed that they might represent early stages in the formation of the ribosome particles. However, since the proteins have not yet been shown to belong to any specific class, and since dextran sulfate causes fairly extensive degradation of ribosomes and their subunits ( 8 6 ) ,the full significance of these complexes is difficult to ascertain.

111. Subsequent Events in the Formation of rRNA A. Cleavage of the Precursor Details concerning the transition in rRNA from a 45 S precursor molecule in the nucleolus to the 28 and 18 S components of the cytoplasmic ribosomes have been obtained from the following types of studies: ( a ) kinetics of incorporation of nucleosides into the various RNA components (10, 78, 79, 83, 88) ; ( b ) alteration of the pattern of incorporation with analogs and inhibitors, e.g., 8-azaguanine (10, 671, thioacetamide (96,108), or puromycin (67,9f, fog), which block specific stages in the transformation sequence; and ( c ) use of refined cellular fractionation techniques to separate cellular structures containing different intermediates (80,85,96). In pulse-chase experiments, including those in which continued incorporation from nonexchangeable nucleotide pools is prevented with actinomycin D, one observes a sequential labeling of 45 S, then 32-35 S and 18 S, and finally 28 and 18 S components (78, 79) (Fig. 5). When part of the RNA guanine is replaced by 8-azaguanine, the early step in this conversion can still occur, but the later step is greatly retarded. Since under these conditions much of the labeled RNA accumulates in the nucleolus rather than the cytoplasm, it was concluded that the early step in the conversion of 45 S component occurs in the nucleolus (10). Rather convincing evidence for this point was obtained by analysis of the labeling of RNA’s from isolated nucleolar, nucleoplasmic, and cytoplasmic fractions (80, 85, 96). I n these cases, sequential labeling of the 45 and 3 2 3 5 S RNA could be observed in a series of nucleolar fractions prepared from cells labeled for different periods or chased in the presence of actinomycin. Moreover, in some cases (80, 8 5 ) , little or no 18 S RNA appears to be associated with the nuclei, indicating that upon cleavage of the 45 S molecule there is a rapid exit of the 18 S component to the cytoplasm. This is in accord with the early appearance of labeled 18 S RNA (67,88), or, more specifically, particles bearing 18 S RNA (10,110), in the cytoplasm (see Section IV,A) . The later stage of conversion involves the transition from 32-35 s

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to the 28 S component. Penman et al. (86) observed that, whereas the major RNA component in the nucleoplasm of HeLa cells sediments a t 28 S, the bulk of the RNA from nucleoli sediments faster than 28 S. Similarly, Busch and co-workers extracted 28 S RNA from both nucleoli and nucleoplasm of Walker tumor cells and 35 S RNA exclusively from the nucleolar fractions (96).Furthermore, it is possible that some of the 28 S RNA found in the nucleoplasm comes from the particulate rim of the nucleolus, which is frequently lost during isolation procedures

FIQ.5. Fate of 45 S RNA after actinomycin treatment. HeLa cells labeled for 30 minutes with C"-uridine (upper left) and further incubated for 20, 60, or 240 minutes in the presence of 5 pg/ml actinomycin D. RNA extracted as in Fig. 4 and analyzed by sucrose gradient sedimentation. Solid curves: absorbance at 260 mp; dashed curves: radioactivity. [From Schemer, Latham, and Darnel1 ('79).I [Birnstiel (26)1. Thus one might suspect that the transition to 28 S occurs in the nucleolus and that release into the nucleoplasm is concomitant with, or immediately follows, this conversion. The molecular event associated with this transition in sedimentation behavior is still unknown. It could conceivably be solely a configurational change, but perhaps more likely it involves a splitting off of a couple of small RNA fragments, about 100 nucleotides long, that remain associated with the large ribosome subunit [ (74,111,112), Brown (26)1. If the conclusions discussed in the preceding paragraphs are correct and the production of rRNA entails the splitting of, say, two out of 7000 diester bonds in the 45 S molecule, one must seriously consider what types of mechanism could possess the necessary specificity to mediate

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such a cleavage. Since all the methylation of rRNA appears to take place on the 45 S molecule shortly after its synthesis (85, loo), it is considered unlikely that methylation could be directly responsible for the cleavage, especially for the 35 S + 28 S transformation. However, methylation could be indirectly implicated by creating specific sites for a couple of nuclease-catalyzed hydrolyses [Comb (25)1. Such sites might be similar to the nuclease-sensitive site in the center of the large rRNA component (113-115). If specificity toward cleavage enzymes is an inherent property of the precursor molecules, one might interpret the effect of azaguanine substitution, and possibly also thioacetamide alkylation, as arising from an altered affinity of the 35 S component for its enzyme. Alternatively, one could derive the necessary specificity from a number of appropriate protein ligands-if indeed much of the 45 S component exists in vivo as a complex with protein (cf. Section 11,D). Presuming that a t least part of the protein is newly synthesized (86), one could then explain the blocking effect of puromycin on the conversion process4 as an effect on the synthesis of this protein (91). At the moment there is no compelling reason to choose between these alternatives or to suspect that both might not be involved to a certain extent.

6. Synthesis of Ribosomal Protein Compared to our knowledge of rRNA, much less is known concerning the synthesis and properties of the protein elements of the ribosome. In part this is because ribosomal proteins belong to a much larger and diverse class of cellular macromolecules than does the rRNA, and consequently the task of sorting and characterizing them is considerably more difficult. Also, the techniques for isolating, preserving, and analyzing ribonucleoproteins have not been developed to the extent of those used in RNA studies, so that there is a much greater possibility of working with structures that are modified from the native state in different ways by different procedures. From sedimentation-diffusion studies and analyses using gel electrophoresis, i t appears that the mature ribosome contains twenty to thirtyfive different proteins, having an average molecular weight of about 25,000 (117,118).Not more than a few of these are common to the two 'Cells given a short pulse of labeled nucleoside followed by a chase in the presence of concentrations of puromycin effectively inhibiting >95% of protein synthesis show a reduced capacity to convert labeled 45 S RNA to 28 and 18 S components (91, 109). Yet in chases carried out in the presence of puromycin and actinomycin, the extent of conversion is identical t o that found with actinomycin alone (67').Therefore, it seems that the maturation of pulse labeled 45 S RNA towards actinomycin insensitivity is accompanied by a concomitant maturation towarda puromycin insensitivity [cf. footnote 2 and (116)l.

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subunits. In gross features, the ribosomal proteins from higher organisms seem sufficiently similar to those of bacteria (119), so that initially, a t least, one is encouraged to extrapolate from one system to the other when attempting to elucidate the mechanism of ribosomal protein synthesis. We can suppose that the structural proteins of the ribosome are those tightly bound to the rRNA and not readily dissociable in solutions of relatively high ionic strength (0.5 M salt) (119-121). These proteins appear to be linked to a particular rRNA component so as to form a “ribonucleoprotein strand,” which can apparently fold in discrete stages into a ribosome subunit by means of Mg2+-dependentprotein-protein interactions (121, 122). At least 30% of the protein can be dissociated from the ribosome by treatment with concentrated CsCl and then reassociated nmenzymatiwlly with a precision sufficient to restore the protein synthesizing capacity of the particle (123-125). The questions concerning us here are: Where is this protein synthesized, and where and how is it assembled into the ribosomal structure? One could imagine that the sites of synthesis and assembly might be different. The ribosomal protein could be synthesized in the cytoplasm and assembled together with the rRNA in the nucleolus. Conversely, both synthesis and assembly could occur in the nucleolus or in the cytoplasm. Let us consider some of the available data that might be useful in formulating a hypothesis. Among the more detailed studies of the sites of synthesis of ribosomal protein have been those by Birnstiel, Flamm, and collaborators (126128) on pea and tobacco cells. These studies featured a mechanical separation of nuclei, nuclear subfractions, and cytoplasm, followed by differential extraction of several distinct protein fractions. Ribonucleoprotein particles with an RNA/protein ratio of 0.5 and of sizes corresponding to monoribosomes and their subunits were obtained from nucleolar fractions considered to be free of cytoplasmic contamination (126). The protein of these particles comprises less than one quarter of the total nucleolar protein; and is not labeled when cells are given short pulses of C14 amino acids. This is in contrast to the rapid labeling of protein associated with the cytoplasmic ribosomes and of a residual nucleolar fraction not solubilized by deoxycholate or 1-2 M NaCl (127, 128). The amino acid composition of the protein from the “residue” ‘The over-all ratio of protein t o RNA in these nucleoli is about 8 to 1. The high proportion of protein in nucleoli has also been noted by Vincent in studies of nucleoli isolated from starfish oocytes (189). It implies that in such cells a considerable portion of nucleolar protein is probably not complexed with RNA, certainly not in proportions characteristic of ribosomes. The bulk of this protein is nonhistonelike and extractable at high ionic strength or with deoxycholate (1%). Its abundance compared to ribosomal protein may well depend on the physiological state of the cell.

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fraction strongly resemblcs that of the cytoplasmic ribosomes (128). From these and other results, it was suggested that ribosomal protein is produced according to the following sequence : protein of “residue” fraction + protein of nuclear ribonucleoprotein particles + protein of cytoplasmic ribosonies. However, because of thc rclatively great amount of protein synthesis occurring simultaneously in the cytoplasm, it is difficult to exclude the possibility that the protein is originally synthesized in the cytoplasm and rapidly transferred to the nucleolus. Efforts to resolve this problem by autoradiographic means have also been inconclusive, essentially for the same reason. I n virtually all cell types studied by this method, there is no preferential labeling of nucleolar protein, even after short pulses of radioactive amino acids (49, 130, 131). After administration of actinomycin D in doses that selectively block ribosomal RNA synthesis, some depression of amino acid incorporation into nucleolar protein was noted (132, 133). However, this was considerably less than the inhibition of nucleoside incorporation into nucleolar RNA, and furthermore it was observed when a sufficiently long time had elapsed after actinomycin treatment so that there was a measurable decrease in nucleolar volume. Thus these results could equally well be explained if the nucleolus were an assembly site, but not a synthetic site, for ribosomal protein. One might pose the question in a slightly different way and ask whether nucleoli are indeed capable of protein synthesis. Beginning with the work of Allfrey, Mirsky, and co-workers [see (194) for refs.] there have been numerous reports of a protein synthetic capacity of isolated nuclei that does not appear to be attributable to a contamination with remnants of cytoplasm (135). In some cases, this activity could be further ascribed to ribosomal particles isolated from nuclei (136, 197). There is some indication that these ribosomes can utilize DNA as well as RNA as a direct template for protein synthesis (136, I % ) , but as yet there is no evidence implicating them in the synthesis of ribosomal protein. Isolated nucleoli supplied with appropriate supplements are also capable of supporting polypeptide synthesis (72, 139), but no one has yet succeeded in localizing this activity in a particulate fraction. Indeed, in the case of plant material, those particles th a t can be liberated from nucleoli appear to be relatively inactive in protein synthesis (127). Therefore it is reasonable to suppose that, if the ribosomal protein is actually synthesized in the nucleolus, the mechanism may be entirely different from that operative in the synthesis of cytoplasmic protein. Recently a search for such an unconventional mechanism has been stimulated by the finding that certain highly purified nuclei contain little, if any, 18 S RNA and thus presumably very few small ribosome subunits (cf. Section II1,A).

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In the same sense, it is also worth considering whether cells must synthesize a specific RNA template for each of the ribosomal proteins. This would not be necessary if rRNA itself could act as the template, as proposed on theoretical grounds (139~) and suggested by recent experiments with bacterial mutants that accumulate protein-deficient particles during methionine starvation (140). However, on the basis of a triplet code, an rRNA molecule would only be capable of coding for approximately one ninth of its weight in protein. Unless one postulates considerable duplication of proteins, which does not seem to be the case on the basis of the gel electrophoresis data, one must assume either that the rRNA specifies only a relatively small part of its protein, or that the rRNA molecules are not all identical and that there is an exchange of the different protein species produced by, say, nine different rRNA templates. This consideration becomes critical in the case of species carrying few rRNA cistrons or where a high redundancy of rRNA cistrons is indicated, as in Drosophila (cf. Section

I,B) -

Another consideration of some relevance here is whether cells contain any preformed pools of ribosomal protein. Recently reported results suggest that in cultured mammalian cells (118) and yeast [Vincent (,%)I there is a pool of preformed protein capable of combining with newly synthesized rRNA and consequently responsible for the fact that inhibitors of protein synthesis such as cycloheximide and, under appropriate conditions, puromycin (cf. footnote 4) do not inhibit the effective chase of 45 S RNA radioactivity into the 18 and 28 S RNA of cytoplasmic particles. In mammalian cells, the pool is believed to comprise on the order of 5-10% of the total ribosomal protein (118). In summary, most of the data discussed in this section are consistent with the idea that a t least part of the ribosomal protein is synthesized prior to its association with rRNA in the nucleolus. Whether the protein is synthesized in the cytoplasm and rapidly transported to the nucleolus, or in the nucleolus by some rather unique mechanism remains to be determined. Consideration of the nucleolus as the site of ribosome assembly is consistent with the observation that 18 and 28 S RNA are observable in the cytoplasm only as ribonucleoprotein complexes having properties of ribosome subunits, and not as free RNA or as complexes having a reduced proportion of protein to RNA (see Section IV,A).

C. Relation to Nucleolar Ultrastructure There is some evidence from fine-structure analyses consistent with the idea that the assembly of subunit particles occurs in the nucleolus. The basic elements common to most nucleoli are RNA-containing fibrils

T H E NUCLEOLUS AND T H E SYNTHESIS O F RIBOSOMES

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about 50 A in diameter, RNA-containing granules about 150 A in diameter, and nucleolus-associated chromatin. It was observed with electron microscopic autoradiography that there is a rapid labeling of RNA in the fibrous portion followed by a slower labeling of the granules (141, 142). The time necessary for appearance of label in the nucleolar granules can be correlated with that observed for the transition from 45 S RNA to smaller components. Since the nucleolar granules have been shown to resemble closely the cytoplasmic ribosomes with regard to cytochemical properties and sensitivity to enzymatic digestions (14% 146), one might suppose that these observations represent a visualization of particle formation. I n pea nucleoli, where the 60 S ribosomal subunits are the major particulate constituent, the granules are seen to be smaller than the cytoplasmic ribosomes (146).Several recent determinations of the RNA components present in nucleoli (Section II1,A) and studies of release of particles into the cytoplasm (Section IV,A) suggest that this may be revealed in many other cell systems when sufficiently detailed observations are made. Furthermore, when ribosome synthesis is not in evidence, such as in early stages of embryogenesis (ldS),or in mitotic division stages of the cell cycle (147),there is a noticeable absence of the nucleolar granules. Suppression of RNA synthesis with actinomycin D also leads to a rapid disappearance of granules from the nucleolar periphery (148) or to their marked segregation from the fibrous elements (146). Thus, on the basis of morphology, it seems plausible that ribosome assembly takes place in the nucleolus. However i t is always difficult to obtain unequivocal proof from this type of data. Investigation by electron microscopic autoradiography of the sites of nucleolar protein synthesis may be instructive in this regard. Also it should be pointed out that passage of ribosomelike particles through the nuclear pore has been looked for and not observed (146).Although this is a t first glance rather disturbing, it may turn out to have a rather straightforward kinetic explanation (Section IV,A) .

IV. Appearance of Ribosomes in the Cytoplasm A. Subunits Our knowledge concerning the mode of entry of the ribosomal particles into the cytoplasm has come mainly from kinetic studies of the cytoplasmic rRNA components (10,80, 88) or of ribonucleoprotein particles containing these components (10, 110,149,150). When pulse-chase experiments of the type discussed in Section II1,A are performed and the cytoplasmic RNA is isolated by cellular fractionation or differential-

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phenol extraction, a distinctive labeling pattern of the rRNA components is observed (10, 80, 88, 160). The label appears first in the 18 S component and then, after a significant lag, in the 28 S component (Fig. 6 ) . 6 Analysis of the ribonucleoprotein particulates of the cytoplasm by gradient centrifugation techniques indicates that the 18 and 28 S components emerge as 40 and 60 S particles (10, 110, 1.69). An example of such an analysis is illustrated in Fig. 7. When selected portions of these gradients are used as sources for the extraction of RNA, the 40 S fractions yield 18 S RNA (10,110) and the 60 S fractions, if sufficiently pure, 28 S RNA (90,149). No significant amount of newly formed rRNA is found in the lighter regions of the gradient, as could be expected if it existed as free RNA or in complexes with small amounts of protein. Since it has been shown that the subunits derived from monoribosomes by magnesium withdrawal sediment under these conditions as 40 and 60 S particles (110) and possess identical buoyant densities (cf. Section IV,B), we can conclude that the observed sequential labeling of the 40 and 60 S particles does in fact represent the emergence of ribosomal subunits into the cytoplasm. I n the studies with cultured mammalian cells, the time necessary for appearance of label in the cytoplasmic rRNA components is found to be in excellent agreement with events in the nucleus. Penman et al. (86) found that there is a concurrent appearance of labeled 18 S RNA in the cytoplasm and labeled 28-35 S RNA in the nucleus, both apparently derived from the breakdown of the 45 S component. Subsequently, as the labeled 28 S RNA appears in the cytoplasm, there is a corresponding loss of radioactive 28 S component from the nucleus. The differential rate of release into the cytoplasm of the two newly formed subunits off ers a reasonable explanation for the preponderance of large subunits in nucleoli. It has been suggested that the large subunit, perhaps because of its greater complexity, takes longer to complete than the small subunit and therefore is retained within the nucleolus for a longer time (88). One could imagine, then, that with cells in a steady state there are equal numbers of large and small subunits leaving the nucleus, but that at any instant the small components entering the cytoplasm are relatively newer if considered from the time of transcription ‘In experiments such aa that illustrated in Fig. 6, in which “cytoplasmic rRNA” is taken to be the >4 S RNA that is extractable from whole cells with phenol and without detergent at 5”C, there is, at the beginning of the chase, a noticeable displacement of radioactivity toward the heavy side of the 28 S peak (10, 78). However, this early-labeled heavy component is, not detected in experiments where the RNA is extracted from cytoplasmic particulate fractions (881, and thus may actually represent a small portion of nuclear RNA extractable by phenol.

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FIQ.6. Series of sedimentation dirtgrams of RNA from L cells labeled for 10 minutes with H3-cytidine and then further incubated for various times with excess unlabeled cytidine in the presence of 3.3 x I&*M actinomycin D. RNA extracted with phenol at 5°C. Centrifugation for 3 hours at 39,000 rpm (SW 39) in 5 to 20% (W/V) sucrose gradients. Solid curves: absorbance a t 260 mp, the three peaks corresponding to 28, 18, and 4 S; dotted curves: radioactivity in counts per minute. Arrows designate the position of the heaviest radioactive peak. [From Perry ( l o ) . ]

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F I ~7.. Zonal sedimentation diagrams of cytoplasmic ribonucleoprotein particles. L cells (a) pulsed for 30 minutes with H3-cytidine or (b) labeled for 30 minutes and subsequently chased for 2 hours with excess unlabeled cytidine in the presence of 0.4 pg/ml actinomycin D. After swelling in hypotonic buffer, cells were homogenized (Dounce) and centrifuged a t 16,000 x g. The supernatant was layered onto a 1530% (W/W) sucrose gradient and centrifuged for 12 hours at 25,000 rpm (SW 25). Solid curves: absorbance a t 260 v ; dashed curves: radioactivity. The encircled symbols, M, 60,40, T, etc., refer to monoribosome, 60 S, 40 S, and “trough” fractions, pooled for further analysis. [From Perry (90).I

THE NUCLEOLUS AND THE SYNTHESIS OF RIBOSOMES

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of the rRNA. This idea was confirmed by the demonstration that the earliest labeled monoribosomes consist of a labeled small subunit attached to an unlabeled large subunit (90) (cf. Section IV,C).’ There is some indication that, in certain cell types, the exit time of the particles may be extremely fast coiiipared to the time of their synthesis and release. This stems from the observations that little or no 18 S RNA component is present in the nuclei of these cells (cf. Section III,A), a situation that would not obtain if the transport time were of the order of several minutes. The rapid transport might explain why the particulate rim of nucleoli appears to be a rather sharply delineated structure rather than a “diffusion source.” Furthermore, if the passage through the nuclear membrane were also rapid, one could understand the frequent inability to observe ribosome-like particles within the membrane. On the other hand, the possibility of a transient alteration of the structure during passage through the membrane still has to be considered (26, 145), especially in view of the distinctive properties of the new particles discussed below.

B. Distinctive Properties of New Particles From sedimentation studies such as those illustrated in Fig. 7, no difference between newly formed and “mature” subunits is evident; i.e., the positions of the absorbance and radioactivity peaks are essentially coincident. However, if the particles are purified by sedimentation in sucrose gradients and subsequently banded in cesium chloride gradients, the new particles are seen to have a distinctive buoyant density (151). This property is manifested by both large and small subunits (Fig. 8 ) . Since the particles have almost identical sedimentation behavior before and after being banded on CsCl (Fig. 9 ) , we may presume that the differences in buoyant density are not procedural artifacts. The metabolic lifetime of the new particles is relatively short. Kinetic experiments (151) demonstrate that they are converted to a form resembling that of the bulk of the subunit particles (Fig. 10) within 0.1 of a cell generation time. It was also observed that the buoyant densities of the “mature” 40 and 60 S particles are indistinguishable from those of the corresponding subunits derived by dissociation of monoribosomes. I n recent experiments, it has been found that the buoyant density of new particles can be made to coincide with that of the “mature” particles ‘In the steady state, the cytoplasm should contain a surplus of small subunits equal in number to the excess of large subunits found in the nucleolus. However, since this amounts to not more than a few per cent of the cytoplasmic ribosomes, it is not readily detectable.

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THE NUCLEOLUS AND THE SYNTHESIS OF RIBOSOMES

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by an appropriate incubation with trypsin (152). Since the RNA components of new and old particles are identical, one interpretation of these experiments is that the new particles have an extra protein component responsible for their lower density characteristic. However, the amount of protein [buoyant density N 1.25 (SS)] necessary to produce the observed density displacement would have a molecular weight in excess of 1.4 X lo5, and it is difficult to understand why such a great amount of protein would not cause the new particles to sediment faster than the “mature” ones in the sucrose gradients. One is therefore led to suspect either that the extra component associated with the new particles is a very low density material, such as the lipoprotein present in endoplasmic membranes [buoyant density < 1.01 ( 1 6 3 ) ] , in which case it need be only about 5 X lo4 MW, or, alternatively, that i t is a protein that induces an unfolding or opening up of the particle structure, fortuitously offsetting the effect of increased inass on the sedimentation velocity.

C. Involvement with Polyribosomes After their appearance in the cytoplasm as 40 and 60 S particles, the newly synthesized subunits appear next in the polyribosomes and later as part of the monoribosome pool (110, 149). It appears that the combination of subunits to form the complete ribosomal particle occurs during, rather than before, the fabrication of the polyribosome structure ( l @ ) ,and that the release of ribosomes from the polysome structure can take place without dissociation of the subunits (10, 149). Indeed, the early labeled monoribosomes, when dissociated in the presence of EDTA, are found to have a much greater proportion of label in their small subunit (Fig. l l ) , as might be predicted on the basis of the difference in times of entry of the two subunits into the cytoplasms. It is Fro. 8. CsCl density gradient centrifugation of ribonucleoprotein particles. Cytoplasmic particles from L cells labeled for 45 minutes with Ha-uridine were fractionated by zonal centrifugation as illustrated in Fig. 7. To achieve greater purity, the fractions corresponding to the 60 and 40 S peaks were rerun on a second 15-30% sucrose gradient (insets). The 60 and 40 S peaks (shown by horizontal bars) were fixed in 670 HCHO for 24 hours, dialyzed, and then banded in density gradients of CsCl together with a marker of fixed monoribosomes labeled with p92 at high specific activity ( p = 1.55). Solid curves: 260 mp absorbanre of 60 and 40 S particles (not drawn in insets) ; dashed curves: H3 radioactivity; dotted curves: P“ marker. Note that the major radioactivity peaks characterizing the newly formed 60 and 40 S particles are shifted toward lower buoyant density from the major absorbance peaks representing the “mature” particles by 0.02 and 0.04 gm cm4, respectively. The lower-density peaks at p = 1.43 and 1.38 correspond to aggregated material. For further experimental details see (161).

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FIG.9. (a) CsCl density gradient profiles of two preparations of 40 S particles: cells labeled for 30 minutes with H3-uridine; 0 , from cells labeled for 30 minutes with H3-uridine and then incubated for 4 hours with excess unlabeled uridine. Dashed curves: H3 radioactivity; solid curve: absorbance a t 260 mp of particles from the first experiment (that from the second experiment is virtually the same and is not shown). Dotted curve: Psz-labeled monoribosome marker ( p = 1.55). The bulk of the fractions shown by the horizontal bars A, B, and C were separately pooled, dialyzed, and resubmitted to zonal centrifugation on 1530% (W/W) sucrose gradients together with a marker of P=-labeled, fixed 40 S particles. (b) Sedimentaiton on sucrose gradients [3 hours, 39,000 rpm (SW 39)l of preparations A, B, C, and of samples of the original preparations of fixed 40 S particles used for the CsCl banding experiments. Note that all preparations sediment essentially the same as the Psz-labeled 40 S marker (vertical line), which was included in each gradient. This proves that the lower buoyant density characteristic of the newly formed 40 S particle is not a procedural artifact.

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not yet certain whether the elements responsible for the lower buoyant densities of the new subunits are lost before, or during the process of, subunit association. It is equally unclear to what extent monoribosomes recycle through the polyribosomes in competition with newly formed particles, and to what extent they equilibrate with the pools of “mature” 40 to 60 S structures.

247

THE NUCLEOLUS AND THE SYNTHESIS OF RIBOSOMES

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I

I

I

I

I

I

I

I

I

I

12

16

20

24

28

4

8

12

I6

20

24

FRACTION

NUMBER

I

28

FRACTION NUMBER

FIQ.10. CsCl density gradient profiles of 40 S particles from experiment shown in Fig. 7. Solid curves: absorbance at 260 mp; dashed curves: radioactivity. Note that the prominent radioactive peak at p = 1.43 decreases almost threefold concomitant with the appearance of radioactivity in the p = 1.49 band, indicating that the metabolic half-life of the low-density 40 S particle is less than 2 hours. [From Perry and Kelley (161).1

essentially the same rate as the 40 S particles led to the proposal that the small subunit forms a complex with the RNA message and thus serves as a vehicle for its transport from nucleus to cytoplasm. However, in a variety of systems, efforts to demonstrate conclusively the existence of such messenger-subunit complexes have been unsuccessful (161,168, 169). Buoyant density studies such as those illustrated in Figs. 8-10 do not reveal any significant amount of radioactive material banding a t higher densities than the subunit, as would be expected of such complexes. Rather, a significant portion of the messenger RNA

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- - - - - __ -0

a I

a I

9

a

I

u?

0

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bands with material of rather low buoyant density. Hence, the cytoplasmic messenger RNA that is not in polyribosomes behaves as if it were complexed with protein, but not attached to subunits (151, 158). This leads one to suspect that its sedimentation in the region occupied by the small subunit may be fortuitous. Furthermore, there is some evidence suggesting that polyribosomes formed from new message may originate on the nuclear envelope (16U) or on specific elements of its associated membrane system (155). If this be the case, then one could envision that polysome formation involves an interaction of even greater complexity with structural elements of the cell.

D. Turnover of Ribosomes It seems quite clear that in rapidly growing cells of higher organisms (78), as in exponentially growing bacteria (161, 16,9),the exigence of producing a large amount of ribosomal RNA in a limited period of time does not allow the organism to engage in an appreciable amount of ribosome turnover. On the other hand, a rapid turnover of ribosomal RNA has been observed in liver cells of the rat (163, 164). In this system, labeled ribosomal RNA is found to decay exponentially with a half-life of from 2 to 5 days, considerably less than .the half-life of the average liver parenchymal cell. The RNA of both ribosomal subunits decays a t the same rate (164). This is a necessity for maintenance of a steady state if, as discussed previously (Section IV,A), the number of large and small subunits entering the cytoplasm per unit time is the same. This turnover provides a rationale for the relatively intense nucleolar activity and rRNA synthesis observed in cells of liver and other less rapidly proliferating tissue. Whether the continued synthesis of new ribosomes is necessary for some regulatory mechanism in the cell or whether it is simply an aspect of the over-all rejuvenescence of cellular constituents remains to be established.

V. Synopsis Granted the license to select the mechanisms that appear most likely on the basis of the preceding discussion and, for the sake of simplicity, to

FIG.11. Series of sedimentation diagrams of 50 and 30 S particles derived from monoribosomes b y chelation with ethylenediamine tetraacetate. The monoribosomes with Ha-uridine followed b y incubation with excess unlabeled uridine and cytidine for 0.5, 1.0, 2.0, 4.0, 12, and IS hours. Note that the earliest labeled monoribosomes contain almost all of their label in the 30 S subunit and that, as chasing proceeds, the proportion of ribosomes containing labeled 50 S subunits steadily increases, [From Perry (901.1

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generalize results from various organisms, we attempt here to summarize this discussion with a “cradle-to-grave” model of the ribosome. The process may be resolved into the following phases (Fig. 12) : (1) Transcription of a set of contiguous rRNA genes a t the nucleolar organizer locus into a covalently continuous precursor molecule containing one or two pairs of rRNA components. Intercalation of 5 S cistrons between each rRNA cistron is purely hypothetical. Regulation of transcription probably entails structural modification of the genetic material. NUCLEOLUS

-v

4 SMALL SUBUNIT

TRANSCRIPTION

LARGE SUBUNIT

MONORIBOSOME

RAPID TRANSPORT

PROTEINCONTAINING

-/

BREAKDOWN \’ \‘

/ \

-

POLYRIBOSOME

FIQ.12. “Cradle-to-grave” model of the ribosome (see Synopsis, Section V).

Methylation of the rRNA and possibly some association with protein seem to occur a t this level. (2) Specific cleavage of the precursor into 18 and 32-35 S pieces and assembly of RNA with preformed protein to form ribosome subunits. Smaller subunits are completed and released from the nucleolus earlier than the larger subunits, the RNA of which is further reduced to 28 S, possibly by removable of two 5 S fragments. (3) Rapid transport of the subunits to the cytoplasm where they appear associated with a protein-containing material that causes them to possess a lower buoyant density than “mature” subunits. Combination of the individual subunits with messenger RNA to form polvribosomes.

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(4) Release of the monoribosome from the messenger RNA and either recycling, dissociation to subunits, or degradation to small molecules. ADDENDUM

On the precursors of rRNA. Separation of the 45 S ribosomal precursor RNA from most of the rapidly sedimenting DNA-like RNA (cf. footnote 1 on page 229) can be achieved by chromatography on methylated albumin columns (166, 166). The rRNA precursor, designated ql, elutes from the column a t a lower salt concentration, and is shown by hybridization tests to contain base sequences homologous to both rRNA components (167). Since in mammalian systems the two ribosomal components have a n appreciably different base composition (see Georgiev, this volume), one might expect that if there were two types of 45 S precursor of equal molecular weight, one for each rRNA component, these should be resolvable on the methylated albumin column. From currently published profiles (168, 169), it appears that ql is not a complex peak. If substantiated by a refined hybridization analysis on individual parts of the peak, this finding could be used as further support for the hypothesis of a single 45 S precursor to both rRNA components. A preliminary estimate from sedimentation equilibrium data indicates that the molecular weight of the 45 S component is about 4.5 million (92), a value close to that predicted on theoretical grounds (cf. Section 11,C). Similar measurements on the 35 S molecule suggest th a t its molecular weight is about 30% greater than the 28 S ribosomal component (92). This seems to eliminate the possibility that the 35 and 28 S components are configurational variants of each other, and suggests further that they differ by more than two 5 S pieces (equivalent to only 4% of the 28 S molecular weight). Besides, it has been recently noted (170) that the 5 S RNA of Xenopus hybridizes with neither the proportion nor the type of DNA expected on the basis of the intercalation model in Fig. 12, thus casting further doubt on the relevance of 5 S RNA to the 3 2 3 5 S + 28 S transition. Detailed studies of the kinetics of methylation of the 45 S component in HeLa cells (171) suggest that a t least some of the methyl groups are attached before completion of the transcription process. The transcription time for one 45 S molecule is estimated to be about 2.3 minutes. By means of a controlled stepwise hydrolysis with snake venom phosphodiesterase Hadjiolov and co-workers (172) have attempted to measure the base composition of various parts of the rRNA chains. From their results i t appears that the 5’-OH ends of both rRNA components are significantly richer than the 3’-OH ends in their relative proportion

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of guanine and cytosine residues. At present the functional significance of this heterogeneity is obscure. Nevertheless, it should be possible to carry out similar analyses on purified preparations of 45 S molecules and in so doing learn more about the nature of these giant precursor molecules. O n the template activity of rRNA. The in vitro template activity of the protein-deficient “relaxed” particles has been found to reside in RNA components distinct from ribosomal RNA (173,174). Thus, the alternative (cf. Section III,B) whereby rRNA acts as the template for the synthesis of ribosomal protein is without substantial experimental confirmation. On newly formed and mature ribosomal subunits. The fact that newly synthesized subunits, as they emerge in the cytoplasm, possess more protein than the subunits that constitute the bulk of the native subribosomal particles (cf. Section IV,B) suggested that the newly formed and mature structures are not in rapid equilibrium with each other, but are separated by polyribosome and monoribosome pools as diagrammed in Fig. 12. In agreement with this idea is the finding that during the course of maturation of rabbit reticulocytes, native subunits constitute a roughly constant proportion of the total ribosomal particulates (176). Since the native subunits persist long after the production of ribosomes has ceased and, in fact, after the ribosome content per cell has undergone a three- ta fourfold reduction, it can be concluded that in this system, too, the bulk of the native subunits are not precursors to the ribosomes. Recently we have examined the sedimentation characteristics of new subunits that have had their extra protein component removed by tryptic digestion. We find that, although these particles now possess buoyant densities almost identical to similarly treated mature particles, they sediment significantly slower. We interpret this to mean that the new particles have a less compact or more asymmetrical configuration than mature ones both before and after the removal of the associated protein component. On ribosome turnover. Further studies on the turnover of ribosomes in rat liver cells indicate that the rate of turnover of the structural protein of the ribosome is indistinguishable from that of the RNA, and that during starvation, the ribosome content per cell decreases because of an increased rate of degradation and a diminished rate of synthesis (176). ACKNOWLEDGMENTS The author would like to thank Miss Dawn E. Kelley and Mrs. Blanche Buckner for their able and enthusiastic technical assistance in performing many of the

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experiments described in this article. The research was supported by grants from the National Science Foundation and the National Institutes of Health.

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The Nature and Biosynthesis of Nuclear Ribonucleic Acids G. P. GEORGIEV Institute of Molecular Biology, Academy of Sciences of USSR, Moscow, USSR

I. Introduction . . . . . . . . , . . . . . . . A. Terminology . . . . . . . . , . . . . . . B. Main Classes of Cellular RNA . . . . . . . . . . C. Fractionation of Nuclear RNA: Main Approaches . . . . 11. Phenol Fractionation of Nuclear RNA’s and Their Characteristics . A. Methods. . . . . . . . . . . . . . . . . B. Molecular Classes of Newly Formed (Rapidly Labeled) Cellular RNA. . . . . . . . . . . . . . . . . . C. Characteristics of Chromosomal D-RNA . . . . . . . D. The R-RNA of the Nucleolochromosomal Complex . . . . 111. Fractionation of Subnuclear Structures and Nuclear Ribonucleoproteins . . . . . . . . . . . . . . . . . A. Principles of Methods . . . . . , , . . . . . . B. Ribonucleoproteins of Nuclear Sap . . . . . . . . . C. RNA of the Deoxynucleoprotein Fraction (Chromosomal RNA) D. RNA of Nucleoli and Nucleonemata . . . . . . . . E. Conclusion: The Nature of the RNA of Subnuclear Structures . IV. The Biosynthesis of RNA in Nuclear Structures and Its Transport to the Cytoplasm . . . . . . . . . . . . . . . . A. Localization and Mechanism of RNA Biosynthesis . . . . B. mRNA Biosynthesis in Chromosomes: Main Stages . . . . C. Transport of mRNA . . . . . . . . . . . . . D. Synthesis and Transport of Ribosomal RNA and tRNA . . . E. Regulation of RNA Biosynthesis. . . . . . . . . . V. Conclusion . . . . . . . . . . . . . . . . . Note Added in Proof . . . . , , . , . . . . . . References . . . . . . . . . . . . . . . . .

259 260 260 261 264 264 269 277 286 288 288 291 299 300 303 304 304 308 315 322 328 340 341 342

1. Introduction Many reviews concerned with studies of nuclear RNA have been published during the last few years (1-6). This is to be expected since the cell nucleus is that cell organelle in which the formation of RNA and probably also of RNA-containing structures takes place (see Prescott in Volume 3 of this series). 259

260

G. P. GEORGIEV

This circumstance explains the large complexity of nuclear ribonucleoproteins as well as the fact that until recently the structure and function of different nuclear RNA fractions has remained obscure. This paper is concerned with the methods of nuclear RNA fractionation and with the nature of the RNA fractions detected in nuclei. It also describes the structural organization of nuclear RNA, that is, of nuclear ribonucleoproteins, the relative role of the different nuclear structures in RNA synthesis as well as some estabished mechanisms of this process, and finally the part that nuclear proteins play in the regulation of RNA biosynthesis.

A. Terminology Three classes of RNA with different roles in protein synthesis can be visualized from the standpoint of their functions. They are: ribosomal RNA (rRNA), transfer RNA (tRNA), and messenger (template) RNA (mRNA). However, it is not always easy to relegate a particular RNA to one of these classes. An example is an RNA having the base composition of rRNA but a different molecular weight. This type of RNA is probably a precursor of true rRNA. On the other hand, an RNA may have some characteristics of mRNA but not be a template in protein biosynthesis [“informational” RNA in the broad sense of the word (7)1. Consequently, it seems useful to classify RNA according to its chemical and physical properties because then one can designate RNA’s with yet unestablished functions. The main types of RNA’s according to this classification are as follows: (1) R-RNA, an RNA having the base composition of ribosomal RNA and probably containing the same nucleotide sequences; (2) D-RNA, an RNA with the base composition of cellular DNA including mRNA (informational RNA in the strict sense) as well (8) ; (3) S-RNA, low polymeric soluble RNA. Recent investigations on RNA fractionation involve analysis both of total RNA of the sample and of labeled RNA in the same preparation. Properties derived from the results of these types of analysis may differ considerably. Hence, RNA determined by its W absorbance is designated as bulk R N A or preformed (pre-existent) RNA, and RNA determined by its radioactivity is designated as newly formed RNA. This RNA may also be called rapidly hbeled R N A if the time of isotope incorporation is suflFiciently short.

B. Main Classes of Cellular RNA Ribosomal RNA is the RNA isolated from well-characterized ribosomes. But to regard an RNA as ribosomal, one should in practice demonstrate its typical base composition [ (G C)/(A U) = 1.6 in higher animals] and its typical two-component sedimentation pattern with

+

+

261

NUCLEAR RIBONUCLEIC ACIDS

peak values of 1&18 S and 28-30 S for the slower and faster components, respectively. Bot,h bulk and newly formed RNA’s can be characterized by these parameters. The main functional determinant of tRNA is its ability to form aminoacyl derivatives and to transfer labeled amino acids to ribosomes. To classify an RNA as tRNA, one should demonstrate its low molecular weight (sedimentation coefficient 4 S) and typical base composition, rich in GC nucleotides (in higher animals). A rapid turnover of CCA end groups and the presence of unusual minor nucleotides are also characteristic of tRNA. Newly formed tRNA can be tentatively characterized by two parameters: by its base composition and sedimentation coefficient. The case of mRNA is more complicated. The template function of mRNA can be strictly proved only if this RNA induces synthesis of a particular, defined protein in ribosomal particles. But such unequivocal proofs are exceedingly laborious and often impossible (9-11).Accordingly, in most cases, one must be content with more or less indirect evidence, such as DNA-like base composition [in DNA of higher animals the ratio (G C)/(A T) is 0.7-0.81, the ability to hybridize effectively with a homologous DNA, and the ability to stimulate protein synthesis in isolated ribosomes (12). The first two characteristics are structural and are indicative of the fact that a large number of cistrons participate in the synthesis of this type of RNA (this is typical for mRNA of most cells). The third, although functional, is relatively unspecific; for example, synthetic “unnatural” RNA’s may also stimulate amino acid incorporation into proteins. However, if a particular RNA possesses all these properties, it can almost certainly be regarded as informational. A number of other properties (rapid incorporation of precursors, rapid breakdown, sedimentation behavior different from that of rRNA) are unspecific and cannot serve as evidence of the informational nature of a particular RNA. There are many cases, however, when the available information is insufficient to classify an RNA unequivocally in a definite functional class. Then determination of its base composition may help to relate it to one of the broad groups, D-RNA or R-RNA. It should be noted that even this classification is not always possible. However, since in higher animals DNA and consequently D-RNA are of the AU type whereas R-RNA is of the GC type, there is no overlapping in composition between these classes (IS) and a clear distinction is possible.

+

+

C. Fractionation of Nuclear RNA: Main Approaches The presence of RNA in the cell nuclei is a well-known fact. This RNA can be detected cytochemically in the nucleolus, in chromosomes,

262

G . P. GEDRGIEV

and in nuclear sap (14-16). Thus, by its localization, nuclear RNA is heterogenous. A special interest in nuclear RNA arises from the demonstration of the important role of the nucleus in RNA biosynthesis. The incorporation of labeled precursors into nuclear RNA is much more rapid than into cytoplasmic RNA (17, 18). Thc removal of the nucleus from a cell inhibits at the same time synthesis of cytoplasmic RNA (19, 20) while isolated nuclei are capable of RNA synthesis (21, 22). This is good evidence for the nuclear localization of cellular RNA synthesis. However, the nature of nuclear RNA has been obscure for a long time. The first characterization of it was made by Elson and Chargaff (23) (Table I ) who found that it is somewhat less rich in GC nucleotide pairs than is cytoplasmic RNA. I n later work, the heterogeneity of nuclear RNA was demonstrated. By extraction with dilute salt solutions (r/2 = 0.05-0.15),nuclear ribonucleoproteins were divided into soluble and insoluble fractions differing in RNA base composition and metabolic activity (21, 24-26'). Further development of this procedure made it possible to obtain new data concerning the nature of nuclear ribonucleoproteins. Salt fractionation enabled investigators to separate the substance of the main nuclear ultrastructures: nuclear sap (extractable by low ionic strength solvents), deoxyribonucleoprotein (extractable by concentrated salt solutions), and the nucleonemata of nucleoli and chromosomes (residue) (27-29). Studies of the RNA and ribonucleoproteins of these fractions revealed some basic facts. Nuclear sap contains ribosomes and transfer RNA (3&33). I n nucleoli and chromosomes, RNA-containing granules are found, probably either ribosomes or their precursors (29, 34). And finally a modification of the extraction procedures enabled investigators to isolate ribonucleoprotein particles containing mRNA (35). All these methods have several weak points. A lengthy procedure for the isolation and extraction of nuclei risks enzymatic damage of RNA. Further, the correlation of the fractions obtained with the cytological structures in nuclei is fraught with uncertainty. These defects are avoided in two other methods of isolation of nuclear RNA's. The first of these is phenol fractionation, introduced independently by Sibatani et al. (36'58)and by Georgiev and Mantieva (39, 40). These authors have demonstrated that the phenol treatment of cells in Kirby's procedure (41) does not extract all cellular RNA and that a portion of RNA remains bound a t the water-phenol interphase. The material a t the interphase consists of cell nuclei, and the RNA nonextractable by phenol t-reatment is nuclear RNA or, more precisely, RNA of chromosomes together with RNA of nucleoli (RNA of the nucleolochromosomal apparatus). This fraction contains all the rapidly

TABLE I BASECOMPOSITION OF NUCLEAR RNA

AND

ITSFRACTIONS

Base composition (mol %) Source

Cell structure

RNA fraction

G

C

U

A

K

=

(G +C)/(A + U )

Ox liver Ox kidney Rat liver

Nuclei Nuclei Nuclei Cytoplasm

-

26.6 26.2 26.2 31.6

25.9 29.6 29.7 30.3

27.0 22.2 23.9 20.1

20.5 22.0 20.2 18.0

1.11 1.26 1.27 1.63

Ehrlich ascites cells

Cytoplasm

Total RNA-E RNA-N 10-40°C 55-60°C

31.4 26.7 23.7 30.1 32.6 24.9

28.5 26.3 23.9 28.0 29.9 20.1

20.8 25.0 27.4 22.4 18.2 27.1

19.3 22.0 25.0 19.5 19.3 27.8

1.50 1.13 0.91 1.40 1.67 0.82

Nucleolochromosomal apparatus

Rat liver

1

10-40°C 31.2 29.3 18.2 21.3 1.53 55-65°C 22.4 24.0 24.7 28.8 0.86 ,Newly Formed RNA (Base Composition Determined from Pa*Incorporation into 2'- and 3'-Nucleotides of RNA) Ehrlich ascites cells 10-40°C 30.8- 30.3 22.6 16.3 1.57 Nucleolochromosomal apparatus 55-65OC 20.7 19.7 29.2 30.3 0.68 Rat liver 10-40°C 34.8 27.6 18.2 19.4 1.65 Nucleolochromosomal apparatus 5545°C 22.4 22.8 23.2 31.6 0.83

Nucleolochromosomal apparatus

Reference

264

G. P. GEORGIEV

labeled RNA of the cell (36, 40). Methods involving treatment by hot phenol (at 6Ck65"C) (42) or phenol treatment in the presence of sodium dodecylsulfate (43, 44) for isolation of this nucleolochromosomal RNA have been described. I n the course of this type of fractionation, the cells are treated with phenol simultaneously with or even before homogenization, to inhibit the activity of cellular RNases so that RNA degradation is minimal. The RNA of nuclei so obtained has been further characterized by sedimentation (42, 44) and the first heavy precursors of ribosomal RNA were discovered (45). Subsequently, it was established that the nucleolochromosomal RNA is a mixture of D-RNA and R-RNA (4.2, 46) and that these RNA classes can be separated. The demonstration of the existence of a DNA-like RNA in animal cells was the important result of these studies. The simple and effective method of phenol extraction a t different temperatures enabled investigators to isolate separately R-RNA and D-RNA (47, 48). The study of these RNA classes sheds some light on their functions. Another approach in the fractionation of cell nuclear RNA is the attempt to isolate minimally damaged subnuclear structures, nucleoli and chromatin. The methods employed are mainly those of Vincent (49) and Dounce (60),which are based on the differential centrifugation of nuclei disrupted mechanically in media of low ionic strength (49473). They have been applied successfully in the isolation and purification of nucleoli and recent concepts of the role of the nucleolus in the synthesis of ribosomes and tRNA arise mainly from this methodological approach. The use of all these methods presently enables us to obtain information as to the properties and functions of various classes of nuclear RNA. The available literature concerning these points is reviewed below in the following order. First, the characterization of nuclear RNA, which demands minimally damaged RNA and is usually achieved by phenol fractionation, is discussed. Second, the characterization of nuclear ribonucleoproteins, for which the methods used are those of salt fractionation and the isolation of subnuclear structures from nuclear homogenates, is discussed.

II. Phenol Fractionation of Nuclear RNA's and Their Characteristics A. Methods Kirby has shown that, when a cell suspension. is treated by a mixture of phenol and water, the cell constituents are distributed between the

NUCLEAR RIBONUCLEIC ACIDS

265

phases according to their solubility, the bulk of proteins in the phenol phase and RNA and polysaccharides in the aqueous layer (41). If the cells are treated with a mixture of 0.14 M NaCl and phenol a t p H 6 practically all the DNA is present as deoxyribonucleoprotein a t the water-phenol interphase (39). The different behaviors of DNA and RNA in this procedure depend on differences in their linkages with proteins. If the salt bonds of deoxyribonucleoproteins are dissociated and the solubility of histones in phenol is increased (for example, by the addition of p-aminosalicylate, dodecylsulfate, or alkaline phenol), DNA is also deproteinized (54, 39). It is essential to note that, in the usual phenol procedure, not only DNA but also a small part of the RNA remains in the middle layer no matter how carefully it has been washed prior to the extraction ( 3 9 ) . Sibatani e t al. (36-38)have shown that about 15% of the cellular RNA remains a t the interphase after phenol treatment and, in incorporation experiments, this RNA shows a far greater specific activity than “extractable” RNA. It was therefore concluded that the phenol treatment results in fractionation of RNA, but the character of this fractionation remained obscure. On the other hand Georgiev and Mantieva (do), studying the properties of deoxyribonucleoprotein a t the interphase of phenol and 0.14 M NaCl a t pH 6, found that this material consisted mainly of cell nuclei retaining their usual morphological appearance, including nucleoli. These “phenolic nuclei” can be purified from cytoplasmic contamination by differential centrifugation in the phenol-0.14 M NaCl system. Purified phenolic nuclei contain all the cellular DNA, variable amounts of RNA, and denatured proteins. It is interesting that the DNA: RNA: protein ratio in phenolic nuclei is approximately the same as in the nucleolochromosomal complex obtained by the extraction of nuclei isolated in aqueous media. A comparison of the results obtained by the phenol fractionation method and by other methods, as well as cytological analysis, shows that, to a first approximation, the RNA of phenolic nuclei corresponds to the RNA of nucleoli and chromosomes, that is, to RNA of the nucleolochromosomal complex (40). Thus phenol fractionation permits the separation of the nucleolochromosomal RNA from other RNA classes of the cell without preliminary isolation of nuclei. It should be stressed that all the rapidly labeled cellular RNA is found in phenolic nuclei, so that phenol fractionation permits the separation of newly synthesized cellular RNA’s. For this reason, it is useful for the investigation of RNA biosynthesis in animal cells (Table 11). The phenol method of RNA fractionation has become widely used

TABLE I1 METABOLIC ACTIYITY A N D BASE COMPOSITION OF RNA FRACTIONS OBTAINED BY HOT PHENOL FRACTIONATION (74)

K Tissue and conditions of incubation

Rat liver, Pa*,30 minutes

Cytoplasm

+

Bulk RNA

RNA fraction

Mouse Ehrlich ascites cells, Cytoplasm Pa*,1 hour

Base composition = (G C)/(A U)

(10°C)

(10°C) l(t40"C

1.63 1.55 1.66 1.40 1.25 0.79 0.80 1.55 1.55 1.39

+

Newly formed RNA -

55-60°C 60-65°C

R

90

R 1.40 1.28 0.71

1.70

1.17

1.10 0.86 0.92

Type of RNA

Specific Per cent of activity total highcpm/mg polymerRNA RNA 90 5.7 1.1

R + D

D R R

1.5

1.5

1.2

} 2.8

97.5

97.5

R + D 1.14 0.84

0.28

D

O'"} 0.20

0.45

Per cent of total activity of highpolymer RNA

104

10.9 10.9

7,100 11,000 7,800 7,000 7,200 8,500

;!9 .:i] 51.7 5

91 3,100 6,200 8,300 12,600 18,700 21,100

30.1 30.1 12.6 12.6

12.2 12.2 11.8 13. 4}25. 2

12.0

'}

15. 30.2 14.4

NUCLEAR RIBONUCLEIC ACIDS

267

and the early conclusions have been confirmed (44, 5548). However, several points should be kept in mind. The applicability of the method depends on the degree to which the native state of the nuclear nucleoproteins is preserved. Autolytic processes occurring when nuclei are previously isolated in aqueous media result in partial solubilization of the nucleolochromosomal RNA during phenol fractionation. Accordingly, the phenol method should be applied to whole cells or a t least to a rapidly prepared homogenate. It should be noted that the cytoplasm also contains an RNA that is not extracted by phenol deproteinization (37, 59). The metabolic properties of this RNA differ from those of the extractable RNA of the cytoplasm, but its nature is obscure. For this reason, if absolutely pure nucleolochromosomal RNA is desired, phenolic nuclei must be separated from cytoplasmic contamination (40). However, in most cases, unpurified phenolic nuclei (that is, from the water/phenol interphase layer) are quite adequate. There are two methods available for the isolation of free nucleolochromosomal RNA. I n the first, phenolic nuclei are treated by a hot (60-65°C) phenol-NaC1 (0.14 M ) mixture a t pH 6 for 15-20 minutes. This results in a relatively complete (about 80-90%) extraction of nucleolochromosomal RNA (42). Another method is based on the observation by Sibatani et al. (36) that phenol extraction in the presence of 0.5 to 1% sodium dodecylsulfate (SDS) liberates all cellular RNA. However, two precautions are necessary to prevent the extraction of DNA. These are, a lowering of the p H of the extracting solutions to 5 to 5.1, and a short heat treatment of the phenol-SDS mixture a t 60°C (43, 44). SDS addition effectively inhibits ribonuclease. However, we have observed that, if RNA is obtained by treatment with phenol and 0.14 M NaCl a t pH 6 and 60°C and only then is SDS added to the RNA-containing extract, the preparations of RNA are not inferior to those prepared by immediate phenol-SDS treatment. These two methods with small modifications have been used by many other authors. Quite recently (&?a), another method for the RNA isolation from the interphase has been suggested in which the lipids are first extracted by ethanol and ether after which RNA is liberated by treatment with EDTA and phenol in the presence of bentonite. The nucleolochromosomal RNA appears in the ultracentrifuge as two peaks with sedimentation coefficients close to those of ribosomal RNA, 18 and 28 S, and contains a small amount of heterogenous material (42). However, its base composition differs from that of ribosomal

268

G. P. GEORGIEV

RNA, being intermediate between that of ribosomal RNA and DNA (Table I). Experiments on the fractionation of the RNA of phenolic nuclei confirm that it contains two subfractions, ribosomal RNA and a DNA-like RNA rich in adenine and uracil. If phenolic nuclei prepared from Ehrlich ascites carcinoma cells are treated with a mixture of 5% p-aminosalicylate and phenol, all the DNA as well as a portion of the nucleolochromosomal RNA (RNA-E) goes into solution. Another portion of the RNA remains in the nuclei and can be extracted by phenol treatment a t 65°C (RNA-N). It was shown that RNA-N resembles ribosomal RNA in base composition, whereas RNA-E has a base composition of the AU type (46). I n this respect, it resembles DNA and may be designated as D-RNA (Table I). The presence of AU-rich RNA in nuclei was shown subsequently (60). It is present in the interphase after phenol treatment of the "nucleolar" fraction. This RNA has a DNA-like composition and can be extracted by water after removal of lipids. Unfortunately, this method yields seriously degraded D-RNA preparations with S values of 2-3 (61, 68).

An improved method of D-RNA isolation is the "hot phenol fractionation" technique. I n this procedure, nuclei are treated sequentially with a phenol-0.14 M NaCl mixture (pH 6) a t different temperatures; the RNA liberated after each such treatment is collected separately. At temperatures lower than 40-50°C, mainly ribosomal RNA is liberated, at 5&55"C a mixture of R-RNA and D-RNA goes into solution, and a t 6045°C relatively pure D-RNA is extracted (Table I ) . Usually treatment a t 40-45OC, 55"C, and 63°C is employed. The interphase after the final extraction contains a small amount of RNA (10-15% of the amount in phenolic nuclei). This RNA may be extracted by treatment a t 70-85OC in the presence of dodecylsulfate and in some cases it appeared to be similar to the RNA of the 60-65OC fraction (D-RNA) with respect to base composition, hybridiaability, and sedimentation properties. The method described has been applied successfully to Ehrlich ascites cells, to rat liver (47, 48), and, somewhat later, to avian liver and reticulocytes, to spleen, hypophysis, and thymus (9, 61, 63-6'6~). As an example, for Ehrlich cells, 2-3 mg of D-RNA and 4-6 mg of R-RNA can be obtained per 100 mg of DNA. It should be stressed again that good and reproducible results with this method as well as with phenol fractionation in general can be obtained only if the starting material is absolutely fresh. The method described enables one to obtain either completely undegraded preparations or preparations damaged to a minimal extent. Ultracentrifugation in sucrose gradients reveals peaks of newly formed RNA

NUCLEAR RIBONUCLEIC ACIDS

269

similar to those observed in preparations of total cellular RNA isolated using all possible precautions against degradation. On the other hand, these preparations, including D-RNA obtained a t 60°C,appear “native” in the viscosimetric test ( 6 7 ) , giving a definite rise in viscosity upon heating. This is good evidence for the absence of intramolecular breaks in RNA molecules (68). Another method includes phenol fractionation of RNA of phenolic nuclei a t different pH values. R-RNA goes to the aqueous phase a t pH 7.6 while a fraction rich in D-RNA is liberated a t p H 8.3 (69). D-RNA can be separated from other cellular RNA’s on columns of methylated albumin and kiesehlguhr (MAK columns). Perhaps because of its secondary structure, D-RNA is bound very strongly to the column and can be eluted either by increase of temperature or by addition of alkali to the eluent (70).However, the high p H is unlikely to be without harmful effects on the integrity of D-RNA. Thus the phenol fractionation method has helped to discover the existence of D-RNA and R-RNA in the nucleolochromosomal complex and has opened ways to their further investigation.

B. Molecular Classes of Newly Formed (Rapidly Labeled] Cellular RNA The combined use of the different techniques of phenol fractionation and of sedimentation analysis of isolated RNA fractions permits a considerable fractionation of the main components of newly formed cellular RNA. It has been mentioned already that phenol extraction of cells in the presence of dodecylsulfate or a t elevated temperatures (60-65°C) permits one to obtain almost quantitatively all cellular RNA, including the nucleolochromosomal RNA. The presence of dodecylsulfate and other inhibitors of RNase facilitates the isolation of RNA in the native state. Preparations of total cellular RNA have been studied, after pulselabeling cells with radioactive RNA precursors, by means of ultracentrifugation in sucrose gradients (@, 44). Such newly formed, pulselabeled RNA has sedimentation properties differing from those of the pre-existing cellular RNA. The latter consists of three components: the ribosomal RNA peaks (rRNA,, and rRNAB) with sedimentation coefficients 28-30 S and 16-18 S, respectively, and an S-RNA peak (4 S component). On the other hand, the newly formed RNA is concentrated in the “heavier” region. Most authors have observed here two peaks with sedimentation coefficients of 45-50 S and 35-40 S, respectively (Fig. 1). A variable amount of RNA was found in the more slowly sedimenting region (25-30 S). After longer incubation times, the relative

270

G . P. GEORGIEV

I

Bottom

l o

Tube number

(b'

Top Bottom

Tube number

TOP

-

OD260

Fro. 1. Sedimentation profiles of total HeLa RNA after labeling with uridine-C" for: (a) 5 minutes, (b) 30 minutes, (c) 1 hour, (d) 4 hours, (e) 24 hours.

NUCLEAR RIBONUCLEIC ACIDS

271

proportions of these peaks decrease and the label acrumulates in the rRNA and tRNA regions. The details of this distribution pattern vary somewhat according to the source and method of isolation of RNA, but the general dynamics of radioactivity “flow” are similar ( 7 l , 7 2 ) . As has already hcen mentioned, all the rapidly labeled RNA is localized in the nucleolochromosomal complex. Heavy (35-45 S) RNA may be extracted both from “phenolic nuclei” and from nuclei isolated in a sucrose medium. On the other hand, RNA isolated by treatment with phenol in the cold or RNA isolatcd from the cytoplasmic fraction after short pulses consists of newly formed RNA only in the 4 S region. When the pulse is prolonged, two peaks of labeled rRNA appear (44). The basc composition of the rapidly labeled cellular RNA is intermediate between that of DNA and rRNA but is usually closer to that of rRNA (43). Therefore all rapidly labeled nonribosomal RNA of the cell cannot be recognized as informational. It has been suggested that, just as total nucleolochromosomal RNA is a mixture of R-RNA and D-RNA, newly formed cellular RNA in the nucleolochromosomal apparatus is likewise a mixture. The experiments using the hot phenol method confirm this suggestion. It follows unequivocally from Tables I and I1 that the rapidly labeled cellular RNA, synthesized in the nucleolochromosoma1 apparatus, is a mixture of D-RNA and R-RNA molecules (7375). Such a separation of RNA components helps to define more accurately their sedimentation properties and their nature since experiments with total cellular RNA, and even with total nuclear RNA, do not resolve peaks sufficiently for separate analysis. The results of experiments on Ehrlich ascites cells using the hot phenol fract,ionation with subsequent gradient ultracentrifugation and analysis of the fractions obtained gave the results shown in Fig. 2 (75, 7 6 ) . The total nucleolochromosomal RNA of Ehrlich cells shows the 18 and 28 S peaks of the preformed RNA and three peaks of newly formed RNA with sedimentation coefficients about 45, 35-40, and 25-30 S. Hot phenol fractionation of phenol nuclei separates these components. I n a fraction extracted a t 4OoC (between 10 and 4OOC) containing R-RNA only (both newly formed and preformed), only one peak of newly formed RNA is found ( 3 5 4 0 S component). Its base composition corresponds exactly to that of rRNA so that this component is designated as R-RNA,. The preformed RNA of this fraction is found mainly in the 28 S zone and, t o a lesser extent, in the 18 S zone. A small UV-absorbing peak is visible also in the radioactivity region ( 3 5 4 0 S) . A fraction solubilized a t 65OC (between 55 and 65OC) and containing only D-RNA (both newly formed and preformed) contains heterogeneous labeled material with a maximum of label a t 25-30 S. Some of the radio-

(C)

(01

5.3

I

30

I

20

I

10

Total

40

I

I

(f)

a

,X'

10

,x/a

20

30

\

40

'\,

55-65O

30 40

10-40°

GtC/AtU=O.81

10 20 G + C / A t U = 1.40

0.15

I

V

I a

I

(i)

I

-1

fX'

a'

1

1

10

10

20

20 x"-a

I-. 1

301 40

30

GtC/A+U= 1.87

AX

0.2

0.4

10-40"

I

1

Y

C

NUCLEAR RIBONUCLEIC ACIDS

273

activity sediments even faster (40-50 S). On the other hand, the main part of the preformed D-RNA has the sedimentation coefficient of rRNAB ( ~ 1 S) 8 ; this is why the sedimentation pattern of bulk nucleolochromosomal RNA is similar to that of rRNA (4.2).Thus newly formed and preformed D-RNA also differ in sedimentation characteristics. The RNA fraction extracted a t 55°C (between 40 and 55°C) contains a mixture of D-RNA and R-RNA. Sedimentation analysis reveals two labeled components. One of them almost coincides with the radioactive peak of the 65°C fraction and the other gives a sharp peak with an S value of about 45. These peaks are well-separated and their analysis is feasible. The peak having a sedimentation coefficient of 45 S is R-RNA (it will be designated as R-RNA,) while the 25-30 S material is D-RNA identical with the 25-30 S material of the 65°C fraction. The conclusions drawn from the base compositions of the fractions are confirmed by hybridization experiments (77, 7 8 ) . R-RNA, and R-RNA, have a very low ability to hybridize with DNA, and unlabeled purified D-RNA does not compete with them for binding sites while purified rRNA does. On the other hand, 25-30 S radioactive D-RNA from the 40-55OC and from 55-60°C fractions effectively hybridizes with DNA and hybridization is suppressed by the addition of unlabeled D-RNA but not of rRNA (Table 111). Thus the following components of rapidly labeled RNA are revealed in the nucleolochromosomal RNA of Ehrlich cells: 45 S RNA-R-RNA, (in the 40-55OC fraction); 3 5 4 0 S RNA-R-RNA, (in the l M O 0 C fraction) ; and heterogeneous D-RNA with a maximum of distribution a t 25-30 S (these RNA's appear in the 55-65°C and partially in the 4&55"C fractions). Preformed nucleolochromosomal RNA contains the rRNAB peak (mainly in the 40-55°C and less in the 1&40°C fractions), the rRNAA peak (mainly in the 10-40"C and less in the 4&55°C fractions), and the D-RNA peak (mainly in the 5545°C and less in the 4045°C fractions). The peak of preformed D-RNA coincides with the 18 S rRNA peak. If the incubation time with the radioactive isotope is increased, the label begins to appear also in the main peaks of each fraction. Hence

FIO.2. Sucrose gradient profiles of RNA fractions prepared from Ehrlich ascites tumor cells by hot phenol method, after P3*: (a, b, d-f) pulse labeling for 1 hour; ( c ) pulse labeling for 4 hours; (g-i) actinomycin chase (actinomycin added for 3 hours after labeling period). (a) Total RNA; (b) total nucleolochromosomal RNA; (c-i) fractions prepared by hot phenol treatment (intervals of temperature are shown on the figures). Letters (R,, R2, D, r A , r B ) refer to peaks of newly formed UV absorbance; --x--, radioactivity (68,76). RNA. -0-,

TABLE I11 THECHARACTERISTICS OF RNA COMPONENTS OBTAINEDBY HOT PHENOL FRACTIONATION I N COMBINATION WITH ULTRACENTRIFUGATION I N SUCROSE

GRADIENTS (78)

Per cent of competition after addition of 5 or 10 times excess of nonlabeled RNA during hybridization Fraction (interval of temperature treatment) ("C)

Time of incubation with PJP (hours)

S value of component studied

10-40 10-40 10-40 40-55 40-55 55-63

1 4 4 1 1 1

35-40 35-40 28 45 25-30

-

(G

+ C)/(A + 1.65 1.60 1.85 1.40 0.88 0.81

Hybridization U) with DNAe D-RNA 1.0 1.0 0.8 1.1 8.9 11.2

0 0

0 75 70

Total ribosoma1 RNA

rRNAA

rRNAB

69 75 63 65 10 0

67 -

53 -

-

NUCLEAR RIBONUCLEIC ACIDS

275

it appears in the rRNAA (the 10-40°C fraction) and in the 18 S peaks of the 4 6 5 5 ° C fraction (mixture of rRNAB and D-RNA) and of the 5 5 6 5 ° C fraction (D-RNA). It is interesting that in these time periods ( 1 4 hours) the specific activity of thc RNA of phenol nuclci is considerably higher than that of the cytoplasmic RNA. Consequently not only heavy R-RNA but also newly formed true ribosomal RNA molecules are localized in the nucleolochromosomal complex. The results obtained with RNA’s isolated from some other tissues investigated are generally similar, but some controversies should be discussed. According to some authors, the 45 S component consists not of R-RNA but of D-RNA. Yoshikawa et al. (79, 80) fractionated the newly formed RNA of human amnion cells grown in tissue culture by chromatography on MAK columns and by ultracentrifugation in sucrose gradients. The RNA most tightly bound to the column is pure D-RNA with a sedimentation coefficient of about 50 S. Newly formed R-RNA with a sedimentation coefficient of 35-40 S is eluted earlier. The authors conclude that there is only one heavy component of R-RNA and that the second heavy peak is D-RNA. Brown and Gordon (81) found that, during early development in amphibia in the absence of R-RNA synthesis, a labeled 45 S component can still be found although this result was not confirmed in fish embryos (82). Finally, it was shown that a t very short incorporation times the heavy zone of the sucrose gradient is enriched with D-RNA (83). These observations should be interpreted with care as some of the experiments mentioned above were carried out using fractionation on MAK columns. It should be remembered, however, that, because of peculiarities of its secondary structure, D-RNA may be eluted from the column a t a higher salt concentration than R-RNA of similar molecular weight and thus the impression may arise that D-RNA is heavier than it really is. The sedimentation coefficient of newly formed D-RNA in the ultracentrifuge depends considerably on the ionic strength of the solution. Hence a t ionic strengths of 0.1 and 0.01, newly formed D-RNA has sedimentation coefficients of 40 and 30 S, respectively, and the values obtained a t slightly varying conditions may be different (84). Aggregation of D-RNA and R-RNA may also distort the picture ( 8 5 ) .Finally, studies on RNA from different sources definitely show two components in R-RNA [see for example (43, 44, 86)].The presence of only one peak of R-RNA in some experiments may be explained by poor resolution. However, the relation between the 25-30 S D-RNA in our experiments (68-76) and the 45-50 S RNA of Yoshikawa e t al. (79, 80) is

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not clear. Several explanations are possible. (1) The difference may depend on the conditions of ultracentrifugation. (2) It may reflect real differences in the molecular weights of newly formed D-RNA isolated from different sources. (3) It is possible that there are several classes of newly formed D-RNA in animal cells and, depending on incubation time, the label may prevail in one or another fraction. I n this connection, we may mention that D-RNA of Ehrlich cells contains, besides the 25-30 S peak, still heavier fractions, although they do not comprise the main portion of newly formed D-RNA (68). These facts indicating the complex nature of heavy RNA clearly demonstrate the importance of preparative fractionation of the total nucleolochromosoma1 RNA before the sedimentation analysis. Some interesting results have been obtained using electrophoresis of newly formed RITA in agar gel with subsequent measurement of the UV absorbance of agar films and autoradiography (87). To a first approximation, the electrophoretic mobilities of RNA’s are inversely proportional to their sedimentation coefficients. The resolution of this method is considerably better than that of sedimentation analysis; for example, the separation of total nucleolochromosomal RNA from rat liver using this method has revealed the presence of four rather than two peaks of newly formed RNA heavier than rRNA (88). RNA fractions from rat liver prepared using the hot phenol method yield electrophoretic patterns similar to those obtained by sedimentation analysis of newly formed RNA of Ehrlich cells. For example, in the 40°C fraction only one heavy R-RNA peak (R-RNA,) is present and, after long labeling rRNAAbut not rRNA, (89). The introduction of the preparative modification of this method may lead to further improvements in the separation of different components of newly formed RNA. I n the course of purification of RNA from phenolic nuclei, a fraction soluble is 2.5 M NaCl and containing low molecular weight RNA was usually discarded. This RNA fraction may contain low molecular weight RNA linked to histones as described by Huang and Bonner (90). This RNA has a relatively high specific activity but it has not been studied in detail. Its further investigation no doubt deserves attention. It should also be noted that many papers dealing with the characterization of newly formed cellular RNA have been published in the last few months. They confirm the presence of a t least two heavy R-RNA classes in nuclei and of relatively heterogenous D-RNA and give further details on their properties. Several of them are discussed in the following sections.

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NUCLEAR RIBONUCLEIC ACIDS

C. Characteristics of Chromosomal D-RNA1 1. NATUREOF CHROMOSOMAL D-RNA A discovery of DNA-like RNA in the nucleolochromosomal complex of animal cells leads to the suggestion that this type of RNA corresponds to newly formed mRNA (4 6 ). It has already been mentioned above that not only bulk but also rapidly labeled RNA isolated in the temperature interval 5 5 4 5 ° C has a DNA-like composition. However, additional evidence is necessary since a similarity in base composition is insufficient for any conclusion as to

Cytoplasmic RNA

I

Total RNA RNA (10-40")

20

40 60 DNAIRNA

I

80

100

0 0

,

10 3.3

,

20 6.7

,

X) mg Cytoplasmic 10 mg Total RNA

FIG. 3. Hybridization between chromosomal D-RNA and DNA. D-RNA was labeled for 4 hours. Right diagram shows the hybridization between D-RNA (100 p g ) and DNA (1 mg) in the presence of un'labeled total cellular and cytoplasmic RN.4 of Ehrlich cells (68, 84).

the informational nature of D-RNA. Such items of evidence are an ability to form molecular hybrids with DNA and a stimulation of the incorporation of labeled amino acids in ribosomes. Hybridizability of RNA from animal cells with the homologous DNA was first demonstrated by Hoyer et al. (91) using DNA-agar columns. It has been shown that newly formed RNA binds to DNA much more effectively than does preformed RNA and that nuclear RNA hybridizes better than cytoplasmic. Samarina et al. (68) studied hybridizability between DNA and different RNA fractions prepared from nuclei by the hot phenol method. It has been shown that the D-RNA of phenol nuclei effectively hybridizes with DNA while the hybridizability of nonribosomal R-RNA is lower by a factor of 10 to 25 (Fig. 3 ) . An increase in the DNA/RNA ratio leads to a proportional increase in the per cent of hybridization until saturation occurs and a further increase in DNA raises the per 'Papers reviewed in these sections prove that) D-RNA is localized in chromosomes and for convenience it will be referred to as chromosomal D-RNA.

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cent of hybridization only slightly. The percentage of hybridized material a t saturation depends on the time of labeling and varies between 30 and 10 to 15. However, even a t long labeling times, when newly formed and bulk RNA coincide, D-RNA gives considerably higher hybridization than does R-RNA. Another characteristic of mRNA is its ability to stimulate protein synthesis on ribosomes (92).A number of authors have shown that RNA prepared from isolated cell nuclei stimulates amino acid incorporation in the Nirenberg system much more effectively than does cytoplasmic RNA ( 9 3 ) . Brawerman e t al. (94) prepared an RNA fraction enriched with D-RNA from rat liver nuclei and demonstrated that it has a greater stimulating activity than the rest of the nuclear RNA. Also it was directly shown that D-RNA from rat liver and Ehrlich cell nuclei prepared by the hot phenol method is many times more effective in the stimulation of protein synthesis in the Nirenberg system than is cytoplasmic RNA or nuclear R-RNA (95, 68). It was inferred that the sti'mulating ability belongs to D-RNA and that it possesses this functional characteristic of mRNA as well. All these three characteristics are indicative of the fact that chromosomal D-RNA is, or a t least contains, newly formed mRNA of animal cells (68). Some interesting biological proofs of the informational nature of chromosomal D-RNA have been obtained recently by Zimmerman and Turba ( 9 ) , who treated rabbit reticulocytes with different RNA fractions from duck reticulocytes and studied the nature of the hemoglobin synthesized under these conditions. It was shown that the addition of duck RNA induces synthesis of duck hemoglobin in recipient cells and that the inducing activity resides in chromosomal D-RNA. Similar results were obtained in studies of induction of antihemoglobin synthesis by RNA isolated from lymphocytes of immunized animals. This phenomenon has been observed by a number of authors who used total cellular RNA for induction. In recent work by Stanchev, RNA was fractionated by the hot phenol method and it was shown that all the activity is contained in chromosomal D-RNA ( 6 5 ) . Lang (95a) demonstrated that rat liver D-RNA may induce synthesis of a-ketoglutarate transaminase in a cell-free ribosomal system. There is no doubt that the results of such induction experiments demand thorough checks and investigation of merhanisms of induction. Nevertheless the data listed are good evidence for the informational nature of chromosomal D-RNA. Thus the informational nature of chromosomal D-RNA can be regarded as proved and this is why the investigation of its properties becomes very important.

NUCLEAR RIBONUCLEIC ACIDS

279

2. PROPERTIES OF CHROMOSOMAL INFORMATIONAL RNA

In the first approximation from the molecular viewpoint, chromosomal informational RNA is very similar to other high molecular weight RNA’s, particularly rRNA (68). It consists of single threads containing helical regions. This is indicated by a typical steep (for RNA) melting curve. D-RNA chains are continuous-they do not contain intramolecular breaks. This is inferred from a characteristic rise in viscosity at elevated temperatures. It should be noted that this property, together with a high sedimentation coefficient (18-30 S), proves the nativity of this RNA. However, some properties of D-RNA are different from those of rRNA. They probably depend on peculiarities of its secondary or tertiary structure. D-RNA has a lower solubility in concentrated salt solutions as compared to rRNA (68). It has a considerably higher affinity for methylated albumin and its elution from MAK columns is effected by more concentrated salt solutions a t elevated temperature (70).The incomplete recovery of newly formed RNA from MAK columns claimed by many authors is probably a consequence of this phenomenon. The same properties are perhaps also reflected in the specific interaction of chromosomal D-RNA with particles participating in mRNA transport. D-RNA with a sedimentation coefficient of 18 S interacts with such particles while rRNA with a similar S value does not (see below). These differences are probably due to a lower helix content in D-RNA as compared to rRNA. I n this respect, D-RNA may resemble denatured DNA. The differences in the secondary structures of D-RNA and R-RNA may correlate with their different functions in protein synthesis although this question is not yet studied well enough. Attempts to obtain double-stranded RNA’s by annealing of D-RNA preparations have yielded negative results. Therefore, there are no complementary chains in chromosomal mRNA and this means that in animal cells only one DNA strand is active in mRNA synthesis (68). Similar data proving the absence of autohybridization between mRNA molecules were obtained on bacterial mRNA (96). Several investigations have been devoted to the sedimentation properties of D-RNA. In the analytical centrifuge, D-RNA reveals only one peak with a sedimentation coefficient of 18 S and some amount of heavier heterogenous material (68). Ultracentrifugation in sucrose gradient reveals a similar picture: there is a main peak with a sedimentation coefficient of about 18 S and more or less heterogenous material in the region of 20-30 S. Rapidly labeled chromosomal D-RNA, as has already been mentioned, is even

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G. P. GEORGIEV

more heterogenous (68, 75,97). The maximum of distribution is usually a t 25-30 S but a considerable part of the label sediments even faster (Fig. 4). It is probable that a t short labeling times chroniosomal D-RNA may have even higher sedimentation coefficients of the order of 45-50 S (79-83). The sedimentation properties of newly formed nuclear D-RNA have also been studied without its separation from the preparations of total cellular RNA. For this purpose, embryonic cells where there is no R-RNA synthesis were used (81, 82, 98), or cells where the synthesis of R-RNA was selectively suppressed by low doses of actinoniycin (83,99). I

25-30 S

I

FIG.4. Sucrose gradient profile of isolated D-RNA from Ehrlich ascites tumor cells labeled with H’-uridine for 30 minutes. -0-, UV absorbance; -A-, radioactivity.

The main conclusion from these experiments is considerable molecular heterogeneity of newly formed rapidly labeled D-RNA (distribution of sedimentation coefficients is between 15 and 60 S). The main component is found usually in the 25-30 S region and a more or less significant part of the material is in the heavy zone of the gradient (4060 S) . Thus data obtained by different methods on RNA from different sources are in satisfactory agreement with each other. It is proved that bulk chromosomal D-RNA from many cell types of higher organisms has a sedimentation coefficient of 18 S while newly formed rapidly labeled D-RNA sediments faster (25-30 S or even more). If we calculate molecular weights of these RNA’s from their sedimentation coefficients using empirical formulas available from the lit-

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erature (100, 101), we obtain the values for chroiiiosomal D-RNA and other RNA fractions of animal cells shown in Table IV. Thus molecular and metabolic heterogeneity are two distinct properties of chromosomal D-RNA. Molecular heterogeneity of chromosomal D-RNA is easily explainable if we recollect that mRNA is a template for synthesis of proteins having different molecular weights. In other TABLE I V SEDIMENTATION CONSTANTS AND CALCULATED MOLECULAR WEIOHTSOF MAIN OF RNA FROM ANIMALCELLS(68) COMPONENTS RNA class

Type of RNA

Sedimentation Molecular constant, S weight X 10-6 -

R-RNA

D-RNA

Ribosomal RNA: rRNAa rRNAB Heavy precursors of ribosomal RNA: R-RNA1 R-RNA2 Total chromosomal D-RNA from Ehrlich ascites cells: Maximum Limits Newly formed D-RNA from Ehrlich ascites cells: Maximum Limits Newly formed D-RNA from rat liver chromosomes: Maximum Limits Newly formed D-RNA from rat liver cytoplasm : Maximum Limits

28 18

1.6 0.65

45 35-40

4.7 2.7-3.7

18 16-30

0.65 0.5-2.0

25-30 16-50

1.3-2.0 0.5-6.0

20-30 15-40

0.8-2.0 0.4-3.7

10-12 8-25

0.2-0.3 0.12-1.3

words, the molecular heterogeneity of mRNA reflects considerable variation in the size of the cistrons or, more probably, of the operons transcribed. 3. METABOLIC HETEROGENEITY OF CHROMOSOMAL D-RNA

A metabolic heterogeneity of chromosomal D-RNA means that its different components are synthesized at essentially different rates (12, 68, 102). Consequently, a t early times the specific activity of the heavier D-RNA components is considerably higher than the specific activity of the main 18 S peak, and peaks of UV absorbance and of

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radioactivity do not coincide. Only a t longer labeling times does the radioactivity curve begin to follow the curve of W absorbance (Fig. 4). There can be several explanations of this phenomenon: (1) D-RNA is synthesized in chromosomes as a relatively long chain (>30 S), which is later split, still in chromosomes, into shorter fragments (about 18 S) . (2) Different fractions of chromosomal D-RNA, although having similar functions, are synt,hesized and turned over with essentially different rates so that the rate of synthesis and removal for heavier RNA fractions is higher than for lighter ones. (3) Chromosomal D-RNA is heterogenous not only metabolically but also functionally. Besides the usual mRNA synthesized in chromosomes and migrating to the cytoplasm, there is a special class of D-RNA functioning in chromosomes themselves at the site of its synthesis. One may suggest that this RNA bears certain regulatory functions.

The last explanation becomes probable because of the fact that the per cent of chromosomal D-RNA is very high. For example in Ehrlich ascites cells about one half of the total D-RNA is in chromosomes. These data also indicate indirectly that not all chromosomal D-RNA is a precursor of the cytoplasmic mRNA. To investigate the nature of the metabolic heterogeneity of chromosomal D-RNA, experiments on competitive hybridization between DNA and various classes of cellular RNA (84) were undertaken. This method permits us to answer the question whether identical or different base sequences are contained in the preparations of RNA studied. For hybridization experiments the following RNA preparations were used: (1) D-RNA labeled with P3? or H3-uridine for 0.5-1 hour (rapidly labeled D-RNA) ; (2) D-RNA labeled for 24 hours (total chromosomal D-RNA) ; (3) the heavy component of rapidly labeled D-RNA isolated by ultracentrifugation in sucrose gradient from D-RNA preparation labeled for 30 minutes; (4) the main component of chromosomal D-RNA with sedimentation coefficient 18 S isolated after prolonged incubation. At first relative hybridization of rapidly labeled and total chromosomal D-RNA with DNA was studied. It was demonstrated that, although total and rapidly labeled chromosomal D-RNA can hybridize effectively with homologous DNA, the per cent of hybridization for rapidly labeled heavy D-RNA is higher than for total chromosomal D-RNA, and particularly 18 S D-RNA (Fig. 5 ) . It was inferred that rapidly labeled D-RNA contains replicas from a larger number of cis-

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NUCLEAR RIBONUCLEIC ACIDS

trons than does the 18 S component or that these classes of RNA have different base sequences. These results are in good agreement with the data of Hoyer et al. (91) who found that a t prolonged incubation, the per cent of labeled RNA hybridized with DNA is decreased. Then the hybridizability was studied between DNA and D-RNA uncomplexed after first annealing with DNA. I n these experiments uncomplexed D-RNA has been transferred subsequently 4 or 5 times into new portions of DNA (Fig. 5 ) . It has been found that after the first transfer the hybridization per cent falls rapidly, but after the third or fourth transfer it stops a t a relatively low but significant constant level, a t both long and short labeling times. 30% labeled I hour

D- RNA labeled 20 hours

20 %

10%

I I I I I I r n Y

I I t I J I r n Y

NN of transfer

FIG.5. Successive hybridizations of chromosomal D-RNA from Ehrlich ascites tumor cells with DNA. DNA was introduced as an insoluble gel, obtained by cross linking DNA films with UV light. After annealing, DNA was freed from RNA by centrifugation and a new portion of DNA was added to the RNA solution (84).

It is concluded that the bulk of D-RNA is represented by replicas from a relatively limited number of cistrons but not less than 0.5% of all DNA. These molecules give constant low hybridization in experiments with transfer of D-RNA into new DNA portions. These RNA’s prevail in samples of newly formed RNA obtained after prolonged incubation with radioactive precursors. Rapidly labeled heavy D-RNA also cont.ains replicas from these cistrons (sequential transfers give the same hybridization level) but a considerable fraction is represented by replicas from other cistrons; that is, it is more heterogeneous and contains more molecular species than the main part of the chromosomal D-RNA. Then a competition between labeled chromosomal D-RNA and unlabeled cytoplasmic RNA for binding sites on DNA was conducted (Figs. 3 and 6 ) . It was shown that this competition does exist and thus the

0.2 I

1

30-25

1

s

I20 0.15

-

P n

I 80

0

5

0.09 40

10

20

40

I

I

Fio. 6. Hybridization with DNA of different components of chromosomal D-RNA isolated by ultracentrifugation in sucrose gradients, and the influence of cytoplasmic RNA on this hybridization (84). Chromosomal D-RNA was isolated from Ehrlich ascites carcinoma cells after (a) 30 minutes or (b) 20 hours incubation of cells with Ps'in v i v a The fractions I and I1 were obtained from sucrose gradients and used in hybridization experiments. Conditions and results of hybridization are as follows: DNA denatured (in the form of gel) (mg) 2 2 2 2

Competitor (mg) Labeled E. coli RNA ( p g ) RNA

I, 40 I, 40 11, 40 11, 40

5 5 -

Cytoplasmic RNA

Hybridization (%)

Competition (%)

5 5

16.4 13.1 8.8 4.3

20 50

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NUCLEAR RIBONUCLEIC ACIDS

transfer of a t least a part of the chromosomal D-RNA into the cytoplasm was proved unequivocally. However, even in the presence of a large excess of cytoplasmic RNA, the inhibition of hybridization is far from complete. On the other hand, the addition of an excess of total cellular RNA as a competitor lowered the binding of chromosomal D-RNA almost to zero values. This result may be explained by the existence of a special class of chromosomal D-RNA that is not transferred into the cytoplasm, but is functioning a t the site of synthesis in chromosomes. The competing effect of cytoplasmic RNA is much more pronounced if long-labeled D-RNA or the isolated 18 S component of D-RNA is used for hybridization, and it is considerably less if pulse-labeled D-RNA (30 minutes of isotope incorporation) or heavy D-RNA from a similar short-term experiment (Figs. 3 and 6) is used. From the results of these hybridization and competition experiments, one may suggest that heavy D-RNA (with a distribution maximum in the 25-30 S zone) contains both the usual mRNA, transferred later into the cytoplasm and nuclear sap (D-RNA,) , and a special type of D-RNA, functioning in chromosomes (D-RNA,) . On the other hand, bulk D-RNA with a distribution maximum in the 18 S zone comprises mainly mRNA for the cytoplasm (D-RNA,). It is possible that the usual mRNA (D-RNA,) is synthesized in the form of heavy polycystronic molecules and is then split into shorter chains with the formation of 18 S D-RNA,. Thus the following hypothetical scheme of interaction between different D-RNA classes can be drawn: chromosomal mRNA of cytoplasm heavy rapidly D-RNA, + 18 S D-RNA + and nuclear sapfunctioning labeled D-RNA TI-RNA, + and breakage in chromosomes

{

{ I

{

The role of the D-RNA functioning in chromosomes is unknown. One may suggest that it serves as a template for nuclear protein synthesis including proteins participating in regulation of gene activity (see below). It may also play a more direct role in regulatory processes. However, further experimental approaches are necessary to answer this question. The suggestion of the existence of two functional classes of D-RNA is in good accord with some other facts. So some investigators point out that a fraction of newly synthesized cellular D-RNA is unstable and degrades upon transfer of the cells to nonradioactive medium or in the presence of actinomycin (3, 103, 104). Under such conditions, heavy D-RNA disappears completely ( 8 3 ) . On the other hand, synthesis of

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G. P. GEORGIEV

cellular proteins is not affected by these conditions. The conclusions were drawn ( 1 ) that the genuine mRNA for the synthesis of most cellular proteins is stable and (2) that, besides it, the nucleus contains a rapidly labeled RNA that is broken in the nucleus (105, 106). The stable mRNA may be identified with stable 18 S D-RNA. It is interesting that maximum template activity of cytoplasmic RNA for protein synthesis resides in the 18 S region of sucrose gradients (107,108). I n summary, we wish to stress that nowadays the presence of newly formed mRNA in cellular D-RNA is strictly proved. But besides it, D-RNA probably contains informational RNA’s with special functions and their investigation is the task of the immediate future.

D. The R-RNA of

the Nucleolochromosomal Complex

It has been already mentioned that, in short incorporation experiments, R-RNA is represented by two heavy peaks, R-RNA, and R-RNA,, with sedimentation constants of 45 S and 35-40 S, respectively (Fig. 1 and 2 ) . What is the nature of these components? Perry (46) has shown that small doses of actinomycin inhibit primarily the synthesis of heavy fractions of newly formed RNA and subsequent incorporation of the label in ribosomal RNA. A conclusion was drawn that heavy RNA peaks are the precursors of rRNA. Similar results were obtained in experiments with the “actinomycin chase” using HeLa cells (103). After pulse labeling, when the radioactivity was predominantly in heavy peaks, a large dose of actinomycin suppressing all cellular RNA synthesis was given. This leads to disappearance of the label from heavy peaks and its appearance in ribosomal RNA. This indicated directly that heavy R-RNA’s are the precursors of true ribosomal RNA’s (Fig. 7 ) . Studying the kinetics of incorporation of radioisotopes in different RNA fractions, Sherrer e t al. (103) suggested the following course of R-RNA conversions: R-RNA, + R-RNA, + rRNAA and rRNA,. Similar conclusions were drawn by Rake and Graham ( 7 1 ) . I n such experiments, the situation is complicated by the breakdown of a part of the newly formed RNA in the presence of actinomycin; however, in the experiments of Sherrer et al. (103) this breakdown could not explain the disappearance of all heavy RNA. We have conducted similar experiments on Ehrlich ascite cells using the hot phenol method for RNA fractionation (76, 109) (Fig. 2). It was shown that, 3-4 hours after actinomycin addition, R-RNA, and R-RNA, disappeared completely from those fractions where they were present before addition of the antibiotic. In the 3540°C fraction, where R-RNA, is found after pulse-labeling only rRNA, could be found aftcr incubation with actinomycin. On the other hand, in the 55°C fraction

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NUCLEAR RIBONUCLEIC ACIDS

where, besides D-RNA, R-RNA, 1s present, only the 18 S labeled component was detected after actinomycin. Although labeled D-RNA is present in this region, it has a peak a t 25 S. The base composition of the 18 S peak is intermediate between those of DNA and RNA. Thus instead of R-RNA,, only rRNAB is present in the 55°C fraction (Fig. 2). Proper controls have shown that incubation in actinomycin does not lead to redistribution of label between different RNA fractions. These data were a basis for the suggestion that there exist two different polycistronic precursors for the two ribosomal RNA components : R-RNA, for rRNA, and R-RNA, for rRNA,.

200

I00

I

I

0

I

0

I

60’

200

0.200

I00

0.100

0

10

20

30

40

50 0

10

20

30

40

50

FIG.7. Effect of actinomycin D chase on the newly formed RNA fractions of HeLa cells prelabeled during 30 minutes with H3-uridine (103). Closed circles = UV absorbance, open circles = radioactivity.

To obtain additional data on the relations between heavy R-RNA’s and true ribosomal RNA, R-RNA, and R-RNA2 were hybridized with DNA in the presence of rRNA’s (78) (Table 111).Both R-RNA, and R-RNA, compete with ribosomal RNA for binding sites and do not compete with D-RNA. These experiments prove that R-RNA, and R-RNA, are the precursors of rRNA. I n experiments with R-RNA,, which is easier to obtain in a highly purified state, rRNAA is a somewhat more effective competitor as compared to rRNA,, but the latter also competes with R-RNA,. However, this circunistance may be due to the presence of about 60% coinmon nucleotide sequences in relatively long regions of rRNA, and rRNA, (110). This latter circumstance im-

288

G. P. GEORGIEV

poses serious restrictions on further use of the competitive hybridization method for detecting precursors of rRNAA and rRNAB. Some indication of a different origin of ribosomal RNA components resides in data on the base composition of different R-RNA’s. Ribosomal rRNA, and rRNAB differ considerably in their base composition, rRNAA being more rich in GC pairs (111, 112). It is interesting to note that the base composition of R-RNA, corresponds almost exactly to the composition of rRNAn (76, 78) (see Figs. 2 and 6) and this is additional evidence in support of a different origin of ribosomal RNA components. Other facts concerning the transition of heavy R-RNA to rRNA’s have been obtained by Perry (112a); they are discussed by him in this volume. It should also be noted that not all heavy R-RNA, a t least in actinomycin, is converted to rRNA. A considerable portion breaks down, but the cause of this phenomenon is obscure ( 7 6 ) . I n summary, one may conclude that the R-RNA of the nucleolochromosomal complex includes both true newly formed ribosomal RNA and its heavy polycistronic precursors.

111. Fractionation of Subnuclear Structures and Nuclear Ribonucleoproteins Phenol methods are very useful for the isolation and characterization of nuclear RNA classes but they do not yield information concerning the intracellular organization of these RNA’s since phenol treatment results in denaturation of proteins. Questions concerning the localization of nuclear RNA and the nature of complexes between RNA and proteins should be resolved by different methods of fractionation. These gentler methods, however, cannot prevent partial RNA depolymerization occurring as a result of RNase action.

A. Principles of Methods 1. SALTFRACTIONATION OF CELL NUCLEI It has already been mentioned that salt fractionation consists of sequential treatment of nuclei by neutral salt solutions of different ionic strength (28, 50, 113, 114). Usually, nuclei obtained a t low ionic strength are treated with salt solutions of ionic strength 0.05-0.15. This results in the liberation of the so-called “globulin fraction.” Extraction with concentrated salt solution (1-2.5 M NaCl) follows, which solubilizes the D N P fraction. The relative content of DNA, RNA, and protein in these fractions obtained from rat liver nuclei is presented in Table V. Detailed cytological and electron microscope studies of these

CHEMICAL

Fraction Globulin

A N D RIORPHOLOGICAL

Main ultrastructural elements

0.05-0.2 M neu-

Free 150-200 A particles, nuclear ribosomes, sRNA, soluble proteins 30 S particles containing mRNA, soluble proteins D N P threads dissociated to DNA and protein

The same, p H 8

Residue

Nature of RNA in fraction

Extraction medium tral salt, p H 6-7

Deoxynucleoprotein

TABLE V COMPOSITION O F THE NUCLEAR FR.4CTIONS

2-2.5 M NaCl

I{

Chromosomal nucleonemata Nucleolar nucleonemata Nuclear membranes

rRNAA and rltNAB, tRNA, some mRNA

I

mRNA mRNA, low M.W. RNA

rRNAa and rRNAB R-RNAI and R-RNA,

OF

R.4T LIVER

Chemical composition ( % of dry weight of nuclei) RNA, 1.5; protein,

Localization in cell nucleus Nuclear sap

23

-

Predominantly in chromosomes

DNA, 22; RNA, 0.5; Chromosomes (inhistone, 33; noncluding material, histone protein, 10 penetrating nucleolar zone) RNA 1.0 Nucleolus and partial protein, 4.0 chromosomes (residual chromosomes) Protein 5.0

Nuclear membrane

290

G.

P. GEORGIEV

fractions and residues remaining after extraction demonstrated their correspondence to certain subnuclear struct,ures and ultrastructures and this explains why this approach is of considerable interest. Observation of nuclei under an electron microscope reveals a number of ultrastructural elements serving as building blocks for such structures as chromosomes and nucleoli. The best characterized element is a deoxyribonucleoprotein thread 100-200 A in diameter (116-1 19, 34), designated below as D N P fibril. These threads are concentrated in chromatin but can be visualized also in the nucleolar zone. Another important ultrastructural element is a thread 100 A in diameter with (sometimes without) attached ribosomelike granules 150 A in diameter (29, 34, 120). This element is called nucleonema (29). Similar ultrastructures were described later by Ris (121) who designated them as “rough threads” and by Busch (122) who called them “nuclear ribonucleoprotein network.” Nucleonemata are concentrated mainly in the nucleolus but are found also in chromatin (chromosomes). Free ribosomelike granules not bound to any fibrillae can also be observed in nuclei as well as granules of another diameter and accumulations of fine amorphous material (mainly in the nucleolus region) [see (123) for details]. These facts prove that the main structures of the cell nucleus are built from several ultrastructural elements. Nucleoli consist mainly of nucleonemata bound to thicker clusters called “nucleolonemata” (1.84) but containing also D N P threads, free granules, and amorphous substances. Chromosomes contain nucleonemata of chromosomes besides D N P threads. Since both chromosomes and nucleoli have common ultrastructural elements and nucleoli can be regarded as derivatives of chromosomes, it is frequently difficult to draw a line between them and usually we use the term nucl6olochromosomal complex or apparatus to designate all threadlike nuclear elements. Amorphous material filling the space between fibrillar elements, including free granules, is designated “nuclear sap.” Salt fractionation of isolated nuclei permits one to separate the substance of some of these basic elements and therefore to obtain RNA or ribonucleoproteins localized in different nuclear ultrastructures. Dilute salt solutions extract from nuclei mainly the nuclear sap, i.e., soluble proteins and free particles not bound to any fibrillae. All fibrillar elements, including D N P threads, nucleonemata, and nuclear membrane, remain. Concentrated salt solutions (0.6-2.5 M NaCl) extract the substance of the D N P threads. The residue, after extraction with concentrated salt solutions, consists of nuclear membranes and nucleonemata (27-29, 34). Concentrated salt dissociates DNP threads into DNA and protein (125), and for this reason the salt fractionation

NUCLEAR RIBONUCLEIC ACIDS

291

method is useful for the analysis of structural elements of nuclear sap and of nucleonemata, but not of intact deoxynucleoprotein. RNA can be detected in all three fractions, in the ratio +3:1:2. It can be inferred from these data that fractionation of nuclear ribonucleoproteins depends mainly on whether they are bound to a fibrillar component or not and does not depend on the localization of a given ribonucleoprotein species within the nucleus. Free granules, both from chromatin and from nucleolus, remain in the globulin fraction while nucleonemata from nucleolus and from chromatin remain in the final residue.

2. MECHANICAL SEPARATION OF NUCLEAR HOMOGENATES Another important approach is mechanical separation of nuclear homogenates (49-5S, 126, 127). The nuclei are disrupted in the appropriate medium (usually a t low ionic strength in the presence of Mg2+) either by sonication or by a French pressure cell. The nuclear homogenates so obtained are fractionated by differential centrifugation, using varying concentrations of sucrose. This type of fractionation gives subnuclear elements differing in their size and density: intact nuclei, nucleoli, one or two fractions of chromatin, ribosomes, and nuclear sap. Since low ionic strength and Mg2+prevent redistribution of the material between fractions, this method permits one to obtain different structural components of the nucleus without their separation on ultrastructural elements. Hence nucleoli obtained by this method contain not only nucleonemata but also DNP fibrils and many soluble components localized in the nucleolus (53, 128, 129). The fractionation thus takes place according to a mechanical principle. The advantage of this approach is the possibility of isolation of nuclear structures with minimally damaged macroorganization. The main defect is extensive cross contamination of fractions. This depends on the fact that the interphase nucleus lacks sharp boundaries between different nuclear structures, which are continuous and penetrate into each other. This is why nucleoli as a rule are contaminated with chromatin functionally unrelated to them. This circumstance limits the resolving power of the method and requires further improvements. The fractions of particles sedimenting after nucleoli are insufficiently characterized. The characteristics of nuclear ribonucleoproteins obtained by both methods are reviewed below.

B. Ribonucleoproteins of Nuclear Sap L‘Nuclearsap” or the “soluble phase” is the material more or less uniformly distributed throughout the cell nucleus and comprising all the nuclear volume between subnuclear fibrillar structures, chromosomes,

292

G. P. GEORGIEV

and nucleoli. I n those cells where chromosomes and nucleoli occupy only a small part of the nucleus, the nuclear sap is easily available for studies. The microchemical data of Edstrijm (130) show that the nuclear sap of oocytes contains a large amount of RNA with a base composition intermediate between that of total RNA and DNA. However, in most interphase nuclei, despiralized chromosomes fill all the nucleoplasm, and nuclear sap occupies only the interstices between nuclear fibrillar elements, D N P threads and nucleonemata. Therefore, for its isolation, one should use extraction by neutral salt solutions, and the possibility of extraction not only of nuclear sap but also of some material of nucleonemata and D N P fibrils should be taken into consideration. However the bulk of these ultrastructures remains in the sediment. Nuclear sap contains about 20% of the nuclear proteins and about 30-50% of the total nuclear RNA (114, 131, 132). These values are obtained for nuclear sap from different sources: calf thymus, rat liver, kidney, spleen, Ehrlich cells, and others. They refer to nuclei isolated in sucrose media. If nuclei are isolated in nonaqueous media, the values are somewhat higher. RNA from the nuclear sap of lymphocytes has the base composition of ribosomal RNA while RNA from rat liver nuclear sap has an intermediate composition, (G C)/(A U) = 1.25 (N,,@4-26) indicating the presence of D-RNA. The rate of incorporation of labeled precursors into this RNA is intermediate between the rates of labeling of nucleolochromosomal RNA and cytoplasmic RNA (21,24-26,133).

+

+

1. RIBOSOMES AND tRNA I n 1960, it was shown in a number of laboratories that extracts of nuclei from calf thymus, rat liver, and other tissues corresponding to the nuclear sap fraction contain ribonucleoprotein particles sedimenting in ultracentrifugation a t 56-105,000 g ( 3 1 4 3 ) .These particles contained highly polymerized RNA and were active in protein synthesis both in vivo (32) and in vitro (31). Their chemical composition (60% RNA, 40% protein) and their appearance under the electron microscope permitted their identification as ribosomes (134-138). Extraction of the nuclear sap removes free ribosomelike granules from the nuclei. Probably the free nuclear granules scattered throughout the nucleus correspond to the ribosomes isolated from nuclear sap extracts. Nuclear sap ribosomes are capable of protein synthesis in witro, and the requirements of the system are similar to those of the cytoplasm (137). Zbarsky and Samarina (139) incubated liver slices prelabeled with CY4-tyrosine and have demonstrated the migration of label from nuclear ribosomes to the

NUCLEAR RIBONUCLEIC ACIDS

293

soluble phase of nuclcar sap. Thus ribosomes isolated from the globulin fraction are probably functional ribosomes participating in the synthesis of soluble proteins of cell nucleus. Such ribosomes can be prepared easily froin thymus nuclei but their yield from liver nuclei isolated in aqueous media is insignificant. This enabled Siekevitz (53) to suggest that ribosomes of the globulin fraction are of extranuclear origin. We are inclined to think, however, that in the case of liver nuclei a loss of a part of ribosomes is possible as has been demonstrated earlier for soluble proteins (140). Frenster e t al. (31) have shown that incorporation of label into ribosomes of the globulin fraction of isolated nuclei occurs only under conditions permitting intranuclear protein synthesis. Checks on the contamination of nuclear ribosomes by labeled cytoplasmic proteins also prove their intranuclear origin ( 3 2 ) . But although the existence of ribosomes participating in protein synthesis in the nuclear sap is proved, their amount can be estimated only approximately. Probably it differs in nuclei from different sources (141). After ultracentrifugation of the nuclear sap extract for 2-4 hours, about one third to one quarter of its RNA remains in the supernatant. This is a less polymerized RNA with a sedimentation coefficient of about 4 S (32, 133). Thus the nuclear sap contains, beside ribosomes and ribosomal RNA, also sRNA. The base composition of the sRNA is intermediate between D-RNA and tRNA and is indicative of the fact that it consists of true tRNA and mRNA fragments. The presence of tRNA in this fraction is proved by its ability to bind activated amino acids in the presence of aminoacyl-tRNA synthetase (SO,148). The formation of aminoacyl-tRNA in the frog oocyte nuclei has been shown histochemically by pulse-labeling with radioactive amino acids and subsequent radioautography, including controls treated with hydroxylamine and RNase. These experiments showed that tRNA is evenly distributed throughout the nucleus (143). If RNA synthesis is suppressed with actinomycin, this fraction continues to incorporate Ps2.After alkaline hydrolysis, the label is present mainly in cytidylic acid and this proves the turnover of the CCA terminus typical of tRNA. Thus, nuclear sap contains tRNA and, if one remembers the presence of aminoacyl-tRNA synthetase in this fraction (SO), the conclusion may be drawn that nuclear sap contains a completely functional system for protein synthesis. It should be noted that dilute salt solutions extract all nuclear sRNA. However this does not mean that it is absolutely evenly distributed in the nucleus (see below). We have already said that in pulse labeling experiments nuclear sap

294

G.

P.

GEORGIEV

RNA is labeled rather actively. It is true for both RNP particles and for sRNA (133). One may suggest that all or part of rRNA and tRNA are transient RNA’s migrating from the site of their synthesis in the nucleolochromosomal apparatus into the cytoplasm. However, to prove this suggestion one should demonstrate that thc labcl is incorporated in rRNA and tRNA but not in contaminating D-RNA. The analysis of the base composition of newly formed high and low polymer nuclear sap RNA after short labels with P3*gives a specificity coefficient, (G C)/(A U ) , of about 1.25. Therefore, besides newly formed D-RNA, about 60% of newly formed RNA of these fractions is represented by rRNA and tRNA and the high incorporation of label into nuclear sap RNA probably reflects synthesis and transport of rRNA and tRNA ( 1 4 ) .

+

2. mRNA

AND

+

mRNA-CONTAINING RIBONUCLEOPROTEINS

It has already been mentioned that the base composition of nuclear sap RNA of rat liver and of Ehrlich cell nuclei is intermediate between D-RNA and R-RNA, indicative of the presence of D-RNA (2?4,26,144). Nuclei from calf thymus give pure R-RNA but, on repeated extraction, an RNA with very high specific activity, most probably D-RNA, begins to solubilize (31). If rat liver nuclei are extracted by several portions of dilute salt solution, the composition of RNA extracted shifts to that of D-RNA (35). The impression arises that, besides ribosomes and tRNA easily extractable from nuclei by dilute salt solutions, D-RNA is gradually extracted from nuclei and its solubilization occurs after breakage of some links between this RNA and the chromosomal complex. Samarina et al. (35) have studied the conditions of extraction of D-RNA from the nuclei and have shown that it is enhanced by raising the p H of the extracting solution (0.14 M NaCl, 0.001 M MgCl,, 0.01 M tris) to 7.8. If the first extraction is carried out a t p H 7.0 and the subsequent one a t pH 7.8, the first extract contains predominantly newly formed R-RNA while the second and the third contain almost pure newly formed D-RNA. By this technique, one may solubilize up to 70% of the labeled D-RNA of rat liver or Ehrlich ascites cells. A part of it, however, always remains in the nuclear residue (Table VI). The possibility of preparing D-RNA in the soluble state without using such denaturating agents as phenol or sodium dodecylsulfate is of considerable interest since it would help to answer questions as to the state of the newly formed mRNA in cell nucleus. Ultracentrifugation of D-RNA containing nuclear extracts in sucrose gradients reveals two peaks of UV-absorbing material having sedimen-

295

NUCLEAR RIBONUCLEIC ACIDS

tation coefficients of 30 and 4 S. The bulk of the newly formed RNA is bound to the faster peak (30 S) (Figs. 8a and b). Analysis of the 30 S peak (35, 145) shows that this is an mRNAcontaining ribonucleoprotein. The nucleoprotein nature of the peak is proved by its following properties: (1) the low ratio of A260/A280, equal to 1.45-1.5; (2) high protein content as shown by the Lowry reactionup to 75% of all the substance of the 30 S peak is protein; these data, however, are not very reliable since not only ribonucleoproteins but some other proteins may be present in the peak region; (3) a more TABLE VI THE DISTRIBUTION OF NEWLY FORMED HIGH-POLYMERIC RNA I N FRACTIONS FROM ISOLATED RATLIVERNUCLEIBY A SALTFRACTIONATION OBTAINED PROCEDURE"

Fraction number 1 2 3 4 5

+6

6

a

Extraction medium NaCl (0.14 M ) , MgClz (0.001 M ) , tris (0.01 M ) , pH 7.0 The same, pH 8.0 The same, pH 8.0 The same, pH 8.0 Residue after extractions 1-4 Residue after extractions 1-4 and 2.5 M NaCl (nucleonemal fraction)

Specific activity of RNA (cPm/ mg RNA)

Per cent of Base compototal radiosition of activity (% of newly total newly formed RNA, formed RNA) (G C)/(A TJ)

+

+

9,200

6.0

1.30

37,500 36,800 45,000 42,000

15.2 14.0 3.5 61.3

0.91 0.78 0.80 1.31

59,000

30.5

1.65

Animals received Pa2 1 hour before killing:

direct proof of the ribonucleoprotein nature of the 30 S particles is a drastic fall in the sedimentation velocity of both radioactive and UVabsorbing materials after deproteinization of the extract (12-18 S instead of 30 s) (Fig. 8a) ; (4) in experiments on amino acid incorporation into proteins, a small but significant peak corresponding to the peak of labeled RNA is revealed in sedimentation diagrams (Fig. 8b). Thus it, is proved that D-RNA can be bound to protein, forming a ribonucleoprotein particle. The nature of the RNA component of 30 S particles was studied. Its base composition corresponds to that of D-RNA, it forms molecular hybrids with DNA, and this hybrid formation may be suppressed by the addition of nonlabeled chromosomal D-RNA.

296

G . P. GEORGIEV

-

Nuclear extracts Native ( , --C-Deproteinized (.--*--)

0.45

600

p 0.30

400

n 0

0.15

z

V

200

(a) 10

30

40

Nuclear extract

0.75

1500

8 0.50

I000

n 0

iF V

500

0.25

FIG.8. Ribonucleoprotein particles containing mRNA, isolated from cell nuclei (146). (a) Sucrose gradient profiles of mRNA-containing nuclear extracts and of

high-polymer RNA prepared from the extracts (results of two separate experiments -&, W absorption; -A- -A-, are shown on the same diagram). radioactivity. (b) Sedimentation profile of mRNA-containing nuclear extracts. RNA -A-, radiois labeled with Pa*,protein with C". 4-,UV absorption; -X-, activity. (c) Interaction of free mRNA and rRNA labeled with P3*with nuclear extracts. Only 18 S components of the corresponding RNA's were used. Their sedimentation profiles are shown on the upper diagram (dotted line). In the first experiment, the RNA of extract wm labeled with C". 4-,W absorption; -x-, Pa'; -0-, C" (dotted line). (d) Interaction of 18 S mRNA labeled with P3' with different components of nuclear extract, isolated earlier by sucrose gradient ultracentrifugaUV absorption, -Xradioactivity. Arrows indicate the position of tion; free mRNA.

+-

-*

It is inferred that a t least the rapidly labeled RNA of the 30 S particles is a chromosomal D-RNA, i.e., the newly formed mRNA of the cell. The RNA of these particles is tightly bound to protein. Sedimentation properties of t,he particles are constant a t ionic strengths of 0.01 to 0.4 and in the presence of 0.001 M EDTA. I n the electron microscope they appear to be discs with dimension of 180 x 180 x 80 A.

297

NUCLEAR RIBONUCLEIC ACIDS

1500

1000

500

(C,)

FIQ.8c and d. See opposite page for legend.

298

G . P. GEORGIEV

It has also been found that similar labelcd mRNA-containing particles can be prepared by the addition of purified labeled niRNA to nuclear extracts (146‘). For these experiments, the 18 S peak of chromosomal D-RNA was used and after mixing with the different components of the extract a considerable part of the label was found in the 30 S region (Fig. 8c). This binding is specific for mRNA since, if labeled rRNA with the same sedimentation coefficient is added, the label is not incorporated into 30 S particles. The study of the binding of mRNA with the different components of the extract has shown that it interacts not with low molecular proteins but with preexisting 30 S particles (Fig. 8d) (146‘). This interaction was also demonstrated in the following manner. The 30 S peak obtained by gradient ultracentrifugation was added to gels of purified mRNA crosslinked by UV irradiation. After 15 minutes of incubation a t O”, about 10% of the label was bound to mRNA gel. Thus mRNA conjugates not with protein but with pre-existing particles. The nature of the 30 S particles remains obscure. It was suggested previously that these particles are complexes of small ribosomal subunits with mRNA (146). This suggestion correlates well with the high degree of homogeneity of the 30 S particles and the mode of their interaction with mRNA. I n this connection, it is interesting that the binding site for mRNA is located on the smaller ribosomal subunit (147). Similar types of particles were also postulated in the cytoplasm of animal cells (148-1 50) . However, recent results concerning the properties of nuclear 30 S particles do not support this concept. The base composition of total RNA isolated from particles is of the AU type. Thus the only RNA in the particles is D-RNA. Another possibility is that the 30 S particles of nuclei are “informosomes” (see below), mRNA-protein particles observed in the cytoplasm of embryonic cells. However some properties of 30 S particles (high homogeneity, buoyant density) differentiate these two types of ribonucleoproteins. One can suggest that the observed chromosomal particles are a special unknown type of protein or lipoprotein complex that conjugates with mRNA. The possible function of mRNA-containing ribonucleoproteins is discussed below in connection with mRNA transport. Concluding this section, it should be noted that using mechanical fractionation of nuclei one can obtain a small amount of material low in DNA but containing ribosomes and soluble proteins (53, 126). However, the bulk of the nuclear sap material obtained under these conditions of fractionation goes with the chromatin debris and possibly nucleoli. The nature of the interaction between soluble proteins and

NUCLEAR RIBONUCLEIC ACIDS

299

nucleoproteins is obscure. This question is surveyed in other reviews (6, 151).

C. RNA of the Deoxynucleoprotein Fraction (Chromosomal RNA) It has been already mentioned that salt fractionation may lead to some loss of RNA from chromosomes. It may well be that, in vivo, mRNA-containing 30 S particles are somehow associated with D N P threads of chromosomes, being separated from them only after repeated extraction. Experiments on the mechanical fractionation of nuclei show that the main part of the D-RNA may be detected in the chromatin debris sedimenting after the nucleoli (15.2). Unfortunately, a more detailed characterization of these D-RNA containing structures is not given. Valuable information concerning the nature of chromosomal D-RNA has been obtained by Edstrom et al. (130, 153) who, using a microchemical technique, has studied the base composition of RNA from different regions of the oocyte nucleus. It was shown that the RNA of chromosomes has a DNA-like composition, that nuclear sap RNA has an intermediate composition, and that nucleolar RNA is of the R-RNA type. We have already pointed out that a considerable portion of the chromosomal D-RNA exists in the form of mRNA-containing 30 S particles. On the other hand, a part of the highly polymerized D-RNA always remains in the nuclei even after repeated extractions with solutions a t pH 7.8. It might well be that this RNA is a part of the complex between template (DNA), enzyme (RNA polymerase), and the product (mRNA). Complexes of this type have been demonstrated in cell-free systems (154). This question is discussed in more detail in the next section. The study of RNA of the D N P fraction (obtained by extraction of nuclei with 2-2.5 M NaCI) actually has not yet been reported. The highly polymerized RNA of this fraction is claimed to have a base composition of an intermediate type, but this may arise from contamination by nucleolar RNA. Huang and Bonner (90) have found that histones dissociated from DNA in concentrated CsCl solution are bound to RNA and this complex can be isolated in a CsCl density gradient. This RNA is of low molecular weight (about forty nucleotides in length) and exceedingly rich in 5,6-dihydrouridylic acid. The authors suggest that this RNA fraction may play a structural role linking histone subunits in large complexes. Another suggestion of the same authors is that this RNA is a component of a repression system (156).

300

G . P. GEORGIEV

Although these results need confirmation, they point to the existence of a whole new class of RNA with unknown functions and probably strictly chromosomal localization. Thus, chromosomes contain mRNA within 30 S ribonucleoprotein particles bound somehow with DNP fibrils. They also contain D-RNA within other not yet known complexes and a special histone-linked lowpolymer RNA.

D. RNA of Nucleoli and Nucleonemata The data on the nature of nucleolar RNA originate from three sources. Edstriim et al. (153, 130) studied the base composition of RNA TABLE VII BASECOMPOSITION OF RNA’s FROM DIFFERENT PARTSOF O ~ C Y T NUCLEI E DETERMINED ULTRAMICROCHEMICALLY (1.30) Base composition (mol %) Structure

G

C

A

U

(G +C)/(A + U )

Chromosomes Nuclearsaps

20.6 23.2 31.7 27.2 20.4 29.3 30.2

25.2 26.6 28.7 29.5 23.8 30.1 27.5

26.0 23.7 18.1 20.1 26.6 21.5 20.9

28.3 26.5 21.7 23.0 29.3 19.1 21.4

0.84 0.99 1.54 1.31 0.79 1.46 1.36

Material

Triturus

Cytoplasm Chromosomes Triturus viridescens Nucleoli Cytoplasm

1

4 Figures for nuclear sap RNA gave large deviations and in the table the data of only one of preparations is presented.

prepared from microsurgically isolated nucleoli of fixed oocytes and showed that, like r-RNA, it is predominantly of the GC type (Table VII) . As we have already noted, the residue after extraction of nuclei by dilute and concentrated salt solutions consists of membranes and nucleonemata-containing structures: nucleoli and residual chromosomes (29, 3 4 ) . Granules of nucleonemata are sensitive to RNase treatment. Uranyl acetate stains these granules and it can be deduced that, in such saltextracted preparations, RNA is present mainly in the granules of nucleonemata. It has been isolated by the treatment of residues, after salt extraction, by SDS-phenol and it was shown that its base composition and sedimentation behavior correspond to those of pure rRNA (74, 155). From this fact and from the similarity of nucleonemata granules and ribosomes under the electron microscope, one can conclude that

NUCLEAR RIBONUCLEIC ACIDS

301

nucleonemata contain rihosoinal RNA within granules that are ribosomes or their precursors. Analysis of the composition of newly formed nucleonemal RNA synthesized during short pulses of Paashows that it is identical with that of ribosomal RNA (‘74, 157, 158). Thus nucleoneniata contain newly formed R-RNA also (Table VI) . These data are in disagreement with the results obtained with calf thymus nuclei. In those experiments RNA of the nuclear residue contained about 25% of D-RNA in addition to R-RNA ( 6 0 ) . However, thymus nuclei D-RNA do not solubilize during the extraction by dilute salt solutions (25, 26). It is possible that the 30 S particles of thymus, containing mRNA, are more firmly bound to chromosomes. On the other hand, during the concentrated salt extraction these particles may be precipitated and then recovered in a “nucleonemal” fraction. However, it should be stressed that residues from a number of other sources contain no detectable amounts of D-RNA (74). A sedimentation analysis of nucleonemal rapidly labeled RNA was carried out by Busch et al. (159). Heavy R-RNA fractions were detected and nucleonemata were the only nuclear component containing heavy R-RNA. Tamaoki and Mueller (160) have shown that this RNA is present not in a free state but as a ribonucleoprotein. They disrupted nuclei by treatment with deoxycholate and polyvinylsulfate and detected a component sedimenting faster than 70 S ribosomes and containing newly formed RNA. After deproteinization this RNA had a sedimentation coefficient of about 35-40 S. Probably it corresponded to heavy R-RNA (Fig. 9 ) . Nucleoli isolated by mechanical fractionation of nuclear homogenates from various sources contain RNA of a definite GC type and do not contain any significant amounts of D-RNA (53, 128, 156). Muramatsu et al. (159) have shown that nucleoli isolated from liver and tumors contain highly polymerized RNA with the specificity coefficient (G C)/(A U) = 1.5-1.8. For rapidly labeled highly polymerized nucleolar RNA, this ratio is even higher: 1.56-2.00 (152). Thus nucleoli contain both pre-existent and newly formed R-RNA. On the other hand, nucleoli isolated by mechanical methods also contain considerable amounts of tRNA (168, 129). Chipchase and Birnstiel (169) isolated nucleolar RNA from pea seedlings and found that it contains two peaks characteristic of rRNA and a considerable amount of sRNA with a sedimentation coefficient of 4 S. Nucleolar sRNA can bind amino acids and possesses a CCA end that turns over rapidly. Thus i t may be regarded as tRNA (128, 1.29). Nucleolar sRNA is an active

+

+

302

G . P. GEORGIEV

acceptor of methyl groups (see l.)clow). This property is also characteristic of tRNA. Nucleolar RNA is capable of limited hybridization with homologous DNA, competing with ribosomal RNA for binding sites (12’9). These experiments prove that the nucleolus contains mainly R-RNA (including true rRNA) and tRNA. As in the salt fractionation mcthod the iriecliilnical fractionation procedure for the isolation of nucleoli does not exclude a certain degradation of nucleolar RNA and ribonucleoproteins. This is why only certain sources poor in RNases are most favorable for studying nucleolar

700

0.5

500

0.3 400

0.2

300

E

3 TN

200 0.1 100

I

0.3

0

3

6 9 12 15 Fraction number

18

FIG.9. Sedimentation profiles of nuclear lysates after digestion with DNase and treatment by deoxycholate and dextran sulfate. Profile of RNA isolated from the “heavy” zone is shown below (160). RNA and nucleoproteins. Using the preparations of nucleoli from such source pea seedlings Birnstiel et al. (161) were able to isolate not only nondegraded rRNA from nucleoli but also nucleolar ribosomes. This was achieved by treating nucleoli with deoxycholate. Identical sedimentation coefficients and the ability to undergo dissociation and reassociation depending on Mg++ concentration make nucleolar ribosomes isolated by these authors indiscernible from cytoplasmic ribosomes. It is significant that in nucleolar ribosome preparations the amount of 60 and 40 S particles is not equal; 60 S subunits prevail. Nucleolar ribosomes are functionally active; they incorporate labeled amino acids in a cell-free system, although to a lesser extent than do cytoplasmic ribosomes (161, 162). However, this may be due to damage during isolation.

NUCLEAR RIBONUCLEIC ACIDS

303

On the other hnntl, nucleoli and nuclconcniata diow in gcneral an active protein synthesis in vivo (114, 126, 163). Thus the nucleolus as an entity contains ribosomes and tRNA. Upon extraction of the nuclear soluble phase (nuclear sap), tRNA and probably some of the ribosomes (free ribosomes of the nucleolus) are extracted and only nucleonemataattached ribosomes or their precursors remain in the nucleolus. These ribosomes contain the most metabolically active rRNA. Nucleonemata contain also heavy R-RNA, a precursor of true ribosomal RNA, although its exact localization is unknown. One cannot exclude the presence of a certain amount of mRNA in the nucleolus.

E.

Conclusion: The N a t u r e of the RNA of Subnuclear Structures

The results of the characterization of nuclear RNA’s and ribonucleoproteins obtained by the methods outlined above can be summarized as follows. (1) Cell nuclei contain R-RNA, D-RNA, and sRNA. R-RNA of nuclei includes true ribosomal RNA (rRNAA and rRNA,) and heavy R-RNA’s, polycistronic precursors of ribosomal RNA (R-RNA, and R-RNA,). Nuclear D-RNA is informational RNA. At least a part of it can be identified with newly formed mRNA, which is later transferred to the cytoplasm. The nucleus probably also contains a specific fraction of D-RNA functioning a t or near the site of its synthesis and then degrading rapidly. Finally, nuclear RNA contains tRNA and probably a special low molecular weight RNA complexed with histones. (2) Newly formed (rapidly labeled) nuclear RNA is represented by similar molecular species. These include heavy precursors of ribosomal RNA, heavy polycistronic D-RNA, and newly formed tRNA of the cell. (3) The soluble phase of the cell nucleus (nuclear sap) contains ribosomal RNA existing in ribosomes of nuclear sap, free tRNA, and probably a fraction of nuclear mRNA in mRNA-containing 30 S particles. (4) Chromosomes contain mainly informational RNA, existing primarily within 30 S particles. Another part of the chromosomal informational RNA is thought to exist within other complexes not yet characterized. The DNP fibrils of chromosomes contain probably a low molecular RNA linked to histones. Nucleonemata of chromosomes contain some part of R-RNA (ribosomal RNA and its precursors). ( 5 ) The nucleolus contains ribosomal RNA mainly in the granules of the nucleonemata, which are probably ribosomes or ribosomal precursors. The nucleolus also contains the heavy precursors of rRNA in the form of ribonucleoprotein complexes. Thus, the nucleolochromosomal R-RNA is completely recovered as a part of the nucleonemata of the nucleolus

304

G. P. GEOBGIEV

and chromosomes. The nucleolus also contains significant quantities of tRNA.

IV. The Biosynthesis of RNA in Nuclear Structures and Its Transport to the Cytoplasm

A. localization and Mechanism of RNA Biosynthesis 1. THE ROLEOF NUCLEI The active role of cell nuclei in RNA biosynthesis is definitely proved by the results discussed in the preceding sections. The hypothesis regarding the nucleus as a site of cellular RNA synthesis was proposed a long time ago (14). It postulates the transport of RNA from the nucleus to the cytoplasm and there is considerable evidence for its support a t the present time. In the present survey, we mention only briefly some fundamental facts proving the nuclear origin of the main part of cellular RNA. For a more detailed discussion, the reader is referred to corresponding reviews ( 1 , 2 , 5 , 6 , l 4 , 1 5 ) . These fundamental facts are the following: (1) The kinetics of the incorporation of labeled precursors into RNA fractions of the nucleus and the cytoplasm agree well with the suggestion of its nuclear origin and subsequent transfer to the cytoplasm. Similar results have been obtained in autoradiographic experiments and in experiments with isolated cellular components. This is valid both for total RNA and for its subfractions, mRNA. rRNA, and tRNA. (2) Experiments on enucleation and on transplantation of labeled nuclei indicate that, in most organisms tested, the incorporation into cytoplasmic RNA requires the presence of the nucleus. However, this is not the case if a considerable amount of DNA is present in the cytoplasm. (3) The synthesis of all classes of RNA is DNA-dependent. DNA sequences complementary to cellular mRNA, rRNA, and tRNA are found in cell nuclei. (4) The presence of newly formed mRNA, rRNA, and tRNA in nuclei is demonstrated. The nucleotide sequences of these newly formed molecules are identical with the nucleotide sequences of the corresponding RNA classes of the cytoplasm. ( 5 ) Cell-free synthesis of all RNA classes requires either the presence of nuclei or the presence of nuclear DNA-dependent enzymes (RNA polymerase).

305

NUCLEAR RIBONUCLEIC ACIDS

Thus the conclusion about the nuclear origin of the main part of cytoplasmic RNA is well-supported. It should be noted, however, that DNA has been found in certain cytoplasmic structures such as mitochondria or chloroplasts (see Granick and Gibor in this volume, and Iwamura in Volume 5). These organelles may thus be capable of autonomous RNA synthesis, independent of the nucleus. The nature of this RNA is unknown. Perhaps it may serve as a template for the synthesis of the corresponding proteins (for example, mitochondrial) . With this exception, the main bulk of cellular RNA participating in cellular protein synthesis is synthesized in nuclei and the main pathways and mechanisms of these processes are discussed below, beginning with the enzymes of RNA biosynthesis. 2. RNA POLYMERASE

It is firmly established nowadays that RNA synthesis in the cell is catalyzed by an enzyme (or enzymes) called RNA polymerase (RNA nucleoside triphosphate nucleotidyltransferase) and proceeds on a DNA template. RNA polymerase was first demonstrated by Weiss and Gladstone (164)in rat liver nuclei. Later, its wide occurrence among animals, plants, and bacteria was shown [see (166)l. The properties of bacterial RNA polymerases purified several hundred-fold have been studied in detail. The RNA polymerase of E. coli has a sedimentation coefficient of about 20 S and a molecular weight of about 600,000 (166).Treatment by agents such as streptomycin or detergents causes dissociation of the enzyme into subunits with a molecular weight about 100,OOO each (sz0 = 4.1). Fuchs et u2. (167) have studied highly purified preparations of RNA polymerase under an electron microscope and have shown that the six subunits are linked side by side in such a way that the native enzyme appears as a rosette or a cylinder with a height of 95 A, external diameter of 125 A, and internal diameter of 40 A. The enzyme catalyzes the reaction n(pppN) + (pN), npp, where pppN are the various nucleoside 5'-triphosphates. The absolute requirements of the reaction are: presence of all four nucleoside triphosphates (for synthesis of natural RNA), DNA, and divalent cations (Mg++ and Mn"). DNA serves as a template determining completely and unequivocally the nucleotide sequence of the RNA synthesized (168). The latter is complementary to the DNA template (169).The RNA synthesized has a fairly high molecular weight 0.1-2 X 106 (166, 170), of the same order as the molecular weight of cellular RNA. It can serve as a template for protein synthesis on ribosomes (171,17g) and one can

+

306

G. P.

GEORGIEV

suggest that functionally active mRNA is synthesized in the RNA polymerase reaction. Both tRNA and rRNA have also been detected in the products of the reaction (175). Thus, if total cellular DNA is used as a template, RNA polymerase synthesizes all the main types of RNA. This fact, together with the ubiquitous distribution of RNA polymerase and its localization in those organelles where RNA synthesis takes place, enables one to conclude that RNA synthesis in vivo is really accomplished by this enzyme. Furthermore, the antibiotic actinomycin inhibits the RNA polymerase reaction, forming a complex with the DNA devoid of priming activity (174). An analogous situation is observed in vivo; the sensitivity of cellular RNA synthesis to actinomycin is equal to that of the RNA polymerase reaction (175, 176). These data point to the RNA polymerase as the enzyme responsible for RNA synthesis in vivo. Since the synthesis of all types of cellular RNA is sensitive to actinomycin and since DNA contains nucleotide sequences complementary to all types of RNA (17’7, 178),one can conclude that all types of cellular RNA are synthesized by this enzyme. It may well be, however, that different RNA polymerases are used for the synthesis of different RNA’s. The important role of RNA polymerase in RNA biosynthesis demands more detailed consideration of the properties of the reaction.

3. ON THE MECHANISMS OF

THE

RNA POLYMERASE REACTION

The synthesis of RNA in the polymerase reaction proceeds by sequential addition of nucleotidyl residues to the free 3’-OH terminus of the growing polyribonucleotide chain according to the scheme : pppXpY . . . pZ pppN + pppXpY . . . pZpN pp (17‘9,180). The mode of participation of the template in this reaction is of obvious interest. It is firmly proved at the present time that doublestranded native DNA serves as a template in vivo. I n cell-free systems from different sources, both single- and double-stranded DNA’s are active as primers. If DNA is double-stranded, RNA synthesis proceeds through a conservative mechanism; if single-stranded DNA is used, the mechanism is semiconservative. I n this latter case, the formation of a double-stranded DNA-RNA complex takes place a t first and then, during the synthesis, the pre-existing RNA of the complex is forced out by the newly formed RNA (181). This complex is not formed when doublestranded DNA is used as template and DNA is not altered in the course of the reaction (154, 166) (Fig. 10). Long double-stranded DNA-RNA complexes were not found in vivo (182,185).

+

+

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On the other hand, RNA synthesis in the cell is extremely sensitive to actinoniycin, which has maximal affinity to native but not to singlestranded DNA (184). Finally, RNA polymerase isolated from some animal sources can use only double-stranded DNA as a template in vitw (185). Thus, double-stranded DNA most probably serves as the teiiiplate in vim. However, although double-stranded DNA participates in RNA biosynthesis, only one of its chains serves as a template in the strict sense of the word. It results in the synthesis of RNA complementary to one chain of DNA. This phenomenon is designated as asymmetric RNA synthesis or asymmetric transcription. RNA

polymerase

Active center

rrrn-ri -5

DNA RNA

FIG.10. The role of template in the RNA polymerase reaction. (a) and (b) Two stages of RNA synthesis on a single-stranded DNA template (semiconservative mechanism), (c) RNA synthesis on double-stranded DNA (conservative mechanism). (d) Model suggested by Butler, illustrating the possibility of local unwinding of double-stranded DNA (216).

The most definite results were obtained using phage DNA where in some instances complementary chains can be separated on a preparative scale. All RNA synthesized in the bacterial cell after phage infection can hybridize with only one of the two chains of phage DNA. It is inferred that during phage infection only one chain is transcribed (186‘189). In cells of bacteria and higher organisms, the preparative separation of DNA complementary chains is much more difficult but some indirect data also give support to the asymmetric mechanism of transcription. All highly polymerized cellular RNA’s including mRNA, consist of single-stranded molecules and cannot hybridize with themselves. That is, they do not form double-stranded RNA molecules on annealing.

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Consequently the population of cellular RNA molecules does not contain chains complementary to each other (68, 96). If bacterial DNA is hybridized with a large excess of RNA, not more than 50% of the DNA can be forced into the complex (96). Thus asymmetric transcription is probably a characteristic of all organisms. The basis of the asymmetric character of transcription remains obscure. RNA synthesis in a cell-free system may be either asymmetric or symmetric depending on conditions. Hayashi et al. (189) observed asymmetric RNA synthesis using circular phage DNA as a template. After the breakage of rings by sonication, both chains of the DNA were copied. Somewhat later, however, asymmetric synthesis of RNA was observed with noncircular templates also (190-192) and only denaturation rendered the synthesis symmetrical. Thus the factors determining transcription of the DNA are inherent in its structure. This is probably connected with the predetermination of the starting points of transcription, which may be singlechain breaks, some unique nucleotide sequences, or other factors. In order to explain the asymmetric complementary synthesis of single-stranded RNA on the double-stranded DNA, it is suggested that a local separation of two DNA strands takes place during the synthesis but is restored immediately after it. This wave of DNA local denaturation should run along all the DNA molecule copied (see Fig. 10).

B. mRNA Biosynthesis in Chromosomes: Main Stages In this section problems concerning localization of mRNA synthesis and the main stages of the RNA polymerase reaction are discussed. 1. LOCALIZATION OF BIOSYNTHESIS OF INFORMATIONAL RNA

Autoradiographic studies of RNA synthesis not only proved its nuclear localization but also permitted an attack on the problem of the role of different intranuclear structures in RNA biosynthesis. In animal cells, RNA precursors are incorporated primarily into chromosomes (193, 194) , which proves the participation of chromosomes in RNA synthesis. A number of facts indicate that informational RNA is synthesized in chromosomes. It has been noted already that chromosomal RNA is D-RNA. Rapidly labeled D-RNA is also localized in chromosomes. Lerman et al. (73) have shown that small doses of actinomycin selectively inhibit the synthesis of R-RNA. RNA, synthesized in the presence of small doses of acetinomycin, consists exclusively of D-RNA (175, 195, 196). I n autoradiographic experiments similar doses of actinomycin completely suppress the incorporation of labeled precursors in nucleolar RNA and inhibit chromosomal RNA biosynthesis only slightly (45, 7 8 ) . One may conclude that all newly formed D-RNA is localized

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exclusively in chromosomes but not in the nucleolus. Embryonic cells having no nucleoli u p to the gastrula stage, as well as anucleolar mutants, do not synthesize R-RNA while D-RNA synthesis proceeds normally (81, 181, 197). It should also be noted that the bulk of RNA polymerase is found in chromosomes (see below). Autoradiographic experiments show that even a t very short incubation times newly formed chromosomal RNA is scattered throughout the nuclear chromatin except for condensed chromatin regions (198). mRNA is thus synthesized by various regions of chromosomes that do not form any particular associations. Newly formed chromosomal D-RNA differs from other cellular RNA classes in respect to behavior during deproteinization. Its solubiliaation demands either the liberation of DNA or heat treatment (see above). This is probably due to a peculiar character of its association with a chromosomal nucleoprotein. Therefore, the study of interrelation between template, enzyme, and product during RNA biosynthesis is of obvious interest. These points are discussed below.

EXISTENCE OF A COMPLEX BETWEEN TEMPLATE, ENZYME, PRODUCT IN THE RNA POLYMERASE REACTION The search for hybrid complexes of DNA and RNA containing one ribo- and one deoxyribopolynucleotide chain has given negative results in both animal (183) and phage-bacteria systems (182).This is natural if we recollect that RNA synthesis in vivo occurs via a conservative mechanism (see preceding section). The scheme of Fig. 10, however, implies the existence of a complex containing double-stranded DNA and single-stranded RNA linked by the enzyme. Indirect bits of evidence for the existence of complexes of this type have been known for a long time. Bonner e t al. (199) have found that the newly formed RNA of pea seedling nuclei is bound to DNA and is rather resistant to RNase action. As judged from the data on complex thermal dissociation, which occurred a t 50-60”C a t any ionic strength investigated, the authors came to the conclusion that RNA is bound to DNA by a protein “linker.” These data may be compared with the fact that chromosomal D-RNA can be solubilized by phenol treatment also only a t 60°C or higher (47,

2. ON

THE

AND

48)* A complex containing RNA, double-stranded DNA, and possibly protein has been isolated from Neurospora crassa by Schulman and Bonner

(ZOO). Interesting facts were obtained from studies of distribution of thc RNA polymerase. A considcrable portion of this enzyme in bacteria is

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bound to DNA, and for its isolation a polymerase-DNA complex must be dissociated, for example, by protamine treatment (165, 1'71). A considerable part of RNA polymerase activity during ultracentrifugation or electrophoresis of bacterial extracts transfers together with DNA although they have different sedimentation constants or electrophoretic mobilities (201). I n cells of higher animals, RNA polymerase is also tightly associated with DNA. After treatment of nuclei with 0.4 M NaCI, which extracts nuclear sap, Weiss (202) found RNA polymerase in the nuclear residue, consisting of chromosomes and nucleoli. RNA polymerase goes into solution if chromosomal deoxyribonucleoprotein is solubilized by 1 M NaCl and precipitates if D N P is precipitated by dilution (203). Huang and Bonner (204) dissociated chromosomal deoxyribonucleoprotein from pea seedling nuclei by treatment with 4 M CsCI. This separates DNA and protein linked to DNA by salt linkages. After ultracentrifugation, DNA sedimented a t the bottom while proteins floated a t the top of the tube. After this procedure, only about 4% of all proteins of the nucleoprotein remain bound to DNA. However, a significant part of the RNA polymerase remains firmly bound to DNA. Thus in animal, plant, and bacetrial cells a considerable part of RNA polymerase is complexed with DNA, and some newly formed RNA is also bound to DNA. The nature of this interaction has been studied in cell-free systems. Bremer and Konrad (17'0) synthesized RNA by a purified RNA polymerase, using DNA from T4 phage as a template. The products of the reaction were studied in a sucrose gradient, in which DNA, newly formed RNA, and RNA polymerase sediment as a single peak. DNase treatment leading to DNA degradation destroys the complex. Deproteinization by dodecylsulfate treatment results in the separation of DNA from RNA. Thus it is no true DNA-RNA hybrid but rather DNA and newly formed RNA held together by the enzyme. If ribosomes are added to such a system, then the formation of the complex cont.aining DNA, RNA polymerase, synthesized RNA, and also ribosomes attached to newly formed RNA takes place (206),as is also shown by the electron microscope, which shows that only one end of the RNA chain is bound in a complex while the rest of the molecule is free and may interact with ribosomes. It should be noted that the newly formed RNA itself cannot be liberated spontaneously from the complex with the enzyme and template in witro (154, 180). A simple method for the detection of complex formation consists of using DNA gels cross-linked by UV irradiation (206). Thin films of native or denatured DNA obtained by drying DNA on smooth glass surfaces were irradiated by UV light and washed free of non-bound DNA,

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after which pieces of the swelled gel were placed in RNA polymerase solution. After repeated washing of unadsorbed enzyme, the gel was placed in the mixture of nucleoside triphosphates and the amount of bound enzyme was assayed by the incorporation of label into RNA (Fig. 11). Under these conditions one can easily observe the formation of a triple complex between enzyme, template, and product. This complex is dissociated by dodecyl sulfate treatment. If gels prepared from denatured

FIG.11. Experiments with insoluble DNA gels demonstrating complex formation between enzyme, template, and product in the RNA polymerase reaction (206). Columns a t the right show the distribution of newly formed RNA between different fractions during washing by buffer and sodium dodecyl sulfate. (SDS = sodium dodecyl sulfate; TCA = trichloracetic acid).

DNA are used, then dodecyl sulfate treatment does not affect a considerable part of RNA, which remains bound in the gel. This RNA is probably a component of the double-stranded DNA-RNA hybrid. The conveniences of the system described are the ease of template removal (since the gel can be transferred simply by a glass rod) and the relatively direct character of the information obtained. These experiments clearly demonstrate the formation of the postulated complex in the course of the RNA polymerase reaction. However, such soluble complexes have not been obtained from cell nuclei but the indirect evidence listed above indicates that they may

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exist in vivo. It may well be that the extraction of newly formed chromosomal D-RNA only a t temperatures exceeding 60°C is explained by the existence of such complexes. However, one should not forget that, besides RNA, chromosomes contain histones, and chromosomal RNA may be complexed with histones. I n model experiments, phenol treatment of artificial ribonucleohistone leads to complex dissociation only a t about 50-60°C (185). Further experiments are necessary in order to resolve the question as to the special behavior of chromosomal D-RNA during thermal fractionation. The formation of a complex between DNA, RNA polymerase, and the product of the reaction in the courge of RNA synthesis suggests three stages in the process (164): (1) complex formation between RNA polymerase and template DNA: DNA+E+DNA.E;

(2) RNA synthesis in the strict sense-after each nucleotide addition to a growing chain, the enzyme moves along the DNA one nucleotide pair; this process results in a stepwise transcription of one of the DNA strands: DNA*E+ n(pppN) + DNA*E-RNA;

(3) liberation of newly formed RNA from the complex (detachment of RNA or dissociation of the complex to the components) after completion of RNA synthesis: DNA*E.RNA+ RNA + DNA + E (or RNA + DNA-E)

The two first stages are connected with RNA biosynthesis and are discussed in the next section. The third stage is discussed in the section devoted to the transport of mRNA. 3. INTERACTION OF DNA WITH RNA POLYMERASE The first stage of RNA synthesis, a formation of the complex between DNA template and the enzyme, is probably of extreme importance for the regulation of RNA biosynthesis. It has already been noted that, in the cell, RNA polymerase exists as a complex with DNA or DNP. On the other hand a considerable amount of RNA polymerase is found in the soluble phase and is not bound to DNA (185, 207-909). Free RNA polymerase, by all its properties, is indistinguishable from the bound enzyme (210).Thus free and bound RNA polymerases exist in equilibrium, which may be regulated by the forces affecting the first stage of the reaction: formation of the enzyme-template complex.

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313

The requirements of complex formation have been studied in vitro. For this purpose, template and enzyme were mixed in the absence of labeled precursors and unbound enzyme was removed either by ultracentrifugation or by simple washing of insoluble DNA gels (211, 206). The amount of bound enzyme was assayed by transfer of the enzymetemplate complex to a substrate mixture containing labeled RNA precursors. It was shown that the first stage of the RNA polymerase reaction, formation of an enzyme-template complex, may take place in the cold and in the absence of substrates, energy sources, and even divalent 211, 212). cations (~?06, An essential role in complex formation between DNA and polymerase is probably played by electrostatic interaction forces. This is suggested by dissociation of the complex in neutral salt solutions of high ionic strength ( ~ 2 or ) in the presence of bases (e.g., streptomycin sulfate) ( 2 0 6 ) . The formation of the complex is relatively unspecific since RNA polymerase may form complexes with nucleic acids other than DNA, although these complexes are less stable. This latter circumstance probably explains the inhibition of the RNA polymerase reaction if RNA is added to the reaction mixture before DNA (211). Although RNA polymerase cannot sharply distinguish the type of the nucleic acid used, the binding of the enzyme with the template cannot be explained only by nonspecific electrostatic interactions. This is supported by the fact that the number of sites where the enzyme can bind to DNA is limited. Bremer et al. (see 164) have measured the maximal number of RNA chains growing on a DNA template and have calculated that only about fifty molecules of the enzyme can bind to DNA with a molecular weight of about lo8. This corresponds to one molecule of enzyme per DNA length of 2 X lo5, which is close to the size of the cistron. The maximal amount of enzyme that can be placed on DNA is ten times greater than this value, and this tight packing is really observed if denatured DNA is used. Naturally the question arises as to the nature of the DNA sites responsible for the interaction with the polymerase. There are several more or less plausible hypotheses. (1) The first possibility is the binding of the enzyme to DNA regions containing free 3? OH ends, suggested by the fact that a complex of DNA and RNA polymerase cannot be a primer in the DNA polymerase reaction (212). This may mean that the free 3’ OH groups necessary for the addition of deoxynucleotides are blocked. It would hardly be observed if the interaction of DNA and RNA polymerase was unspecific. Moreover, the complex of DNA with RNA polymerase is more resistant towards exonucleases liberating 5’ deoxynucleotides while

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its sensitivity towards endonucleases is not altered as compared with that of free DNA (212). It is interesting to recall, in this connection, that aminoacyl-tRNA inhibits the RNA polymerase reaction to a lesser degree than free tRNA (213). It may well be that esterification of the 3’ OH terminal group in a tRNA molecule inhibits its interaction with RNA polymerse and therefore decreases the competition of tRNA and DNA for the enzyme. A question arises as to whether free 3’ OH terminals really exist in the native DNA molecule. The first possibility is that single breaks may exist in one of the DNA chains while the complementary chain remains continuous, or the breaks in this second chain may be masked by polypeptide bridges. A small amount of amino acid is found reproducibly even in the most purified DNA preparations. On the other hand pronase, hydroxylamine, and phosphoamidase lower the molecular weight of DNA preparations and the content of amino acids (214, 2 1 4 ~ )The . suggestion of breaks in one chain of DNA could also explain the asymmetry of RNA synthesis. (2) One may also suggest that RNA polymerase recognizes certain nucleotide sequences in double-stranded DNA. It has been shown, using P,y-labeled nucleoside triphosphates, that RNA chains synthesized on native DNA templates begin almost exclusively with purine nucleotides. It is inferred that the enzyme should interact with the DNA chain having a pyrimidine nucleotide terminus. If denatured DNA is used, this phenomenon is less pronounced (179). (3) Finally, it has been suggested that local denatured regions of the DNA double helix may be attachment sites for the enzyme (164). This is indirectly supported by the greater affinity of RNA polymerase for denatured DNA (164,166). Further clarification of the nature of enzyme attachment sites, corresponding probably t o the starting points of transcription, is of utmost importance for the understanding of regulation of RNA synthesis in chromosomes since the availability or blocking of these sites will determine if a given gene can be transcribed. If nucleoside triphosphates and Mg++ions are present in the medium, complex formation between DNA and a polymerase results in the initiation of RNA synthesis. The elongation of RNA chains in the cell-free system in the course of the reaction can easily be observed. If RNA is isolated a t different times after the beginning of the reaction, one can see that the sedimentation coefficient and consequently the molecular weight of the RNA increases while the number of molecules synthesized does not (170).This observation is in good agreement with the con-

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cept that, after attachment to the template, RNA polymerase moves along the DNA chain and transcribes it. The necessity of enzyme movement follows from the fact that the enzyme transcribes DNA chains that are one or two orders of magnitude longer than the diameter of the polymerase ( 1 6 7). In the cell-free system, the rate of the synthesis is about two to threc RNA nucleotides per second. It corresponds to an enzyme movement of about 500 A per minute and the synthesis of an RNA chain with a molecular weight about 50,000 per minute. However, the rate of RNA synthesis in vivo may be different. I n some experiments on mRNA synthesis in bacteria undergoing induction, the values for the rate of RNA formation are of the same order as in cell-free systems (215). Unfortunately, there are no data on the rate of RNA synthesis in cells of higher animals. Generally it may vary considerably depending on conditions. Many aspects of the detailed mechanism of the RNA polymerase reaction remain obscure, in particular the mechanism of local unwinding of double-stranded DNA. One of hypothetical schemes of interrelations between the enzyme and the template is presented in Fig. 10 (216). This scheme postulates both unwinding and subsequent rewinding of double-stranded DNA molecules as caused by the rotation of the enzyme during its passage along it. Data obtained with the electron microscope indicate that DNA may well pass through the internal channel in the cylindrical molecule of polymerase formed by its six subunits (167). Synthesis of RNA in the cell-free system ceases after a relatively short period of time and can be restored only by the addition of more enzyme. This is explained by the inability of the complex of DNA with the polymerase and RNA to dissociate in vitro (170). This problem is discussed in more detail in the section concerned with mRNA transport. I n conclusion, we would like to stress that, although many facts are established only for the bacterial enzymes, they may well be of general nature. The next section is devoted to the problem of the liberation of newly formed mRNA from the chromosomal complex and of the transport of mRNA to the cytoplasm.

C. Transport of mRNA 1. MRNA AND MRNA-CONTAINING NUCLEOPROTEINS OF THE CYTOPLASM Information concerning mRNA and mRNA-containing structures in the cytoplasm is absolutely necessary for studies of the fate of mRNA synthesized within chromosomes. The occurrence of mRNA in the cytoplasm was demonstrated soon after its observation in cell nuclei, when

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Hoagland and Askonas showed that the postmicrosomal fraction of cytoplasm contains RNA capable of stimulating amino acid incorporation into isolated ribosomes. This RNA had sedimentation coefficients of about 6-14 S (217). Stimulation of amino acid incorporation by cytoplasmic RNA was also observed by others (93-95). Pulse-labeled cytoplasmic RNA can be effectively hybridized with DNA, but a t longer labeling times this property is lost (91). Hybridizability with DNA means that a given RNA is complementary to a considerable number of cistrons and this is typical of mRNA. Penman (et al. isolated pulse-labeled RNA from polysomes and found that this RNA sedimenting in the region 6-14 S has a sedimentation behavior different from that of ribosomal RNA. The base composition of this RNA differed from that of ribosomal RNA, but it did not correspond exactly to the base composition of DNA (218). The cytoplasm of early embryos also contains several newly formed RNA components with sedimentation constants of 6-35 S, which are not identical to ribosomal RNA species ($19, 197). These RNA's cannot also be rRNA precursors since early embryos do not synthesize rRNA a t all. Recently i t has been shown that these RNA components can be effectively hybridized with DNA (220) and therefore they may be regarded as cytoplasmic mRNA. I n order to detect cytoplasmic mRNA, Samarina used selective suppression of R-RNA synthesis by small doses of actinomycin. Under these conditions, practically all newly formed cellular RNA is DNA-like, and studying its distribution and properties one can obtain useful information concerning the content of D-RNA in various subcellular structures. This method permitted the demonstration of the presence of D-RNA not only in the nucleus but also in the cytoplasm of the animal cell (99, 196). Thus cytoplasm contains a DNA-like RNA capable of hybridization with DNA and stimulating amino acid incorporation in cell-free systems of protein synthesis. All these facts indicate that the cytoplasm of animal cells really contains mRNA, although it remains to be proved that all these three properties belong to the same RNA. Brawerman and Hadjivassiliou have observed that D-RNA can be dissociated from template RNA (221);however, these data should be thoroughly checked. First of all, the dissociation is not complete and therefore it may arise from different specific template activities of different D-RNA's. It has been shown also that certain types of R-RNA, in particular precursors of ribosomal RNA, can stimulate amino acid incorporation into ribosomes (222). The existence of mRNA in the cytoplasm is thus proved, although

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its exact molecular characteristics are not yet established. Most authors agree that its mean sedimentation constants lie within the range 1216 S, and 8 mean molecular weight determined from these constants is of the order of 0.3-0.5 x 106 (99, 618, 2.23). These values were obtained in experiments on the distribution of labeled RNA after short radioactive pulses using rat liver and some other cell types. The selective suppression of R-RNA synthesis by actinomycin has been used in some experiments. The values so obtained are in good agreement with the data on the size of polysomes in HeLa or liver cells. Polysomes containing ten to fifteen ribosomes are most numerous, this size corresponding to a molecular weight of about 0.3-0.45 x lo5 for mRNA contained in them (2.24, 225). Other authors who used template activity in the Nirenberg system as a test for mRNA identification reported somewhat higher sedimentation constants and molecular weights for these RNA species. I n their experiments, the sedimentation constant was about 18 S and the corresponding molecular weight about 0.6 X lo6 (107, 108). The explanation of these differences is not yet clear, but both sets of data give szo values for cytoplasmic mRNA that are very close to the sedimentation constant of the main peak of chromosomal D-RNA and considerably lower than the sedimentation constants of newly formed chromosomal D-RNA (68, 79, 80). Thus, polycistronic mRNA synthesized in chromosomes must be split a t or near the site of its synthesis and then transported to the cytoplasm. Migrating RNA having a molecular weight 3 to 4 times lower than newly formed chromosomal RNA may be monocistronic (Table 111). Identity of part of the chromosomal D-RNA and cytoplasmic mRNA is shown by competitive relations between them in hybridization experiments (84, 266). The finding of mRNA in cytoplasm together with its complementarity with DNA and the identity of its base sequences with those of newly formed chromosomal mRNA prove the existence of the transport of mRNA from chromosomes to the cytoplasm. However, these facts do not yield any information about the actual mechanisms of this transport. In order to attack the problem of the transport mechanisms, the nature of mRNA containing nucleoproteins from various subcellular fractions has been studied. Hoagland and Askonas (917) found that a part of the activity stimulating amino acid incorporation is present in a fraction of rat liver homogenate sedimented for 8 hours a t 105,000 g, that is, in particles lighter than ribosomes. The main part of the RNA having template activity and a DNA-like composition could be detected in polysomes. Spirin (227, 228) described in embryonic cells a special class of particles containing mRNA. These particles were called

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“informosomes.” Informosomes contain several components with discrete sedimentation coefficients, and each component contains an RNA with a definite molecular weight (98). Ultracentrifugation in CsCl density gradients gives RNA contents in informosomes of about 2540%. It has been suggested that informosomes are a form of conservation of mRNA when it does not participate in protein synthesis. Informosomal protein possibly makes mRNA inactive in protein synthesis. Informosomes were also isolated from polysomes (227). Studying the distribution of D-RNA in rat liver cytoplasm under the conditions of partial actinomycin block, Samarina (99, 196) has shown that about three quarters of the cytoplasmic mRNA can be detected in

Qi

I

Zu oH t

I000

i

400 ;2.0

E

u 500 200 ; 1.0

10 20 30 Fraction number (0)

40

10

20

30

40

Fraction number (b)

FIQ. 12. Sedimentation profile of cytoplasmic extracts from HeLa cells after labeling RNA for 30 minutes and 2 hours. Profile of RNA isolated from 45 S peak is shown above (2.48).

polysomes of the endoplasmic reticulum while the remaining one quarter can be found in particles remaining in the supernatant after 2 hours a t 105,000 g but sedimenting a t higher speeds (180,000 g for 2 hours). It should be noted that, even in the absence of actinomycin, the newly formed RNA contained in these particles has a DNA-like composition. The specific activity of RNA from these particles exceeds considerably the specific activity of RNA isolated from other cytoplasmic structures, for example, from polysomes of the endoplasmic reticulum especially at short labeling times. At longer labeling times, this difference decreases, indicating perhaps a precursor-product relationship between mRNA’s of these structures. Thus light particles may be suggested as the transport form of mRNA. These particles have been studied in detail using ultracentrifugation in sucrose gradients (Fig. 12) . Their sedimentation con-

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stants are about 4045 S and they contain .mRNA as evidenced from their template activity and hybridizability with DNA (149, 150). These particles may interact with polynucleotides, in particular with polyuridylic acid (149).Besides D-RNA, they perhaps contain R-RNA as judged from the sedimentation analysis and determination of base composition. RNA moieties of these particles have sedimentation constants about 18 S. Similar particles were found after infection of HeLa cells by vaccinia virus. In this case, the 40 S particles contained viral mRNA and possibly an 18 S rRNA (148). It is suggested that the 40 S particle is a complex between mRNA and a minor ribosomal subunit. However this is not proved and it is possible that the 45 S peak consists of both ribosomal subunits and mRNA-containing particles. The relation of nuclear mRNA protein particles, informosomes, and 4 0 4 5 S particles is a t present still obscure. Recently, Perry ( 2 2 8 ~characterized ) 40 S particles by CsCl gradient centrifugation and observed these particles to possess a lower density than small ribosomal subunits. They were interpreted as subunit precursors on the basis of “chase” experiments. This work, however, leaves open the question about the binding of mRNA to these particles. Thus in the cytoplasm of animal cells, mRNA is incorporated into special ribonucleoprotein particles (40-45 S particles) of yet unknown nature. These particles may be a transport form of mRNA. 2. ON THE MECHANISMS OF MRNA SEPARATION FROM AND ITSTRANSPORT TO THE CYTOPLASM

THE

TEMPLATE

This section is concerned with the nature of factors involved in separation of mRNA from the template and its transfer to the cytoplasm. Attempts to study the process of separation of mRNA from the template were undertaken using both whole cells and cell-free systems. Experiments in vivo show that, if RNA synthesis after pulse-labeling is suppressed by actinomycin, only a limited transfer of newly formed RNA from the nucleus to the cytoplasm may take place (75, 229, 230). Predominantly R-RNA is transferred while the main part of D-RNA remains bound in chromosomes a t the site of its synthesis. Thus blocking D-RNA (and consequently mRNA) synthesis inhibits its transport. This result may not be explained by a specific action of actinomycin since another agent (UV radiation) that causes a block in RNA synthesis also inhibits RNA transport. Thus it seems probable that transport of mRNA depends on the continuing synthesis of the latter (2.90).On the other hand inhibition of protein synthesis by puromycin does not affect RNA transport in the cell. Consequently, suppression of RNA synthesis

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inhibits its transport directly and not via the inhibition of protein synthesis (230). Since actinomycin binds to DNA a t any site containing deoxyguanylic acid, it inhibits not only initiation of RNA synthesis but also its growing, creating a hindrance for movement of RNA polymerase along template polydeoxynucleotide chains. If we suggest that separation of mRNA from the template may occur only a t the end point of transcription, then it becomes clear why an inhibition of mRNA synthesis results in blocking its separation: uncompleted RNA molecules are fixed in chromosomes. Thus experiments on the inhibition of RNA synthesis in vivo permit the suggestion that the first stage of RNA transport, its separation from the template, may occur a t end points of transcribed regions. Attempts to study this process in vitro have given negative results. It has been already noted that, in the RNA polymerase system, further synthesis stops after completion of RNA chains. Addition of more enzyme restores the reaction (154). Thus, in vitro, one molecule of enzyme can probably be used only once for the synthesis of one RNA molecule. If polymerase is present in excess, the amount of RNA synthesized exceeds the amount of primer DNA, but the liberation of RNA does not take place, The liberation of RNA probably demands some special factors absent in a cell-free system. Although their nature is unknown, several possibilities are discussed. a. Ribonuclease. Addition of minute amounts of ribonuclease to a cell-free system liberates RNA completely from the complex, although it is accompanied by some RNA degradation (206'). It is known that cell nuclei of various origins contain ribonuclease [see (671. The action of ribonuclease is naturally unspecific, but one may suggest (although it seems not very probable) that RNase may liberate RNA from the complex if it is immobilized a t certain regions of DNA within the deoxyribonucleoprotein complex. b. Ribosomes. Nirenberg et al. (205) demonstrated the possibility of a participation of ribosomes in the detachment of mRNA from the template. They showed that ribosomes may attach to newly formed RNA complexes with the template and enzyme. Such complexes can be seen under the electron microscope (205). However, the attachment of ribosomes to newly synthesized RNA does not result in a liberation of the latter from its complex with DNA. Moreover, in vivo monosomes as a rule do not contain intensively labeled mRNA and are labeled later than polysomes. Thus the participation of ribosomes in the detachment of mRNA from the complex seems doubtful.

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32 1

c. Small Ribosomal Subunits. Another hypothesis suggested by our laboratory (76, 146) and by Henshaw et al. (149) postulates that detachment of mRNA from the template is accomplished not by whole ribosomes but by their small subunits. Then the latter transfers mRNA into cytoplasm where, after its binding with large subunits, the center of protein synthesis is formed. This hypothesis is in accordance with a group of indirect facts. Among them are the existence of two different channels for the synthesis of smaller and larger ribosomal RNA components and ribosomal subunits, the latter being bound only in cytoplasm (see below). The mRNA binding site is located on small ribosomal subunits. Moreover, isolated small subunits have greater affinity to mRNA as compared with the whole ribosomes (149) and they interact a t lower Mg++ concentrations. It is possible that the 45 S particles of cytoplasm might be the complexes of mRNA with precursors of ribosomal subunits or with ribosomal subunits themselves. However recent results indicate that nuclear mRNA containing ribonucleoproteins does not contain ribosomal RNA. These data do not exclude the possible role of small subunits in mRNA transfer but it seems more probable that their participation is limited by the late stages of transport and a t the first stage (liberation of mRNA from the enzyme-template complex) the specific nuclear 30 S particles are involved. d. SO S Particles. As mentioned above, the main form of mRNA in the cell nucleus is as a specific nucleoprotein particle with a sedimentation constant of 30 S. These particles differ from ribosomal subunits in that they have a base composition of the AU type, excluding the presence of ribosomal RNA. Also the buoyant density in CsCl after formaldehyde fixation of 30 S particles is lower than the density of ribosomal subunits (and of informosomes also). The fact of specific interaction of mRNA with these particles may be connected with their participation in the liberation of mRNA from the chromosomal complex. The relation between 30 S particles, on the one hand, and informosomes, on the other, is of obvious interest. One may suggest that, in most cells, mRNA from the chromosomal complex incorporates into 30 S particles and migrates to the cytoplasm where it immediately becomes a component of polysomes and serves as a template for protein synthesis. I n embryonic cells where a large amount of mRNA is conserved it may exist as another complex-informosomes, inactive as templates but protected from degrading enzymes. Further study of this question may help us to understand the intricate mechanisms of mRNA transport in more detail.

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e. Other Possibilities. It should be taken into consideration that 30 S particles might react with the liberated mRNA but that liberation of mRNA from the template occurs via some other mechanisms, for example by RNase. Finally, it seems very probable that mRNA liberation and transport consists of several steps. In this case, different mRNA-protein complexes may exist in the cell, including 30 S particles of chromosomes (first step) and complexes of mRNA with small ribosomal subunits (second step). One can see, however, that now the problem of mRNA liberation and transport is quite obscure and only some speculations may be suggested. Concluding this section, we summarize the data concerning the interrelations of different fractions of informational RNA. Template RNA in the animal cell is relatively stable (104-106, 231). Blocking the synthesis of RNA influences neither the in vivo incorporation of amino acids in proteins nor the stimulating activity of cellular RNA in in vitro experiments for several hours or days. The cytoplasmic mRNA has a distribution maximum in the 1 4 1 8 S zone under ultracentrifugation. Similar sedimentation properties (maximum of distribution 4 1 8 S) are characteristic of the main component of chromosomal D-RNA. The latter, as was pointed out above, consists of molecules with base sequences identical with or similar to cytoplasmic mRNA. This fraction of chromosomal D-RNA is metabolically stable. One may consider that heavy polycystronic mRNA is synthesized in chromosomes. Part of it is further split into shorter chains (distribution maximum about 18 S) included in 30 S ribonucleoprotein particles, transported in this state to sites of protein synthesis, and formed into polysomes. I n polysomes, mRNA is stable for periods of from several hours to several days. Another part of the heavy D-RNA synthesized in chromosomes functions in the place of its synthesis and rapidy breaks up. This labile nuclear D-RNA may be, for example, the messenger RNA for regulatory proteins of the deoxynucleoprotein complex (see Section IV,E) . It was shown that almost the only protein fraction whose biosynthesis is sensitive to an actinomycin block of RNA synthesis is the histone and nonhistone protein of the deoxynucleoprotein complex (232, 233).

D. Synthesis and Transport of Ribosomal RNA and tRNA 1. LOCALIZATION OF RIBOSOMAL RNA BIOSYNTHESIS

The data indicating the presence of ribosomal RNA in nucleoli or, more exactly, in nucleolar and probably chromosomal nucleonemata have

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already been mentioned above. Newly formed nucleolar RNA also contains R-RNA. Experiments on salt fractionation of isolated nuclei show that after short labeling about 8&90% of newly formed R-RNA is found in the nucleonemata of nucleoli and chromosomes. This RNA has the highest specific activity compared to other fractions of nuclear RNA (74, 78, 133). These facts corroborate the suggestion of the nucleolar origin of ribosomal RNA, but additional facts are necessary to prove it. Numerous autoradiographical studies on the kinetics of incorporation of labeled precursors into RNA of chromosomes and nucleoli support the hypothesis of an independent synthesis of ribosomal RNA in the nucleolus (2S4-%%). Nucleoli were shown to be responsible for a considerable part of cellular RNA synthesis since the destruction of nucleoli by a UV microbeam causes a 70% decrease in the incorporation of labeled precursors into cytoplasmic RNA (236, 237). Direct experiments with cell-free systems have shown that isolated nucleoli contain RNA polymerase and are capable of RNA synthesis in vitro (238-240). Thus the nucleolus seems to be not the site of RNA accumulation but rather the site of its synthesis. The question of the identity of the RNA polymerases of chromosomes and nucleoli remains unsolved although they are indistinguishable by several criteria (238). To clarify further the nature of the RNA synthesized in nucleoli, the sensitivity of the synthesis of different RNA classes to antimetabolites was compared. Actinomycin D in small concentrations inhibits selectively the synthesis of R-RNA without any effect on D-RNA synthesis (73-76). In radioautographical experiments, small doses of actinomycin inhibit selectively only the synthesis of nucleolar RNA (46, 78, 241). Consequently the nucleolus can be regarded as a t least one of the sites where R-RNA is synthesized. It is interesting in this connection that cells lacking nucleoli do not synthesize ribosomal RNA. This is true for early embryos and also for anucleolar mutants a t later stages of development (81). I n neither case did the sedimentation profiles and base compositions of newly formed RNA's reveal the presence of either ribosomal or heavy R-RNA. All the newly formed RNA consisted entirely of D-RNA. The beginning of R-RNA synthesis coincides with the appearance of nucleoli (197). ' Of course these experiments are not definitive since one may suggest the synthesis of RNA outside the nucleolus and its subsequent transfer to the nucleolus. This is why experiments on localizing DNA complementary to R-RNA within the nucleus are of extreme importance. Zbarsky e t al. (242) isolated the substance of nucleonemata using a salt fractionation method. The base composition of a small DNA fraction remaining in the residue after extraction by a concentrated salt solution

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differed from the composition of bulk cellular DNA, being more rich in GC pairs. (G C)/(A T) was about 1.00-1.1 in this fraction instead of 0.7-0.8 for total DNA. The authors concluded that their DNA preparations were enriched with DNA complementary to R-RNA. A special case is the DNA of the fungus Blastogladuella emrsonii, which contains much DNA of the GC type. A considerable portion of this DNA is concentrated in nucleolus while another part of the GC DNA is found in the chromatin. On the other hand, the nucleolus does not contain the common DNA of the AT type representing the main fraction of chromosomal DNA (243). It may well be that GC-rich DNA is a template for rRNA of this fungus, and is concentrated in the nucleolus. A number of authors have studied the distribution of DNA capable of hybridization with rRNA between chromatin and nucleolus. Chipchase and Birnstiel (1.29) have shown that in pea seedlings this DNA fraction, corresponding to 0.2-0.3% of the total cellular DNA, is uniformly scattered throughout the nucleus and is not concentrated in nucleoli. On the other hand, experiments with HeLa cells have shown that concentration of this DNA in perinucleolar chromatin is considerably higher than in other places ($44). It should be noted, however, that the methods available for the isolation of nucleoli do not permit obtaining preparations containing only nucleolar chromatin and therefore the results of the mentioned work cannot be interpreted unequivocally. To overcome this difficulty Ritossa and Spiegelman (246) studied the DNA complementary to ribosomal RNA in Drosophila larvae having four, three, two, or one nucleoli. The quantity of DNA complementary to rRNA was proportional to the number of nucleoli. An important observation proving that R-RNA synthesis is localized in the nucleolus itself is that RNA synthesized in isolated nucleoli is of the GC type in base composition (240). Thus it is established that a considerable part of ribosomal RNA synthesis takes place in nucIeoIar deoxyribonucleoprotein fibrils. However the possibilit-y that a fraction of this DNA is localized in chromosomes (perhaps in D N P fibrils bound with chromosomal nucleonemata) remains open. The data listed above permit one to conclude that a nucleolus is the cellular organelle where synthesis of a t least part of the R-RNA takes place. Let us discuss some aspects of this process in ,more detail, first of all, which fraction of R-RNA is definitely of nucleolar origin. Heavy R-RNA in most cases cannot be obtained from isolated nucleoli since it is degraded during the isolation procedure. Nevertheless, Steele et al.

+

+

325

NUCLEAR RIBONUCLEIC ACIDS

(158) succeeded in dctecting R-RNA in preparations containing nucleoneniata; the latter, however, are found both in the nucleolus and in chromosonies. I n order to identify R-RNA species localized in nucleolar nucleonemata, an autoradiographical study of nuclei during the hot phenol fractionation was undertaken (78). It has already been noted that hot phenol fractionation permits the separation of heavy R-RNA precursors. The nuclei obtained by cold phenol treatment contain labeled RNA both on chromosomes and in nucleoli, which are easily distinguishable under the microscope. After extraction a t 40°C the nucleoli are still preserved, while after extraction at 55°C they disappear. It was shown that after treatment a t 40°C the label is removed almost exclusively from nucleoli. It is inferred that the 40°C RNA fraction of phenol nuclei represents nucleolar RNA. Consequently, a considerable part of heavy 3 5 4 0 S R-RNA (R-RNA,) and newly formed 28 S rRNA (rRNAA) is localized in the nucleolus. Thus the nucleolus may be regarded as a site where the formation of the precursors for rRNAA takes place and where this precursor is transformed into true rRNAA chains. A similar conclusion has been made in two recent papers (86,246) where it was shown that isolated nucleoli contain newly formed 28 S but not 18 S rRNA. Data on the site of synthesis of R-RNA,, as well as information about its further interconversions, are meager. Phenol treatment a t 40°C leaves a part of the label in the nucleolus, and thus a nucleolar origin of 45 S RNA is possible. It is also not excluded, however, that it is synthesized not in nucleolar but in chromosomal nucleonemata. This suggestion is in good agreement with the fact of preparative separation of newly formed rRNAA and rRNAB by hot phenol fractionation (76) and with the absence of newly formed rRNAB in isolated nucleoli (86, 246). Such spatial separation of synthesis of two ribosomal RNA components is not impossible since, as is described in the next section, small and large ribosomal subunits may exist independently from each other both in nucleus and in cytoplasm.

2. THEASSEMBLYOF RIBOSOMES AND

THE

TRANSPORT OF R-RNA

The interaction of R-RNA with the protein probably takes place a t the first stage of R-RNA synthesis. Tamaoki and Mueller (160) have shown that, in nuclear lysates obtained by deoxycholate treatment, a fraction of labeled RNA sediments a t or faster than 70 S. Deproteinization decreases the sedimentation constant to 35-45 S (Fig. 9). Con-

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sequently, the formation of ribonucleoprotein strands, folding later into ribosomes, takes place a t the stage of polycistronic R-RNA, and this strand, not free R-RNA, is split into fragments. It is inferred that the assembly of ribosomes begins in the nucleolus. It should be noted that in vivo the nucleolus possesses an active protein synthesis. This is shown in experiments with isolated nucleonemata and nucleoli themselves as well (52, 11.6, 126, 163). The latter can incorporate amino acids in cellfree systems (162). It is thus not improbable that proteins interacting with heavy R-RNA are synthesized in nucleoli (or in nucleonemata in general). A number of authors have suggested (74, 164, 247) that R-RNA can be a template for ribosomal proteins. This hypothesis, while attractive, needs further experimental elaboration. The synthesis of ribosomal protein is most probably very important for the stabilization of R-RNA. I n experiments with actinomycin chase (actinomycin block after short pulse labeling) a considerable part of the label incorporated into heavy R-RNA is always degraded. Another part of this is recovered later in cytoplasmic ribosomes (74, 76, 229). It may well be that complex formation between R-RNA and protein is a limiting step in ribosome formation and the free R-RNA not included in a ribonucleoprotein structure is degraded. The splitting of heavy R-RNA gives precursors of ribosomal subunits different from completed ribosomes in some aspects (228a). Detailed studies of these particles are absent. Whether they are completed in the nucleus or in the cytoplasm is not known. On the other hand, some data indicate that joining of completed large and small ribosome subunits takes place outside the nucleus in the cytoplasm. Nucleoli, for example, contain a considerable excess of large free ribosomal subunits (161). A number of authors have shown that newly formed R-RNA in cytoplasm is found both in small and large ribosomal subunits but not in whole ribosomal particles (Fig. 12). I n contrast to free subunits, monosomes are labeled to a lesser extent than the ribosomes in polysomes (248, 160). So, small and large ribosomal subunits migrate to the cytoplasm separately. Thus the cell possesses two different channels for the formation of the two ribosomal RNA components and consequently of two ribosomal subunits. The detailed characterization of ribonucleoproteins contained in nucleolar and chromosomal nucleonemata, which are probably the site of ribosome formation, is of extreme interest nowadays, since these nuclear substructures may contain particles intermediate in the assembly of ribosomes.

NUCLEAR RIBONUCLEIC ACIDS

3. THEROLEOF

THE

327

NUCLEOLUS IN TRANSFER RNA SYNTHESIS

It has already been mentioned that nucleoli contain considerable amounts of tRNA. The existence of DNA sequences complementary to tRNA and the presence of newly formed tRNA in nuclei (13.3, 249) proves the nuclear origin of tRNA, although these facts do not give any indication of the exact localization of tRNA synthesis within the nucleus. To obtain this information Sirlin et al. (250) studied the incorporation of H3-pseudouridine, a specific tRNA precursor, in the intracellular structures of Hironomus. At short labeling times, the radioactivity is concentrated predominantly in nucleoli. A new approach to this problem has been exploited recently. It has been shown that 5,6-dichloro- and 4,5,6-trichloro-1- (b-D-ribofuranosyl) -benzimidazole inhibit chromosomal RNA synthesis while nucleolar RNA synthesis remains unaffected. Under these conditions the incorporation in nuclear sRNA remains unchanged (251). Thus the nucleolus is probably a site of tRNA synthesis as well. Unfortunately, there are no reports as yet on the precise localization of DNA complementary to tRNA. Another interesting aspect of these investigations is the study of tRNA methylation. Sirlin et al. have shown that the incorporation of methyl-C14 methionine is a measure of tRNA methylation if protein synthesis is blocked by actinomycin. It was shown by autoradiography that methylation occurs in nucleoli (652). Birnstiel et al. isolated nucleoli and chromatin and studied methylation in a cell-free system. Again it was shown that only the nucleoli are able to methylate RNA (253).Thus nucleoli seem to participate in two stages of tRNA formation: (1) its synthesis oh a DNA template resulting in appearance of unmethylated tRNA; (2) methylation of certain bases completing tRNA formation. The stages and mechanisms of tRNA transfer to the cytoplasm are unknown. Thus the nucleolus possibly together with the chromosomal nucleonemata represents an apparatus for the synthesis of nonspecific components of the protein-synthesizing machinery, the ribosomes and tRNA that later accomplish the synthesis of proteins under the direction of mRNA. We have discussed a number of questions concerning the localization and stages of RNA synthesis in nuclear structures, as well as some questions concerning the assembly of ribonucleoproteins within the nucleus. The last problem discussed in the present survey is the analysis of factors regulating these processes.

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E. Regulation of RNA Biosynthesis 1. SOMEGENERAL MECHANISMS OF

THE REGULATION AT THE GENIC LEVEL Regulation of RNA synthesis, or in other words regulation a t the level of transcription, is an important step in determining the protein and enzyme profiles of the cell (that is, its differentiation). There are of course other important regulatory mechanisms, in particular that a t the translation level in ribosomes, but they are probably less important in determining cellular differentiation. The regulation of RNA synthesis is studied in most detail in bacteria where Jacob and Monod discovered the repressor-operator mechanism of regulation (964). The essence of this well-known mechanism is the following. Within the genome the groups of cistrons for metabolically related enzymes are clustered in operons. Each operon has an operator gene and the transcription of the operon begins from the operator, If the operator is switched off, RNA synthesis in the whole operon is blocked. Furthermore, regulator genes exist in the genome and the products of these genes, repressors, may react with operators and therefore switch the operon off in RNA synthesis. The interaction between repressor and operator depends on the presence of effectors which either are necessary for the interaction or prevent it. I n bacteria, effectors may be either substrates or end products of metabolic pathways catalyzed by enzymes coded by the respective operon. Thus the appearance of the substrate in the medium leads to a chain process : interaction of the substrate with repressor, dissociation of the complex between repressor and operator, synthesis of mRNA on the operon, synthesis of enzymes, utilization of the substrate. When the substrate is removed, the repressor inhibits operon action again. In this way the bacterial cell easily adjusts itself to varying environmental conditions. Consequently, each gene in bacteria has the potential to synthesize mRNA. McCarthy and Bolton (96) have shown that the growing bacterial culture contains mRNA of all cistrons although in different amounts. The rate of mRNA synthesis on different cistrons is thus regulated. However, the nature of repressor substances remains unelucidated. There is experimental evidence that repressors may contain allosteric proteins (966-957).The involvement of polynucleotide chains responsible for the recognition of specific DNA sequences is also not excluded (257). A similar mechanism of regulation may well exist in higher organisms

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although there is no definite proof of its existence. Since cells of metazoa exist in media of relatively constant composition, such specific regulators as hormones or mediators and not the subdrates or products of reactions may well play the role of effectors. On the other hand, cells of higher organisms probably possess some other regulatory mechanism providing lasting switching-off of a considerable portion of the genome. This is suggested by the nonequivalence of cell nuclei: nuclei isolated a t early stages of embryogenesis as well as nuclei of germ cells may undergo normal development if transplanted to the embryo, while nuclei of many differentiated cells have lost this property (258, 2559). Although not conclusive, this fact is in a good agreement with the hypothesis of nuclear differentiation. On the other hand, it is known that the DNA complement, that is, the genetic set, is identical in all cells of each organism as shown most definitely by experiments on competitive hybridization (260). Thus higher organisms must possess a mechanism for the continuing exclusion of a number of genes from mRNA synthesis and this mechanism must be preserved in cell division. It should be noted in this connection that the structural organizations of genomes in bacteria and higher organisms show considerable differences. In bacteria, the genome is composed of free DNA or DNA with a small amount of proteins. The genome of higher organisms is an intricate nucleoprotein complex containing a special class of basic proteins, the histones (116). It might well be that a more complex organization of the genome reflects the aquiring of new regulatory mechanisms. However, it should be noted that the irreversibility of inactivation of a part of the genome is not absolute. It was shown recently that erythrocyte nuclei transplanted in tumor cell cytoplasm begin to synthesize both RNA and DNA, which are not synthesized under normal conditions (261). This shows again the complexity of the problem. Many facts suggest some relationship between the structural organization of certain regions of chromosomes and their participation in mRNA synthesis. The so-called condensed chromosomes (e.g., a metaphase chromosome with maximal folding of chromosomal material and aggregation of D N P threads) do not synthesize RNA. Metaphase chromosomes are absolutely inactive in RNA synthesis (262). In lampbrush chromosomes, one can differentiate chromocenters where D N P is aggregated and so-called loops-despiralieed regions of chromosomes (263).Autoradiographic experiments show that RNA synthesis proceeds exclusively in the loops (264). It is interesting in this connection that, at different stages of oogenesis, loops are formed by

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different regions of chromosomes, so that different DNA’s participate in RNA synthesis. Frenster et ul. (265) isolated condensed chromatin and extended despiralized chromatin from homogenates of sonicated lymphocyte nuclei (heterochroniatin and euchromatin, according to their terminology). The main part of the newly foriiied RNA is connected with euchromatin. However, these experiments do not prove that RNA is synthesized in the euchromatin regions since newly formed RNA may simply sediment together with it. Experiments on hybridization of nuclear mRNA with the DNA from hetero- or euchromatin are necessary in order to resolve this question unequivocally. Here it should be noted that the nature of differences between condensed and extended chromatin remains obscure. However, the existence of structural morphological differences between active and repressed chromatin suggests differences in their chemical organization. A number of approaches to the analysis of these differences constituting the basis of genic regulation of differentiation can be visualized at the present time. The main hypotheses explaining the biochemical mechanisms of the steady exclusion of certain genes from RNA synthesis are discussed below. 2. ROLEOF HISTOKES IN

THE

REGULATION OF MRNA SYNTHESIS

The hypothesis that histones are the regulators of gene activity and consequently of mRNA synthesis is one of the most intensively studied although not yet finally proved. This hypothesis was first formulated by Stedman and Stedman in 1947, who postulated that histones act as gene inhibitors (at that time the authors thought that not DNA but acid proteins are the genetic substance) (266) . Only in 1962 was it shown that histones may act as inhibitors of RNA synthesis on the DNA template (204, 267). The main supporting facts are the following. (,a) The removal of histones from isolated cell nuclei or from chromatin stimulates RNA synthesis in them to a large extent. Bonner and Huang isolated chromatin from pea seedling nuclei and showed that it has a limited ability to serve as a primer in RNA synthesis. After splitting of histones (using dissociation and subsequent ultracentrifugation in 4 M CsCI), the total RNA polymerase activity of deoxyribonucleoprotein increased some 25-fold. The addition of histones to the incubation medium again inhibited RNA synthesis, and, if histones were added in excess, the inhibition was essentially complete (204). Allfrey et $uZ. (267), who removed histones from calf thymus nuclei by trypsin treatment, also observed considerable activation of incorpo-

NUCLEAR RIBONUCLEIC ACIDS

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ration in RNA (two to threefold) although i t was less expressed than in the first case. It has also been shown that treatment of cell nuclei by concentrated salt solutions (0.4 M and higher) activates the RNA polymerase present in the nucleus (203, 268, 269). It is known that these conditions lead to a splitting of a portion of histones from DNA. ( b ) These observations were confirmed with a purified RNA polymerase system (270).Soluble nucleohistone prepared according to Zubay and Doty ( 2 7 l ) , containing about 50% of DNA and about 50% of histone, is inactive as a template in the RNA polymerase system of E. coli. If a t least a part of the histone is removed, the template activity of the complex increases sharply. The nuclear residue remaining after the extraction of nucleohistone by the above procedure has a lower histoneto-DNA ratio and is considerably more active as a template than the soluble nucleohistone. Artifically prepared nucleohistones, obtained by association of DNA and histones by dialysis from concentrated salt solutions, are inactive as primers in the RNA polymerase reaction (272). (c) Histones act not on the enzyme but on the template. If an excess of DNA is added to the incubation medium containing RNA polymerase, DNA, and histone, the synthesis of RNA is restored. This proves that not enzyme but template is inactivated by histone (270). (d) The inhibitory action of histones is not due to the insolubility of nucleohistone, originating from the interaction of histone with DNA (273). Soluble preparations of nucleohistone are still devoid of priming activity (272). On the other hand, if only a part of proteins is extracted from chromatin (for example, by treatment by 0.6 M NaCl) the resulting insoluble preparations are nevertheless active as templates in RNA synthesis (269). The observations outlined above suggest that histones may act as specific inhibitors of RNA synthesis in vivo. This suggestion is discussed below. The question as to which histones act as inhibitors of the RNA polymerase reaction raises many difficulties. It has already been noted that histones may be divided into several classes, and each class probably contains several molecular species of histones. These classes are : lysinerich histones (LH), moderately lysine-rich histones (MLH) , and arginine-rich histones (AH) [see (274)1. According to Bonner et al. (270, 272) only LH and to a lesser extent MLH may act as repressors while AH does not inhibit RNA synthesis. Moreover, nucleohistone preparations containing AH only but having a histone-to-DNA ratio similar to that of native nucleohistone have a template activity about the same as that of native DNA. Artificially

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prepared nucleohistone containing LH or MLH or polylysine is completely inactive (272). It is interesting in this connection that nucleohistone prepared by the method of Doty and Zubay has a melting point 15-20°C higher than native DNA. The results of Bonner e t al. ($70, 272) indicate that this effect is correlated with the inhibition of the priming activity of deoxynucleoprotein. Artificial nucleohistones containing LH, MLH, or polylysine have a higher melting temperature than DNA while nucleohistones containing AH do not. It should be noted that other investigators obtained different results. Allfrey and Mirsky have shown that RNA synthesis in nuclei is inhibited if AH is added, while the addition of LH was without effect. Later it was shown that the inhibitory action depends on the method of isolation of the histones. I n preparations obtained by acid-ethanol fractionation, AH is inhibitory, and if histones are fractionated on CMcellulose L H possesses inhibitory action (276). Only lysine-rich histone increased the D N P melting temperature. These data suggest that the inhibitory action may be due to a small fraction that, depending on the method of isolation, goes into one or another large fraction. I n this case, the inhibitory action may not necessarily be connected with the elevation of the DNA melting temperature. Hindley has studied the template activity of chromatin preparations after removal of histones by increasing concentrations of NaCl (268). Removal of L H has no effect on the priming activity of deoxyribonucleoprotein, while removal of AH stimulates it. Data conflicting with the results of Bonner were obtained also with the reconstituted histones (276). Finally, it has been shown in a number of papers that various histones may inhibit RNA synthesis, and that synthesis on DNA's containing more AT pairs is inhibited to a greater extent (277, 278). Probably one may consider that, in particular conditions, histones may act as inhibitors of RNA synthesis. In order to know which histone is a true repressor acting in viva one should demonstrate the specificity of the in vitro inactivation, in other words, demonstrate that, in in vitro conditions, the particular histone inhibits the same parts in the genome that are repressed in vivo. There is considerable indirect evidence as to the possible repressor functions of histones. It is known that a phenotypical expression of the activity of genes is conserved in cellular divisions. Therefore, doubling of the DNA during the cellular cycle must be connected with a simultaneous doubling of repressor proteins, and it appears that this is really the case (279). Later investigations have shown that histones are syn-

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thesized not only in those cells that synthesize DNA but also in differentiated, nonproliferating cells, although in the latter the intensity of histone synthesis is lower (274, 280). However, i t has been shown recently that this is not true for all histones. I n the absence of DNA synthesis, only AH turns over while synthesis of LH and MLH occurs only in those cells that actively synthesize DNA (281). These observations are in accord with the suggestion that some histones may provide long and lasting inactivation of certain groups of DNA molecules. Another fact is the ability of histones to provoke condensation of chromosomes. Activation of DNA in RNA synthesis is connected with the transition of D N P from the condensed to the extended state. Addition of histones to lampbrush chromosomes results in the disappearance of loops, that is, in condensation of the extended D N P contained in loops (282).Removal of a part of the histones (LH) from nuclei results in some disaggregation of condensed chromatin (283). If addition or loss of histone leads to a transition of D N P from an extended to a condensed state and vice versa, and if extended chromatin is active, one should suggest the existence of two kinds of DNP: one containing histone and the other either not containing histone a t all or containing only a fraction of it. This speculation is indirectly suggested by the data of Bonner (27’0) who showed that the D N P remaining after extraction of nucleohistone by water contains considerably less histone. The melting curve of this D N P is close to that of free DNA and it may serve as a template in RNA synthesis. These data should be confirmed. The relative content of histone in heterochromatin and euchromatin isolated from thymus nuclei was quite similar (284). In histochemical experiments, the ratio of DNA to histone appears to be approximately the same both in the active regions of chromosome “puffs” and in the repressed regions (286). Thus there is no direct evidence correlating the content of histones in chromosomes and their functional state, although it seems probable that such correlations do exist. It may be concluded from the above that histones may act as inhibitors of certain loci of DNA in RNA synthesis in vim. But i t should be taken into consideration that since histones easily react with acid polymers, in particular with nucleic acids, they may inhibit a number of metabolic processes in vitro. For example, they inhibit the DNA polymerase reaction (686, 287), glycolysis and synthesis of ATP in nuclei (288), and a number of other reactions. Therefore, it is not clear which of these effects corresponds to real processes in vivo. This is why this problem needs further experimental elaboration. One of the approaches has been exploited by Bonner et al. (289) who

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studied protein synthesis in a cell-free system containing ribosomes and RNA polymerase from E . cooli; D N P from various parts of pea seedlings served as a template. I n this system, mRNA synthesized on a D N P template was used as a template in protein synthesis. It was shown that if D N P from the cotyledon is used as a primer about 1.5% of the protein synthesized in the system shows the immunochemical characteristics of globulin. This globulin is actively synthesized in cotyledon cells in viva On the other hand, if D N P from the apical bud is used as a primer, the incorporation of label in globulin in vitro is negligibly small. Normally, apical buds do not synthesize globulins. DNA’s isolated from both organs behave in a different way: they stimulate the incorporation into globulins equally but to a small extent (Table VIII). TABLE VIII THE NATURE OF THE PROTEIN FORMED UNDER THE INFLUENCE OF RNA ON DNA OR DNP TEMPLATES FROM DIFFERENT ORGANS SYNTHESIZED OF PEA SEEDLINGS (289)

Template for RNA synthesis Apical bud chromatin Cotyledon chromatin Apical bud DNA Cotyledon DNA

Total incorporation of amino acid into protein (cpm)

Incorporation into globulin (CPd

Incorporation into globulin (% of total incorporation into protein)

15,650 41 ,200 23,600 14 ,000 15,200 14,200 5 ,600 50,100

16 54 341 226 60 72 22 157

0.10 0.13 1.45 1.61 0.40 0.51 0.39 0.31

These experiments suggest that, in vitro, intact D N P contains the factors permitting RNA synthesis on a limited number of cistrons and that these factors are absent in purified DNA. These data also demonstrate that these factors block DNA binding sites for the polymerase, since in these experiments an excess of RNA polymerase was supplied exogeneously. Unfortunately, this result has not been confirmed on other material, although it has been shown that chromatin from tissues with active RNA synthesis is a better primer than chromatin from inactive tissues (290,291) . The experiments discussed above permit a commonplace conclusion to be drawn, namely, that agents blocking binding sites for the polymerase are proteins. There is no information about the nature of these

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335

proteins; that is, it is not known whether they are histones, histone fractions, or nonhistone proteins. Another approach to the problem is through DNA-RNA hybridizat.ion and competitive hybridization, which permits the analysis of the nature of the RNA synthesized on a given deoxynucleoprotein template (269). In this approach, RNA is synthesized in a cell-free system using DNA or DNA protein from some organ as a template. The reaction is catalyzed either by endogenous polymerase or by exogenous enzyme. After the reaction, hybridization of the isolated synthesized RNA with template DNA is studied. mRNA isolated from the cells chosen as a source of DNA or D N P is used in competition experiments. The competition in such experiments may indicate the similarity of RNA molecules synthesized in vivo and in vitro; if there is no competition, one can conclude that factors restricting RNA synthesis only a t a part of the cistrons are lost. Such experiments were conducted with D N P preparations from Ehrlich ascites tumor cells (269) (Fig. 13). If RNA is synthesized on a DNA template by bacterial polymerase, the newly formed RNA is effectively hybridized with DNA. If chromatin is used as a template, the extent of RNA synthesis is considerably lower, and the hybridizability of RNA with DNA is less pronounced, but the competition between the RNA synthesized in vitro and mRNA is more clearly expressed. Such experiments suggest that fewer cistrons are active when D N P is used as a primer, compared to DNA. Other experiments have employed a stepwise dissociation of chromosomal D N P by neutral salt solutions of increasing ionic strength. After the removal of the dissociated protein, the D N P complex was tested in an RNA polymerase system. Upon increasing the ionic strength of the extraction medium to 0.4, the template activity of the chromatin remained low. However, in the interval 0.4-0.6, an increase of the template activity of the chromatin was observed, although it remained insoluble in the incubation medium of the RNA polymerase reaction. I n this interval of ionic strengths, about 1520% of the protein was split from the chromatin complex. A further increase of ionic strength to 1.5 leads to a gradual dissociation of other proteins from DNA and is accompanied by a further increase of template activity that, however, correlates with the rise in D N P solubility. The increase of chromatin template activity in the interval of ionic strengths 0.4-0.6 is accompanied by an increase of hybridization of the RNA synthesized. On the other hand, a certain fall in competition with mRNA is observed, and the value shifts to that of RNA synthesized on purified DNA. Consequently, the dissociation of a relatively small portion of protein upon increase of ionic strength from

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G. P. GEORGIEV

0.4 to 0.6 leads to a randomization of RNA synthesis: all DNA cistrons become equally accessible for exogenous RNA polymerase. These results suggest that certain regulatory mechanisms functioning in viv'o are preserved in the cell-free system used and that proteins removed in the interval of ionic strength 0.4-0.6 are responsible for the repression of most of the DNA cistrons. This protein fraction consists of a part of the LH (fl) histone and nonhistone protein (1:l). The nature of the active material is now under investigation. It deserves mention that, if RNA synthesis on a chromatin template is catalyzed by its own enzyme, the removal of protein by increase in ionic strength

20

Hybridizability

10

* 0.15

0.4 0.6 0.8 r12

10

1.5

2.0

FIG.13. Alteration of priming activity of D N P and ability of RNA synthesized on D N P template to hybridize during stepwise dissociation of D N P from Ehrlich ascites cells by KC1 or NaCl solutions with increasing strength (269).

to 0.6 does not affect the character of the RNA synthesized. Although the synthesis is activated, the degree of hybridization and of competition with mRNA does not change. It is inferred that endogenous polymerase is localized only in connection with derepressed cistrons and is absent in repressed cistrons. Thus it is probable that in vivo the complex formation between the enzyme and the template, that is, the first stage of RNA biosynthesis, may be a limiting reaction. The above experiments outline some approaches to ascertaining the nature of the repressors of mRNA synthesis in the chromosomes of higher animals. These experiments also permit an attack on the problem of the structural organization of DNP complexes possessing priming activity. For this purpose, DNP was isolated from the solution a t ionic strength 0.6, that is, under conditions when it is derepressed and possesses template activity. This D N P can be separated from dissociated proteins by gel filtration on a Sephadex G-200 column. This DNP contains considerable amounts of histone (whole f3 and f2 fractions), that is, AH (srginine-

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rich) and MLH (moderately lysine-rich). The ratio of protein to DNA in this preparation is 1.25. About three quarters of this protein is histone. Such D N P does not precipitate in solutions of low ionic strength. The electron microscope permits one to visualize threads with a diameter of 30 A in such preparations. The melting temperature of such preparations is considerably higher than that of free DNA. Consequently, the properties of D N P isolated in 0.6 M NaCl are simiIar to those of D N P obtained by the technique of Zubay and Doty (besides a lack of lysine-rich histone) (271). However, D N P isolated in 0.6 M NaCl, in contrast to D N P isolated in low ionic strength, reveals considerable template activity in the RNA polymerase system (269). One may suggest that the genes derepressed in vivo are represented not by free DNA but by a nucleoprotein or nucleohistone. Addition of a small protein fraction, perhaps of a histone nature, transforms these genes into an inactive state, so that they cannot support the RNA polymerase reaction. One may also suggest that this “repressor” histone is responsible for the transition of D N P into a condensed state, probably because of cross linking between individual D N P threads. I n a recent publication, Littau et al. presented experimental data in favor of the hypothesis that these cross links between D N P threads consist of lysine-rich histone molecules (283). However, one should not think that histone components of active D N P play only a structural role and do not participate in regulation of gene activity. It may well be that they regulate the rate of RNA synthesis on derepressed cistrons; that is, they may be involved in mechanisms of rapid and transient regulation of gene activity. It is interesting in this connection that Huang and Bonner have shown that, in chromosomes, histones are complexed with a special class of low-polymer RNA (90). According to their hypothesis, the repressor represents a complex between several histone molecules and this RNA. This RNA participates in the recognition of the DNA that should be repressed. It may be suggested from the above that proteins of D N P , mainly histones, provide lasting inactivation of genes responsible for differentiation and accompanied by condensation of chromatin, as well as the regulation of the rate of RNA synthesis in working cistrons. However, a t the present time we have only reached that stage of experimentation where we can suggest hypotheses but not prove them. Elaboration of new experimental approaches will help to resolve these questions in the near future. 3. OTHERPOSSIBLE REGULATORY MECHANISMS

a. Acid Proteins and Other Polyanions. Another possibility that should be taken into consideration is the participation of acid proteins of

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DNP in the regulation of gene activity. Nonhistone prot,eins of DNP form a group of insufficiently characterized proteins insoluble in acids and showing a higher content of dicarboxylic amino acids. I n contrast to a histone: DNA ratio that is rather constant in different cells, the ratio of nonhistone proteins to DNA varies widely and, as a rule, is higher in tissues with intensive RNA synthesis (292, 131). This ratio is very low in condensed chromatin regions and is very high in extended chromatin., although these values may be influenced by artifacts. The addition of acid proteins or other polyanions to DNA templates in vitro stimulates RNA synthesis but the stimulation is small and probably nonspecific (284, 293). These data are the basis for a suggestion of Frenster that nonhistone proteins of DNP may act as derepressors. This hypothesis postulates that all DNA molecules are bound to identical sets of histones and in such a state all of them are blocked, However, several nonhistone proteins interact with nucleohistone to force histones out of the complex with DNA and, as a result, the DNA may participate in the RNA polymerase reaction. This hypothesis is quite interesting but it lacks clear experimental confirmation. The possibility of complex formation between nuclear phosphoproteins and histones was demonstrated recently by Langen and Smith (294). Complex formation lowered the inhibitory effect of histones, but phosphoproteins were unable to force the histone out of its complex with

DNA. b. Role of RNA in R,epression and Derepession of the Genome. According to another suggestion, a special class of informational RNA participates in the regulation of gene activity. It is known that chromosomes of metazoa contain large amounts of D-RNA (48). This D-RNA is metabolically and functionally heterogeneous. A part of i t functions a t the site of its synthesis and is degraded there (68,84). Hadjivassiliou and Brawerman (5’81) have shown that a part of this

D-RNA does not possess template activity. It has been suggested that this D-RNA ,may interact specifically with DNA, leading to its repression or derepression (293). However, we incline to the idea that this RNA participates in regulation of gene activity in an indirect way, serving as a template for the synthesis of nuclear regulatory proteins (histones). It is interesting in this connection that RNA functioning a t the site of its synthesis in the nuclei breaks down rapidly (104). On the other hand, histones and nonhistone proteins of the DNA complex are the only known group of proteins in animal cells the synthesis of which is inhibited rapidly upon inhibition of RNA synthesis (23&233). Therefore, it seems probable that

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rapidly turning-over, unstable D-RNA participates in the synthesis of proteins of the D N P complex. We have already noted that a special class of low-polymer RNA complexed with histones may participate in the regulation of mRNA biosynthesis. However, new experiments are necessary in this direction. c. Polyamines and O t k r Low Molecular Compounds. It has been shown in a number of papers that certain less complex compounds may affect the RNA polymerase reaction. For example, polyamines such as spermidine or putrescine activate the RNA polymerase reaction and increase the specificity of interaction of the enzyme with the template, preventing the complex formation between the polymerase and RNA (296, 296, 211). Since these compounds are found in vivo they might regulate RNA synthesis on some particular genes. d. Structure of DNA. Finally, one cannot exclude the possibility that repression or derepression of genes is due to alterations in DNA structure. These alterations may result in different activities of DNA regions in RNA polymerase reaction. For example, purified DNA from phage T2 catalyzes the synthesis of predominantly “early” mRNA in an RNA polymerase system (297). Although not very probable in higher organisms, this idea must be considered in the discussion of regulation of mRNA biosynthesis. Thus, in addition to a histone mechanism of regulation of genic activity, other possibilities are suggested. It may well be that several different mechanisms of regulation of RNA synthesis are operating in vivo.

4. SOMEASPECTSOF REPRESSOR ACTION It has been shown in preceding sections that proteins are the most probable candidates for gene inhibitors. This is valid both for the steady and lasting repression responsible for differentiation and for a transient control of the rate of RNA synthesis. Repressors of both kinds contain proteins. The question then arises as to which mechanisms are responsible for the recognition of DNA regions by a repressor. There are several possibilities: it may be a special low-polymer RNA (90). This RNA may recognize a corresponding DNA according to the complementarity principle. ( b ) Recognition of certain nucleotide sequences by a repressor itself. An example of such recognition is given by aminoacyl-tRNA synthetases or methylases of nucleic acids.

( a ) Presence of RNA in repressors. For histones

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(c) Recognition of sites altered by chemical modification of bases. This

suggestion has no experimental support so far (297). ( d ) A special localization of repressor synthesis (in the case of histones). One may suggest that each operon contains regions coding for histones. Synthesis of histones takes place in ribosomes on mRNA still attached to the template. In this case, a histone repressor is situated near its operon and will interact with it and not with a neighboring one. Only further experimentation may help to make choices among all these hypotheses. Another very fundamental problem is the problem of the nature of factors responsible for switching genes on and off. Works published in the past 2 years have demonstrated the important role of cytoplasmic factors in the activation and repression of genes. For example, Harris has found (261) that fusion of nuclear erythrocytes with tumor cells activates RNA and also DNA synthesis in erythrocyte nuclei. On the other hand, if nuclei of somatic cells are transplated to embryonic cells, these nuclei lose the ability to synthesize R-RNA and synthesize only D-RNA, typical of nuclei of early embryos. Transplanted nuclei begin to synthesize R-RNA only a t that stage of development a t which it begins in normal embryos (298). I n both cases, cytoplasm dictates the pattern of nuclear activity. It is interesting in this connection that nuclear differentiation, for example in the case of nuclear erythrocytes, is a t least partially reversible. The nature of the cytoplasmic factors remains unknown but the experiments described mark ways for further progress in this field. Allfrey et al. (299) have shown that acetylation of histones decreases their inhibitory action on the template. Acetylation of amino terminal groups in histones may be observed both in vivo and in vitro. If RNA synthesis is activated (e.g., by partial hepatectomy) , the increase of acetylation of histones is one of the earliest metabolic events (300). The authors have suggested that acetylation of end groups in histones may be a mechanism for gene activation. However this hypothesis demands further experimental support. These are of course only first approaches to the study of the mechanisms of development and cellular differentiation but they imply that essential progress will be made in this direction in the next few years.

V. Conclusion The data discussed in the second part of the present survey may be briefly summarized as follows.

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Three types of cellular RNA are synthesized in cell nuclei: mRNA, rRNA, and tRNA. All are synthesized by RNA polymerase. Synthesis takes place with double-stranded DNA as a template but only one of its chains is copied; the synthesis is asymmetrical. The mechanism of this phenomenon remains unknown. A complex between template enzyme and product is formed during RNA synthesis. The requirements of complex formation between the template and enzyme and of RNA synthesis itself are different. mRNA is synthesized in chromosomes. The liberation of mRNA from the complex with enzyme and template and its transport to the cytoplasm are probably accomplished by special ribonucleoprotein particles with a sedimentation constant of 30 S. A part of the nuclear RNA remains and probably functions in chromosomes a t the site of its synthesis where it is rapidly degraded. It is suggested that it may participate in the synthesis of proteins of the D N P complex. rRNA synthesis takes place in nucleolar nucleonemata and possibly also in nucleonemata of chromosomes. Synthesis of two types of ribosomal RNA proceeds via separate channels from the stage of polycistronic precursors. The assembly of ribosomes takes place also in nucleonemata and probably separately since small and large subunits of ribosomes are transported to the cytoplasm and only then are joined to ribosomes. Synthesis of tRNA as well as its methylation also take place in the nucleolus. Higher organisms probably possess two mechanisms for the regulation of RNA synthesis: a mechanism of steady repression of genes and a rapid mechanism for the regulation of the rate of mRNA synthesis. Histones play an essential role a t least in the first mechanism. Nonhistone proteins and RNA of D N P complex may also participate in the regulation of mRNA synthesis. The identification of true repressors, the clarification of mechanisms of interaction of repressor and template, and the factors switching genes off and on are not yet concluded. Studies of these problems are of utmost importance for elucidating the secrets of cellular differentiation.

NOTEADDEDIN PROOF Many papers submitted for publication in 1966 are devoted to further studies of nuclear D-RNA (301-303).It wash shown that sedimentation constants of most rapidly labeled D-RNA’s are very large and reach values of 60-70 S and more. These giant D-RNA molecules are metabolically labile and degrade soon after synthesis without leaving the nucleus. Thus the main part of this RNA corresponds to the D-RNA, described in Section II,C but has even larger sedimentation constants. On the other hand, further studies of the hot phenol fractionation

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procedure show that after short labeling with radioactive RNA precursors, a significant part of the labeled D-RNA is not liberated a t 63". This D-RNA may be extracted by phenol-SDS treatment a t 85". In the actinomycin chase, this fraction loses 7 5 4 0 % of its radioactivity in contrast to the 63O-fraction of D-RNA, which is stable in the conditions of the actinomycin chase. Thus the separation of metabolically labile and stable D-RNA's has been achieved (304). A new method for the purification of nuclear D-RNA is proposed by Penman and Smith (305) who observed the solubilization of D-RNA after treatment of isolated nuclei by DNase in 0.5 M KCl. The R-RNA of nucleoli remained in the residue after such extraction. The D-RNA obtained is of relatively high molecular weight, and thus the described procedure may be useful for further investigation of nuclear ribonucleoproteins containing mRNA. Some additional properties of particles containing nuclear mRNA have been described (306). It was shown that these particles may be reversibly dissociated into RNA and protein by increasing the ionic strength of the solution. The particles obtained during the reassociation procedure do not differ from the original with respect to sedimentation properties or electron microscopic appearance. The dissociation-reassociation reaction was also visualized by CsCl density gradient ultracentrifugation. Interesting results correlating with those presented in this paper were obtained by Paul and Gilmour (307), who studied the properties of RNA's synthesized by RNA-polymerase on DNA or D N P templates. On the basis of hybridization experiments, i t was calculated that only about 10% of the DNA in D N P preparations is active as a template. Some indications were also obtained that different parts of genomes are repressed in D N P preparations from different tissues in such experiments (308).

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Replication of Phage RNA CHARLES WEISSMANN AND SEVEROOCHOA Department of Biochemistry, New York University School of Medicine, New York, New York

I. Introduction . . . . . . . . . . . . . . . . 11. General Properties and Biology of RNA Phages . . . . . . 111. Mutants of RNA Phages. . . . . . . . . . . . . IV. Messenger Function of Phage RNA . . . . . . . . . V. Replication of Phage RNA . . . . . . . . . . . A. In Vivo Studies . . . . . . . . . . . . . . B. Enzyme Studies . . . . . . . . . . . . . . VI. Conclusions and Summary . . . . . . . . . . . . VII. Appendix: Identification and Analysis of Viral RNA. . . . . A. Identification of Virus-Specific Double-Stranded RNA . . . B. Analysis of Double-Stranded RNA (Specific Dilution Assay) . C. Determination of “Minus” Strands. . . . . . . . . D. Determination of ‘‘Plus” Strands . . . . . . . . . E. Determination of Base Sequence Homology between Viral RNA’s . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . .

353 354 355 357 358 359 377 386 387 387 388 392 392 395 395

1. Introduction Viruses containing RNA as genetic material are widespread in nature, afflicting animals, plants, and bacteria alike. They have, with few known exceptions, one feature in common that sets them aside from most other viruses and living systems, namely, the single-strandedness of their genome. This property has an important consequence for the reproduction of RNA viruses; the genome itself can serve as a messenger for the synthesis of virus-specific proteins, so that the process of transcription, which plays a central role in genetic expression in general and in the replication of DNA viruses in particular, is bypassed altogether. The replication of the single-stranded RNA genome has long presented an intriguing problem. For one thing, viral RNA synthesis could not be attributed to enzyme systems responsible for RNA synthesis in the normal host. For another, it was not clear by what mechanism a 353

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replica of a single-stranded RNA could be produced. I n fact, early failures to detect the presence of viral “minus” strands or virus-specific double-stranded RNA had led to the suggestion that replication might proceed by a direct copying mechanism, rather than by a process based on the principle of base complementarity underlying DNA replication and transcription. Reports from several laboratories have cast some light on these problems. It has been shown that viral RNA replication is carried out by a novel enzyme system arising in host cells after infection and that complementary viral RNA (“minus strands”) accumulates in almost all hosts infected with RNA viruses studied so far, be they animal, bacterial, or plant cells. This essay is concerned with the replication of the RNA of RNA phages. However, reference is made to the replication of other RNA viruses, either to point out similarities or to stress differences. Zinder (%a) has reviewed the discovery, general biology, and the genetics of RNA phages. Earlier work on the replication of phage RNA has also been reviewed (1, l a , 2, 2 u ) .

II. Geneml Properties and Biology of RNA Phages RNA phages are small polyhedral viruses, about 200 A in diameter, having a particle weight of about 4 X lo6. They consist of 180 capsomeres, each of a molecular weight of 17,000 ( S ) , and an RNA molecule of about 106 molecular weight (4-7). A number of RNA coliphages have been isolated and described in some detail, such as f2 ( 4 ) , MS2 ( 8 ) , R17 ( 9 ) , M12 ( l o ) ,fr (11, l a ) , f,,, ( I J ) , &a ( 1 4 ) , and several others (15).The first six phages named are closely related serologically (16). Analysis of the coat protein of R17 and M12 reveals that these may differ by as little as one amino acid (3) ; however, the analyses of pancreatic ribonuclease digests of R17 and M12 RNA suggest more extensive differences between the two phages (17). The RNA’s of MS2 and f2 show about 7576 homology in their base sequences as determined by annealing techniques (18, 19). MS2 and Qp show no sequence homology (18, 19) and their coat proteins appear to be quite different ( 2 0 ) . The coat proteins of f2 and fr differ a t several sites ( 2 1 ) . RNA phages specifically infect male E . wZi ( 4 ) or bacteria containing the E . coZi F agent ( 2 2 ) . The phage attaches to the male-specific F-pili (i?2-25) and enters the eclipse phase. The phage RNA may be introduced into the host via these pili, the coat protein remaining outside (26).The actual penetration of the RNA into the host cell, but not the attachment of the phage particle, is dependent on divalent cations such as Mg++ (27); between the attachment and penetration phase the phage

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RNA becomes susceptible to RNase (28). The specificity of RNA phages for male bacteria arises from the attachment step, since spheroplasts of the female strains as well as those of Salmonella, Shigella, and Proteus may be infected by the purified viral RNA (69,30). After infection in broth a t 37OC, an eclipse period of 15 minutes is observed, following which infectious particles accumulate within the cell. Lysis of infected bacteria occurs between 22 and 60 minutes, and each cell may yield up to 40,000 virus particles, of which, however, only a fraction (5-50oJo) is viable, depending on the phage and the growth conditions (4, 9, 31). It is of interest that in the case of phage fr, under certain conditions (low temperature), synthesis and release of phage may occur without lysis of the host cell ( 3 2 ) ,which continues to reproduce while synthesizing virus. Infected bacteria continue to synthesize host-specific DNA, RNA, and proteins, albeit a t a diminished rate (4, 28). The formation of RNA phages is independent of host DNA synthesis (33, 34) and no base sequence homology between the phage RNA and the DNA of either infected or noninfected host cells has been detected (36). Exposure of infected E . coli spheroplasts to actinomycin D, a t concentrations that inhibit host cell RNA synthesis completely, does not abolish RNA phage synthesis (36). These findings speak against the participation of host DNA in the synthesis of RNA phages. At high concentrations of actinomycin, RNA phage synthesis is also greatly inhibited (37) ; the reasons for this inhibition are not understood. However, similar observations have been made in the case of animal RNA viruses (38, 39). Protein synthesis is required not only for the formation of the coat protein but also, during the early infective period, for synthesis of viral RNA. This is shown by the fact that chloramphenicol added within 2 4 minutes of infection completely prevents formation of viral RNA and decreases its synthesis when added within 10 to 15 minutes after infection (31, 40). These findings are in accordance with the subsequent discovery that one (or more) virus-specific RNA-synthesizing enzymes are produced in the host within minutes after infection (41-43).

111. Mutants of RNA Phages Conditional lethal mutants of RNA phages obtained by treatment with nitrous acid or fluorouracil have been important tools for the study of viral RNA replication. Host-dependent mutants of f2, sus (sus = suppressible) have been selected because of their capacity to grow on E . mli S26RlE, a strain containing the suppressor gene Su-1 ( 4 4 ) , while failing to multiply on the isogenic suppressorless (Su-) strain 526 (45). These mutants are therefore defined as N1 or amber mutants, a class

356

CHARLES WEISSMANN AND SEVER0 OCHOA

of mutants in which the lesion arises from the conversion of a “sense” codon (i.e., a codon specifying an amino acid) to the “nonsense” codon UAG (46, 47). The presence of this untranslatable codon results in an interruption of the polypeptide chain a t the corresponding site (48). In organisms containing an amber or N1-type suppressor gene (such as Su-1, Su-2, or Su-3) the nonsense codon is translated as serine, glutamine, or tyrosine, respectively (44). The insertion of one of these amino acids permits the synthesis of an intact polypeptide chain, which, however, is not necessarily functional. Temperature-sensitive mutants (designated ts) of f2 have been obtained either by mutagenesis with fluorouracil (49)or nitrous acid or as spontaneous revertants of host-dependent mutants (60, 61) and have been selected because of their capacity to grow a t 34°C but not a t 42°C. Host-dependent and temperature-sensitive mutants fall into three classes, which define three cistrons of the f2 genome (61). Class 1. The prototype of this group is ts-6. Temperature-shift experiments show that the temperature-sensitive function is required throughout the period of viral replication. Under nonpermissive conditions, no coat protein antigens and no viral RNA are formed; for reasons discussed on p. 362 it was concluded that class 1 mutants lack the function required for the synthesis of “minus” strands (62). Possibly the host-dependent mutant sus-10 also belongs to this group (61). Class 2. Under nonpermissive conditions, mutants such as ts-16 or sus-1 are deficient in a function required late in viral replication. Although an almost normal burst size is observed, the virus particles produced are nonviable and defective. They are not capable of adsorbing to host cells, they contain a smaller amount of RNA per particle than normal f2, and the RNA itself is partly degraded and noninfective for spheroplasts (50,51). Since the RNA synthesized under nonpermissive conditions was normal and infective, and since mutants of this class coinplement with known coat protein mutants (class 3), Lodish et al. (60) concluded that the coat protein itself is normal and that the maturation step required for the assembly of progeny particles is either altered or lacking in these mutants. This might result in the formation of defective particles, the RNA of which becomes damaged either during assembly of the virion or subsequently. Class 3. A typical mutant of this class is sus-3. A nonpermissive host (S26) infected with a class 3 mutant produces no phage antigens (45),although penetration of the viral RNA into the’ host cell occurs normally. When a mutant containing the sus-3 lesion is grown on the permissive host S26RlE, which contains the Su-1 suppressor, viable phage particles are formed. However, the coat protein differs from its wild-type counterpart in that the sixth amino acid from the amino-

REPLICATION O F PHAGE RNA

357

terminal end is serine rather than glutaiiiine (56a-54). In vitro experiments show that, in a nonsuppressed system, only a polypeptide fragment consisting of N-formylmethionine followed by the first five amino acids of the coat is formed, in accordance with the hypothesis that the codon specifying the sixth amino acid (or the seventh, if N-formylmethionine is considered) is a nontranslatable nonsense codon. In view of the fact that the lesion of the class 3 mutants was localized unambiguously in the coat proetin cistron, and under the assumption that no further mutation was present, it was surprising to find that the RNA replication of these mutants is also profoundly affected in nonpermissive hosts. When class 3 mutants are grown on nonpermissive hosts, viral RNA polymerase activity and the amount of virus-specific doublestranded RNA formed are abnormally high (5 to 15 times the wild-type level). Moreover, the double-stranded RNA is fragmented, for reasons that a t present remain unexplained. It may be noted, however, that an enzyme capable of cleaving double-stranded RNA is present both in normal and in infected E. coli ( 5 5 ) .When grown on the permissive host (containing the Su-1 suppressor), viable phage particles are produced and viral RNA polymerase and double-stranded RNA are only moderately increased (56, 57). From these and other findings, it was concluded that formation of coat protein is required to regulate the synthesis of the viral RNA polymerase responsible for the formation of doublestranded RNA (67). Mutants of different classes complement each other, in the case both of host-dependent (58) and of temperature-sensitive mutants (59). Streptomycin suppression of the lesion in the host-dependent mutant sus-3 has also been observed (60), confirming the notion that this phenotypic repair, first observed with certain E. coli mutants, is operative a t the translational level (61).

IV. Messenger Function of Phage RNA After penetration into the cell, parental phage RNA is found associated with the 70 S ribosome fraction. The f2 RNA functions as a messenger in the cell-free system of protein synthesis derived from E. coli (66).The protein synthesized under the direction of f2 RNA consists largely of f2 coat protein, as judged by analysis of tryptic digests, as well as of histidine-containing polypeptides. Later work indicates that this system forms intact coat protein subunits (6s). Since the coat protein of f2 phage does not contain histidine, viral proteins other than coat protein appear to be made in vitro in the presence of f2 RNA as messenger. Whereas the NH,-terminal amino acid of R17 coat protein is alanine, the product synthesized in vitro by the E. coli system, in the presence of R17 RNA, contains N-formylmethionine as the NH,-termi-

358

CHARLES WEISSMANN AND SEVER0 OCHOA

nal amino acid, followed by alanine ( 6 4 ) . This result is analogous to that obtained in similar experiments with RNA from a class 3 mutant of f2 mentioned in the previous section ( 5 3 ) . These findings suggest the presence of an initiation signal in the coat protein cistron, apparently a codon that directs the incorporation of N-formylmethionine as the first amino acid of the polypeptide chain. This is mediated by a special methionine tRNA (met tRNA 11) whereas the incorporation of methionine a t internal positions is mediated by another methionine tRNA (met tRNA I) (66).Under the in vitro conditions, formylmethionine remains attached to the polypeptide but in vivo this residue is removed, leaving alanine as the NHAerminal amino acid (63,6 4 ). f2 RNA also directs the formation of f2 coat protein in a cell-free system from Euglena gracilis, thereby providing an example for the postulated universality of the genetic code ( 6 6 ) . I n the presence of MS2 RNA, the cell-free E. coli system forms three proteins (67). One of them, lacking histidine, has been identified as coat protein. Coat protein synthesis starts immediately upon addition of MS2 messenger whereas histidine-containing proteins appear after a lag of about 5 minutes. Moreover, coat protein is formed in much larger amounts than the other proteins. These findings were thought to indicate that control mechanisms regulating the order and the frequency of translation of individual cistrons in a polycistronic messenger are operative also in vitro. Some aspects of the mechanism of suppression have been clarified by using RNA’s from amber mutants of RNA phages as messengers. As previously mentioned, when the RNA contained a nonsense codon (UAG) in the coat protein cistron, only an amino-terminal fragment of that protein was formed (64, 63). Suppression was obtained under in vitro conditions when tRNA, prepared from E. coli containing the Su-1 suppressor gene, was added to the system (68, 64). Serine was inserted at the site corresponding to the nonsense codon and serine tRNA was identified as the factor responsible for suppression.

V. Replication of Phage RNA The earliest event that can be related to the synthesis of phage RNA is the conversion of parental RNA strands to a double-stranded form1 by ‘As pointed out elsewhere in the text, it is conceivable that the “plus” and “minus” strands within the replicating complex, as it occurs in wiwo, may not have the tightly hydrogen-bonded, RNase-resistant structure that is observed after its isolation by standard methods involving phenol extraction (79, 118). This possibility must be borne in mind whenever mention is made of the involvement of doublestranded RNA in replication (cf. page 379).

REPLICATION OF PHAGE RNA

359

synthesis of a complementary (“minus”) strand (69, 7 0 ) . Subsequently, several hundred additional “minus” strands, most of which can be isolated as double strands, are formed in each infected cell. At least a part of the double-stranded RNA, which, after an isolation procedure involving RNase, was characterized as a double helix consisting of equivalent amounts of “plus” and “minus” strands, appears to be derived from a more complex structure, the replicative intermediate (71). This intermediate has been shown to turn over during RNA synthesis in a fashion compatible with the concept that the “minus” strands serve as templates for the synthesis of “plus” strands, whereby the newly formed “plus” strands displace their pre-existing counterparts from the replicating complex (72, 7 3 ) . Two enzyme preparations have been purified from phage-infected E. coli; their action in vitro gives rise to viral RNA, which was characterized either by annealing techniques (74, 75) or by infectivity assays ( 7 6 ) . I n all cases, “minus” strands either are present in the enzyme preparation or are formed during the early phase of synthesis (74, 77). In vivo, completed RNA strands are incorporated into virus particles after a delay of 10-15 minutes (31, 7 8 ) . Lodish et al. (50) demonstrated the existence of a maturation step in the late phase of virus synthesis: chloramphenicol or fluorouracil, administered to infected cells 20 minutes after infection, as well as certain mutants (sus-1, ts-16) grown under nonpermissive conditions, cause the formation of defective, nonviable virus particles deficient in RNA, although viral RNA synthesis itself is normal ( 5 0 ) . Similar particles are formed in the presence of proflavin (79). Four viral functions have been clearly defined by biochemical and genetic criteria: (1) synthesis of “minus” strands; (2) synthesis of “plus” strands; (3) synthesis of coat protein; (4) a maturation step. (1) and (2) are associated with the formation of one or two virusspecific polymerases. An analysis of MS2-infected, actinomycin-treated E . coli spheroplasts suggests that six different virus-specific proteins are produced in the infected host (37).

A. In Vivo Studies 1. FATEOF PARENTAL RNA

The interpretation of experiments designed to follow the fate of labeled parental RNA is complicated by the fact that the viability of RNA phages is often relatively low (20% and less), and that there may be some uncertainty as to whether a given labeled fraction is actually participating in the replicative process.

360

CHARLES WEISSMANN AND SEVER0 OCHOA

Davis and Sinsheimer (80) demonstrated that less than 3% of the radioactivity of labeled parental RNA is recovered in the progeny phage, and there was no indication that any of the RNA is transmitted to the progeny as an intact strand. On the other hand, the parental strand is conserved in the host cells in which it demonstrably initiates infection and it can be recovered intact a t the end of infection (81). Sedimentation analysis of the contents of host cells infected with P32-labeled phage showed that, within minutes after infection, the parental RNA strand was found mainly in association with the 70 S ribosomal fraction (81, 71) and/or with the polysome fraction (8.9). Similar analysis of the purified RNA from cells infected with PSzlabeled phage revealed a striking change in the physical state of the labeled parental RNA strand shortly after infection. Part of the labeled RNA (original s ~ = 27-28 , ~ S) sedimented broadly around 16 S (83, 7 1 ) .A considerable fraction of the 16 S RNA was ribonuclease-resistant1 (Fig. 2 ) . The conversion of the parental strand into what has been called “replicative intermediate” (71)can be followed by the appearance of ribonuclease-resistant parental RNA (69, 70, 83), which is first detectable around 5 minutes after infection (Fig. 1 ) . After 12 minutes, 12% of the input phage RNA is resistant to RNase. Since only 20% or less of the labeled phage is viable, 12% may represent a substantial fraction of the RNA actually initiating infection. After 12 minutes, the RNaseresistant fraction again decreases. As shown by Geidushek e t al. (84), resistance to RNase is a striking property of double-stranded RNA and, in fact, the isolated, RNase-resistant radioactive RNA showed all the properties expected of a double strand, namely: ( a ) a sharp thermal transition to a ribonuclease-sensitive state [103”C in 0.15 M sodium chloride, 0.015 M sodium citrate (SSC)] ( 7 0 ) ; ( b ) an s ~ , , , ~of about 13.4 S (86),a value compatible with double-helical RNA of molecular weight about 2 x lo6; and (c) a buoyant density about 0.02 gm/cm3 less than that of single-stranded RNA (70).Moreover, specific dilution assays (see Appendix) showed that the radioactive component of the double strand was indeed a viral “plus” strand. After heating and reannealing in the presence of an excess of the homologous unlabeled viral RNA (but not of any other RNA) , the radioactive strand was displaced from the double strand and thereby became RNase-sensitive. Erikson et al. (85) isolated the fraction of parental-labeled RNA sedimenting around 16 S and demonstrated that after heat denaturation a considerable amount of the radioactive RNA sediments a t 27 s, the characteristic value of single-stranded viral RNA. This shows that the parental RNA is

REPLICATION OF PHAGE RNA

361

[L

W

I-

LL

a

0 0 a

TIME AFTER INFECTION (min)

FIO.1. Formation of double-stranded MS2 RNA and “minus” strands following infection of E . coli with MS2 phage. The continuous curve (Pa-labeled parental RNA) illustrates the formation and subsequent decrease of ribonuclease-resistant double-stranded RNA containing P”-labeled parental RNA strands. The dashed curve showa that “minus” strands increase throughout infection. Most “minus” strands are recovered in a double-stranded form. Note that different ordinates are used for the two curves. The dotted line shows the time course of intracellular MS2 phage production (70).

conserved intact within the replicative intermediate and can be released from the duplex by denaturation. Appearance of the radioactive parental RNA in a double-stranded form is attributed to the synthesis of a complementary (or “minus”) strand, with use of the “plus” strand as template. This conversion is prevented by the addition of chloramphenicol to the bacterial culture immediately preceding infection, suggesting that the synthesis of a virusspecific enzyme is required for this step (83,7 1 ) . The decrease of the labeled double-stranded RNA later on (Fig. 1) can be explained by assuming either ( a ) that the double-stranded RNA is degraded or separated into two single strands or ( b ) that the parental strand is displaced from the duplex by a newly formed “plus” strand (see the scheme of semiconservative, asymmetric replication, Fig. 9). The kinetics of the process, in particular the slow rate of displacement, can be accounted

362

CHARLES WEISSMANN AND SEVER0 OCHOA

for if a parental strand, after being displaced from the duplex as a single strand, is again converted to a double-stranded form, and passes through several such cycles until it is finally trapped in a single-stranded state, possibly as the consequence of some discrete damage. The curve of Fig. 1 would then be the result of nonsynchronous cycling of a large population of parental strands ( 1 ) . Several observations support interpretation ( b ). When host cells are infected with Ps2-labeled sus-3, a mutant of f2 that causes the formation of excessive amounts of the enzyme responsible for the formation of “minus” strands, the amount of parental Pa2RNA converted to a doublestranded form increases throughout the period of observation and reaches 50% of the input after 1 hour (67). Since the double-stranded RNA accumulating under these conditions is defective (6 S rather than 14, 16 S), the result of this experiment can be explained by the assumption that the parental RNA is trapped in double strands that can no longer participate in replication because of their defectiveness. Moreover, i t is obvious that extensive intracellular degradation of double-stranded RNA to acid-soluble fragments, as called for by hypothesis ( a ) , does not occur. Lodish and Zinder (62) isolated a temperature-sensitive mutant of f2, ts-6, that neither showed parental strand conversion nor caused viral RNA replication within the host a t 43”C, although adsorption of the virus and penetration of the RNA took place normally. After transferring to 34”C, both the conversion to a double-stranded form, and, in due course, viral RNA synthesis occurred. This finding suggests that the processes associated with the conversion of the parental strand to a duplex are related to the early stages of RNA replication. Further experiments show that, in the case of this mutant, formation of “minus” strands but not that of “plus” strands is blocked a t high temperatures. When cells infected with Psz-labeled ts-6 were grown a t 34”C, allowing double-stranded, Ps2-labeled RNA to form, and were then transferred to 43OC, the labeled double-stranded RNA disappeared almost entirely within 3 minutes (86a). This means that, under conditions where “minus” but not “plus” strand synthesis was inhibited, parental Ps2-labeled strands were displaced from the duplex ,and were no longer reconverted to double strands. Using spheroplasts synchronously infected with Ps2-labeled virus, Kelly et al. (86) found that within the first few minutes of infection the labeled RNA periodically became resistant to RNase. At 90 seconds, 80%, at 150 seconds, 20%, and a t 210 seconds, again 70% of the parental RNA was RNase-resistant. However, this resistance was only detectable as long as the crude lysate was tested. Upon deproteinization, the RNA

REPLICATION OF PHAGE RNA

363

was completely sensitive to ribonuclease. The periodic appearance of ribonuclease resistance was correlated with the periodic appearance of a form of parental RNA that (after phenol extraction) sedimented a t 20 S rather than 27 S. This phenomenon appears to be unrelated to the conversion to a double-stranded RNA described above. The authors suggest that the 20 S form is a special conformation of the 27 S single-stranded RNA, with which it is interconvertible, and that the periodic RNase resistance observed in crude lysates may be explained by its periodic participation in a protein-synthesizing complex. These observations require further elaboration. No conversion of labeled parental RNA to a double-stranded form has been observed in the case of animal viruses (87-89). To recapitulate, parental phage RNA, after entering the host cell, is found in association with the ribosomal fraction. I n the first few minutes after infection, periodic events appear to take place that render the parental RNA resistant to RNase as long as it is examined prior to deproteinization ; after deproteinization, this RNA behaves like singlestranded viral RNA, except that its sedimentation coefficient is reduced to 20 S. This event may reflect processes associated with protein synthesis. After 5-6 minutes, parental RNA, in deproteinized extracts, is found in a form sedimenting a t 14-16 S, the replicative intermediate. Treatment of this fraction with RNase yields parental RNA in a doublestranded form. About 15 minutes after infection, the amount of parental RNA found in this form is maximal and decreases thereafter, possibly by displacement from the duplex (see p. 374). During the entire infective process, the parental RNA is conserved intact within the host and is not incorporated into progeny phage. 2. VIRUS-SPECIFIC RNA SPECIESFORMED IN BACTERIA INFECTED WITH RNA PHAGES

a. “Minus” Strands and R’eplicative Intermediate. As already mentioned, virus-specific double-stranded RNA is detected some 5 minutes after infection of E. coli with P32-labeled MS2 phage. As shown in Fig. 1, the process is followed through the conversion of the labeled parental strand from an RNase-sensitive to an RNase-resistant state. This shows that “minus” strands, i.e., strands with a base sequence complementary to that of the parental RNA, are formed. Annealing assays show that “minus” strands are synthesized in large excess over those associated with parental RNA strands (Fig. 1). Fifteen minutes after infection an MS2-infected cell may contain about 30, and 45 minutes postinfection as many as 500 equivalents of viral “minus” strands, most of which

364

CHARLES WEISSMANN AND SEVER0 OCHOA

are double-stranded. The bulk of the double-stranded RNA sediments around 16 S and, while some of it sediments more rapidly, very little sediments more slowly (Fig. 2). However, under certain conditions, such as conversion of infected cells to protoplasts followed by treatment with actinomycin (89, or after UV irradiation of the host cells (go), considerable amounts of double-stranded RNA with a sedimentation coefficient of about 6 to 7 S are found. The sedimentation coefficient of the double-stranded RNA present after infection with phage mutants such as mutant sus-3 of f2 (67) or mutant 9 of MS2 (91) is almost exclusively 6 to 7 S. I n view of the circumstances under which this material is found, its formation may reflect some abnormal process. This is

FRACTION

NUMBER

FIG.2. Conversion of parental P=-labeled R17 RNA into the replicative intermediate. RNA was isolated from E . coli infected with P" R17 at (a) 0, (b) 6, and ( c ) 12 minutes after infection and analyzed by sucrose density gradient centrifugation. - - .O - - 0 - -, total radioactivity; 0-0, RNase-resistant radioactivity -@-, absorbance. From Erikson et al. (71).

the ,more likely as the yield of virus, if any was formed a t all, was decreased under these conditions. Figure 3 shows the sedimentation profile of RNA from infected cells treated with RNase a t low and high concentrations prior to centrifugation. As described by Fenwick et al. (!V), mild RNase digestion causes the ribonuclease-resistant RNA to sediment somewhat more slowly in a narrow band a t about 12-13 S, whereas high concentrations of the en-

REPLICATION O F PHAGE RNA

365

zynie (50 pg/nil) cleave the double-stranded RNA to fragments sedimenting homogeneously a t 8-9 S. If double-stranded RNA has the same hydrodynamic behavior as double-stranded DNA, the relationship Sw,w = 0.0882 M”.”fi (96) suggests an s ? , , , ~of 13.4 S for a duplex of inolecular weight 2 X loo consisting of two full-length viral strands. It has been suggested (72) that the radioactive material sedimenting a t 14 S and above may consist of double-stranded RNA with one or more growing single “plus” strands attached (Fig. 4 ) . Mild ribonuclease treatment prior to sucrose gradient centrifugation would hydrolyze the single strands, thereby making the double-stranded RNA sediment more homogeneously and with a lower sedimentation coefficient. More vigorous digestion leads to cleavage of the double-stranded RNA itself. Lodish and Zinder (66) have presented evidence suggesting that the virusspecific 1 6 1 6 S RNA may in fact consist of two components: ( a ) a double-stranded molecule without single strands attached, sedimenting a t about 14 S, and ( b ) a double-stranded molecule with attached single strands, sedimenting a t 16 S and above. Component ( a ) would predominate after long periods of labeling and may represent double-stranded material not engaged in replication. Component ( b ) would be predominant after short labeling periods and may represent the actual replicating intermediate shown in Fig. 4. b. Other Virus-Specific RNA’s. The synthesis of virus-specific RNA has also been studied in cells in which host-specific RNA synthesis was blocked by actinomycin or UV irradiation (72, 86). These host-virus systems showed varying degrees of damage so that the observations must be viewed with some caution, particularly regarding the appearance of 6 S double-stranded RNA. Kelly et al. (86) examined RNA from actinomycin-treated infected spheroplasts by sucrose gradient sedimentation. They found four kinds of virus-specific RNA: a t 6 S, the “abnormal” double-stranded RNA ; a t 1&16 S, the “replicative intermediate”; a t 20 S, single-stranded RNA considered to be viral “plus” strands in a special conformation; and a t 27 S, normal viral RNA, a s it is found in the virus itself. These species of newly synthesized virus-specific RNA correspond to the various forms in which, as noted above, labeled parental RNA itself appears after infecting the host. The “replicative intermediate” seemed to consist of both single- and double-stranded RNA, as judged by its buoyant density (86) and by the fact that treatment with very small amounts of ribonuclease degraded about 30% of the radioactive RNA to acid-soluble fragments. The undegraded fraction sedimented a t 12 S (76)and had a lower buoyant density, characteristic of double-stranded RNA (86).

14 S

(a) RNase

FRACTION

0.5 j.iq/rnl

NUMBER

13 S

(b)

5 0 pg/rnl

RNase 8 S

(c)

-

3

.c

E

I

(...O...) (96).

cells, labeled with C"-uracil, was treated (a) with D N w alone, (b) with DNase and 0.5 pg/ml RNase, ( c ) with DNase and 50 pg/ml RNase. Alter phenol extraction, the samples were centrifuged through a aucrose density gradient, with E. coli riboRadioactive double-stranded RNA (+); radioactivity due to fragmented DNA somal RNA added as a marker (-).

FIG.3. Effect of RNase on the sedimentation behavior of double-stranded MS2 RNA. Nucleic acid from MS2-infected

No RNasc

0

a!

M

5

Z

$

367

REPLICATION O F PHAGE RNA

The question has been raised whether the “minus” strand in the duplex is continuous and of the same length as the “plus” strand, or (bl

(C)

RNase

50 y g h l

FIG.4. Schematic representation of the replicative intermediate : (a) as isolated after deproteinization [modified from Fenwick et al. (72’11; (b) after mild treatment with RNase the “tails” are cleaved off; (c) after intensive RNase digestion, the duplex is fragmented.

whether it consists of a series of short, complementary segments basepaired with the “plus” strand (93). I n order to decide between these alternatives, labeled 14-16 S RNA, containing replicative intermediate, was isolated and sedimented through a sucrose gradient either before or after heat denaturation. Labeled “minus” strands were localized by annealing each fraction with an excess of nonlabeled “plus” strands and determining the acid-insoluble radioactivity. As shown in Fig. 5, “minus” strands were found in the 14-16 S region in the case of nondenatured sample and in the 27 S region after denaturation. This shows that the “minus” strands within the duplex are for the most part full-length viral strands, c. Properties of Purifid Double-Stranded RNA. As already mentioned, part of the so-called replicative intermediate consists of ribonuclease-resistant RNA. Large-scale purification of this fraction is usually carried out after phenol extraction followed by treatment with DNase and pancreatic RNase in solutions of moderate ionic strength. The RNase-resistant RNA is isolated by exclusion chromatography on Sephadex G-200 (94, 96). Further purification may be obtained by

368

CHARLES WEISSMANN AND SEVER0 OCHOA

cesium sulfate density gradicnt centrifugation or by column chromatography on niethylated albumin-silicic acid. Several modifications of this basic procedure have been employed. Langridge et al. (96‘)used Mg++ to coprecipitate double-stranded MS2 RNA with the phage-induced enzyme, RNA synthetase, prior to phenol extraction and enzymatic

..

I

‘gs $TOTAL

a FRACTION

FIQ.5. Demonstration that the replicative intermediate contains a full-length “minus” strand. Labeled RNA from MS2-infected cells was centrifuged through a sucrose density gradient (top) and the fractions sedimenting around 16 S, containing the replicative intermediate, were pooled and recentrifuged directly (middle) or after heat denaturation (bottom). Total = total radioactive nucleic acids; Minus = “minus” strands, as detected by annealing with excess unlabeled MS2 RNA. Heat denaturation of double-stranded RNA (~m,,., 16 S ) liberates a “minus” strand sedimenting around 27 S.

-

digestion. Kaerner and Hoffmann-Berling (97) coprecipitated the doublestranded RNA with DNA using ethanol and then digested the sample with DNase and RNase. Advantage has been taken of the fact that a considerable fraction of ribosomal and single-stranded viral RNA may be precipitated by 1.5 M sodium chloride leaving double-stranded RNA in solution (94). Ammann ‘et ,aZ.(98) followed this step by chromatography on methylated albumin and avoided the use of ribonuclease alto-

369

REPLICATION O F PHAGE RNA

PROPERTIES O F

TABLE I MS2 RNA, DOUBLE-STRANDED MS2 RNA, A N D RIBOSOMAL RNA (96) MS2 RNA or R17 RNA

Par am eter

-

Phosphorus (%) Orcinol test (A166 units per p mole of purine) Cysteine test e(P)260 m p

Base composition (%) Buoyant density in CSZSOI szo,w (high ionic strength)

8600s A, 23.7a U, 24.4 G, 27.1 C, 24.8 1.626

Double-stranded MS2 RNA

11.8

Less than 2 % DNA 6670 24.6 (calc. 24.05) 23.7 (calc. 24.05) 25.9 (calc. 25.95) 25.8 (calc. 25.95) 1.609

7450' 25.1 20.4 32.6 21.9 1.646b

26 S (0.2 M NaCl)c 8.45 S (SSC) 20.4 S (0.02 M NaC1)c

8.1 S (0.01 SSC)

T, (high ionic

47 (SSC)

103 (SSC)

strength) ("C)

-

T, (low ionic strength) ("C) Hypochromicity ( %) X-ray diffraction pattern

-

9.58 (calc. 9.46) 11.8

sz0,,,,

(low ionic strength)

E. coli ribosomal RNA

87 (0.1 X SSC)

-

24 S, 18 S (0.1 M KC1, 0.05 M Tris)b 19 S, 15 S (0.005 M KCI, 0.0025 M Tris)' 54 (0.1 M sodium phosphate buffer)O

-

18

26.5

21.9

Not oriented

Highly orientedd

Not oriented

J. H. Strauss, Jr. and R. L. Sinsheimer, J . MoZ. Biol. 7 , 43 (1963). W. M. Stanley, Jr., Thesis. University of Wisconsin (1963). R. Gesteland and H. Boedtker, J . Mol. B i d . 8, 496 (1964). d R . Langridge, M. A. Billeter, P. Borst, R. H. Burdon, and C. Weissmsnn, Proc. Natl. Acad. Sci. U.S. 63, 114 (1964). a

b

gether. An analytically pure preparation of double-stranded viral RNA from MS2-infected E. coli has been described in detail ( 9 5 ) . Table I shows some of the analytical data on this material. The base composition is that expected of an RNA duplex consisting of an MS2 RNA strand and its complement and is very different from that of the host cell RNA. The molar absorbance, c (P), is lower than that of single-

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CHARLES WEISSMANN AND SEVER0 OCHOA

stranded RNA ; the absolute and relative (thermal) hypochromicity is higher. The T, of the helix-coil transition is dependent on the ionic strength and is substantially higher, under comparable conditions, than that of single-stranded RNA and higher than that of native DNA with a similar base composition. The X-ray diffraction pattern is well-resolved and characteristic of a double helical structure (96). The purified doublestranded RNA was fairly homogeneous in size, as revealed by ultracentrifugation studies. The sedimentation coe5cient of various samples ranged from 8 to 9.5 S, which corresponds to a molecular weight of 450,000-750,000 under the assumption that the Studier (92) relationship between sedimentation coefficient and molecular weight of DNA is valid for double-stranded RNA. Examination of the preparation in the electron microscope by Dr. A. K. Kleinschmidt revealed filaments similar in width and stiffness to those of double-stranded DNA or doublestranded reovirus RNA with a most frequent length of about 0.2 p. Assuming a value of 2.3 X lo6 per micron of double-stranded RNA (99, 100) this corresponds to a molecular weight of 460,000, in good agreement with the value determined by sedimentation. Since the molecular weight of MS2 RNA is about lo6 ( 6 ) ,the purified double-stranded RNA is thus between one quarter and one fifth of the size expected for a duplex consisting of an intact MS2 RNA strand and its complement. The small size of the double-stranded RNA is the consequence of treatment with high concentrations of ribonuclease in the purification procedure as indicated by the results described in the previous section. Smallscale preparations of double-stranded RNA were obtained by digesting RNA from infected cells with RNase a t low concentrations (0.1-1 pg/ ml) and isolating the 13 S peak after sucrose gradient centrifugation (cf. Fig. 3). They showed a most frequent filament length of 0.8-0.9p , corresponding to a molecular weight of about 2 X 106 (la,101). The purification method of Ammann (et al. (98) yielded a large proportion of full-length filaments as judged by electron microscopy. This preparation was noninfective in its native state ; however, upon thermal denaturation infect,ivity was demonstrable. I n conjunction with experiments described above, this suggests that double-stranded RNA isolated without RNase treatment contains full-length viral “plus” and “minus” strands. The resistance of purified double-stranded RNA to pancreatic ribonuclease A and B, as well as to ribonuclease TI, is very pronounced in 0.15 M sodium chloride but not a t low ionic strength. After heat denaturation, double-stranded RNA is as susceptible t o pancreatic ribonuclease as single-stranded RNA. Reannealing occurs readily and almost quantitatively a t 80-9OoC a t sodium chloride concentrations above 0.15

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M . Reannealed double-stranded RNA possesses the same properties as the native material (96,86).

3. VIRUS-SPECIFIC RNA SPECIES FORMED IN ANIMALAND PLANT TISSUES INFECTED WITH RNA VIRUSES The formation of virus-specific double-stranded RNA was first demonstrated in Krebs I1 ascites cells infected with encephalomyocarditis (EMC) virus, which contains single-stranded RNA (94). This doublestranded RNA was found to be infectious even after treatment with RNase. Since the virus causes the synthesis of an RNA polymerase in the infected host (102,103), it is very probable that encephalornyocarditis viral RNA functions as a messenger. Inasmuch as doublestranded RNA cannot serve as a messenger (log), the mechanism by which double-stranded EMC RNA initiates infection is obscure. Cell RNA polymerase might utilize this double-stranded RNA as a template for the synthesis of viral “plus” strands, or the component strands of the duplex might become separated, yielding messenger “plus” strands and “minus” strands. No infectivity has been demonstrated so far for double-stranded phage RNA (98). Double-stranded polio RNA was detected by Baltimore et al. (106) and characterized in terms of its annealing behavior (106) and base composition (107).Infectivity of double-stranded foot and mouth disease RNA (108) and of double-stranded polio RNA (109) has been reported. I n cells infected by mengovirus, specific double-stranded RNA was detected by Homma and Graham (89) but not by Tobey (87). The formation of virus-specific double-stranded RNA has also been reported in cells infected with Maus-Elberfeld (ME) virus (110), Semliki virus ( I l l ) , turnip yellow mosaic virus (TYMV) (112), and tobacco mosaic virus (TMV) (113, 11.4). Failure to find double-stranded RNA in cells infected with Western equine encephalitis virus has been reported (115). 4. SHORT-TERM LABELINGEXPERIMENTS

It is postulated that a structure consisting of enzyme and viral “minus” and viral “plus” strands occurs as an intermediate in RNA replication. After isolation involving deproteinization, this structure is recovered as the so-called replicative intermediate already described, which may consist of an RNA double strand with single strands attached. Several lines of evidence suggest that double-stranded RNA or a structure giving rise to double-stranded RNA upon isolation is involved in viral RNA replication*: ( a ) In MS2-infected bacteria the formation of viral “minus” precedes that of “plus” strands (73). ( b ) As

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CHARLES WEISSMANN AND SEVER0 OCHOA

described in a previous section, radioactive parental RNA is converted to a double-stranded form within a few minutes after entering the host cell. (c) A complex involving “minus” strands appears to be involved in the in vitro synthesis of viral “plus” strands by RNA synthetase (74, 75). Findings with the in vitro system and the fact that radioactive parental RNA converted to a double strand is subsequently displaced from the duplex led to the suggestion that double-stranded RNA, or a structure giving rise to double-stranded RNA upon isolation, is an intermediate in viral RNA replication (74). Short-term labeling experiments lend strong support to this idea. Fenwick et al. (1964) (72) labeled with radioactive uridine R17infected bacteria in which host RNA synthesis had been inhibited by irradiation with ultraviolet light. Upon sucrose gradient centrifugation of the isolated RNA, the label was found almost exclusively in the 16 S region, corresponding to the replicative intermediate. If cells thus labeled for 10 seconds were subsequently incubated in unlabeled medium (chase), there was a marked decline of ribonuclease-resistant label within the 16 S fraction, which originally was 70% ribonuclease-resistant, while 27 S radioactive RNA, characteristic of viral RNA, appeared. This suggested a precursor-product relationship between the 16 S and the 27 S RNA. These experiments were confirmed and extended by Billeter et al. (73). Following exposure of MS2-infected E . coli to C14-labeled guanine for 8 seconds, the distribution of radioactivity between total and RNaseresistant RNA was determined both immediately after the pulse and at various times after terminating the pulse by the addition of excess unlabeled precursors. Whereas immediately after the pulse about 25% of the total acid-insoluble radioactivity was in RNase-resistant RNA (it should be noted that host RNA synthesis proceeds actively in these cells), this value dropped to about 9% during the first 2 minutes after quenching and to 6.8% after 7 minutes (Fig. 6). The loss of radioactivity from pulse-labeled double-stranded RNA was not due to a breakdown of the duplex, since the sedimentation pattern of the ribonuclease-resistant RNA did not change a t any time after the pulse, nor was it caused by separation of the two strands of the duplex because, as shown below, the radioactive “minus” strand was conserved within the double-stranded structure. Loss of radioactivity was due to displacement of labeled “plus” strands from the duplex. The proportion of radioactive viral “plus” and “minus” strands in double-stranded RNA was determined with use of the specific dilution assay (see Appendix), ( a ) immediately after an 8-second labeling period and ( b ) a t various t’imes after dilution of the label. The results of the dilution assays are shown in Fig. 7. It may be seen that (a)after brief

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exposure to labeled precursors, about 87% of the ribonuclease-resistant radioactivity was in viral “plus” strands and ( b ) upon dilution of the label, this value fell to about 60 and 40% after 2 and 12 minutes, respectively, indicating displacement of the “plus” strand from the duplex. The internal controls added to the dilution assay, consisting of double-

I

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FIQ.6. Decrease of labeled double-stranded RNA formed during a short pulse of C’4-guanine. MS2-infected cells were exposed to C“-guanine far 8 seconds. RNA was prepared from samples taken immediately after the pulse and at different times after dilution of the label. Double-stranded RNA was determined as RNaseresistant radioactivity (73).

stranded RNA uniformly labeled in both strands with P32, showed, as expected, 50% of the label in “plus” strands. For further analysis, the proportion of viral “plus” strands present both in double- and in single-stranded RNA a t various times after the pulse of radioactive precursors was determined by means of the double isotope specific dilution assay (see Appendix). The results of a typical experiment are summarized in Fig. 8. At the termination of the pulse some 60% of the labeled “plus” strands were in double-stranded RNA. This value dropped to about 8% in the following 4 minutes as the proportion of labeled f‘plus” strands in double-stranded RNA decreased. “Minus” strands were synthesized during the labeling period and appeared largely in double-stranded RNA. However, in contrast to the “plus” strands, the “minus” strands showed no significant turnover. The

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CHARLES WEISSMANN AND SEVER0 OCHOA

ratio of total “plus” to total “minus” strands synthesized was about 10. These findings are compatible with the mechanism outlined in Fig. 9. It is suggested that replication of phage RNA proceeds in two steps:

10

20

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20

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20

MS2 RNA ADDED ( pg FIQ.7. Distribution of radioactivity between the “plus” and “minus” strands of double-stranded RNA immediately after a pulse of CX4-guanineand at various times after dilution with unlabeled guanine. C“-Labeled double-stranded RNA was prepared from the pulsed cells at the times indicated and analyzed by the specific dilution assay. Samples were heated and renatured in the presence of increasing amounts of unlabeled MS2 RNA. The radioactive RNA remaining RNase-resistant at “infinite” concentrations of MS2 RNA (see extrapolation in inset) is due to labeled “minus” strands ; the radioactive RNA converted t o an RNase-sensitive form is due to “plus” strands. Double-stranded RNA, uniformly labeled with Pa’ (containing equal amounts of labeled “plus” and “minus” strands), was added as an internal standard to each sample (73).

(a) synthesis of “minus” strands using “plus” strands as template; and ( b ) synthesis of progeny ‘‘plus” strands by an asymmetric semiconservative process in which a complex of “minus” and “plus” strands is an intermediate. If a strictly semiconservntivc mcchnnism werc opcrntive in step ( b ) , all of the viral RNA labeled upon exposure of infected cells to radioactive RNA precursors, for a time shorter than required to displace the equivalent of one viral “plus” strand from the duplex, should be recovered in double-stranded RNA. In other words, there should be no free labeled “plus” strands after a sufficiently short pulse. However, reduc-

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375

FIQ.8. Distribution of radioactivity between “plus” and “minus” MS2 RNA strands immediately after a pulse of C”-guanine, and at various times after dilution of the label. Solid or shaded areas, radioactivity in double-stranded RNA; nonshaded areas, radioactivity in single-stranded RNA. Immediately after the pulse, 60% of the “plus” strands are found in a double-stranded form; subsequently “plus” strands are displaced from the duplex and less than 6% remain double-stranded. The proportion of “minus” strands in a double-stranded form remains constant (73).

FIQ.9. Scheme of viral RNA synthesis and the fate of labeled precursors. Two steps in synthesis are postulated: A, synthesis of a “minus” strand leading to the formation of a replicating complex; B, synthesis of “plus” strands by the replicative complex, by a mechanism in which newly formed strands displace their counterparts from the complex. After short labeling C”-guanine is incorporated into “plus” and “minus” strands in the ratio 1011. After dilution of the radioactive label, labeled “plus” strands, but not labeled “minus” strands, are displaced from the complex.

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CHARLES WEISSMANN AND SEVER0 OCHOA

tion of the labeling time to 2 seconds gave essentially the same results as labeling for 8 seconds; the proportion of plus strands in doublestranded RNA did not exceed 5040%. These findings could be explained if some of the newly formed “plus” strands in step ( b ) displace their counterparts from the duplex and some do not. If the component strands of the duplex dissociate as synthesis of a new “plus” strand proceeds through base pairing with a “minus” strand, reformation of the duplex on completion of the new “plus” strand might occur randomly with equal chance for displacement of either the old or the new strand. Such a mechanism has been suggested (118, 117) to explain the finding that, when a DNA-RNA hybrid template directs the synthesis of the complementary RNA of +X174 by RNA nucleotidyl transferase, about half of the RNA in the hybrid is displaced after one round of synthesis. Alternatively, if hydrogen bonding between the “plus” and the “minus” strands of the replicating complex occurs only during the isolation procedure, the randomization may take place a t this stage. Another point deserving comment is that the displacement of the radioactive “plus” strands from the duplex was incomplete even 12 minutes after termination of the pulse. It is assumed that this is, due to conversion of some of the released labeled “plus” strands to doublestranded material through the continuing synthesis of new “minus” strands. Experiments by Lodish and Zinder (62) cast further light on the role of double-stranded RNA. As already mentioned, E . coli infected with ts-6, a mutant of the RNA phage f2, synthesizes double-stranded RNA, viral RNA, and virus a t 35°C but not at 43°C. However, once viral replication is initiated and some double-stranded RNA formed a t 35”C, viral “plus” strand synthesis (but not synthesis of double-stranded RNA) occurs also a t the higher temperature. This mutant then appears to induce the synthesis of a temperature-sensitive step (1) enzyme and a stable step (2) enzyme. Short-term labeling experiments carried out at 35°C gave similar results as those described above. However, a t 43°C where no further “minus” strands are synthesized, displacement of radioactivity from double-stranded RNA is virtually complete. There is some evidence that the double-stranded RNA of TYMV is involved in viral RNA turnover. Mandel e t al. (112) found that, after short-term labeling of Chinese cabbage leaves infected with TYMV, the base composition of the radioactive RNA in the RNase-resistant fraction is similar to that of TYMV “plus” strands, whereas, after long labeling periods, the analysis is that expected of a TYMV duplex. I n the case of animal viruses, the involvement of double-stranded RNA has been suggested by the finding (110) that double-stranded Maus-

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Elberfeld (ME) RNA is preferentially labeled in the “plus” strand after short-term labeling experiments. 5. SUMMARY

Within minutes after infection, part of the parental RNA entering the host is converted to a replicative intermediate characterized by ( a ) its sedimentation behavior and ( b ) its partial RNase resistance. It has been suggested that this intermediate consists of double-stranded RNA with growing single-stranded tails attached to it (Fig. 4). Subsequently, additional intermediate is made de novo and several hundred viral double strands finally accumulate in the infected cell. Short-term labeling experiments show that a t least part of the double-stranded RNA turns over during the period of active “plus” strand synthesis. At this time, radioactive precursors are incorporated into “plus” strands of double-stranded RNA much more rapidly than into “minus” strands, and dilution of the radioactive label results in the displacement of “plus” but not of “minus” strands from the duplex. These findings suggest that a substantial fraction of “plus” strands is formed by a semiconservative, asymmetric replication mechanism. It is possible that a t any one time only a limited number of double-stranded RNA molecules is involved in replication while the remaining molecules are either discarded or kept in reserve and do not form part of a replicating complex. Furthermore, it is conceivable that the replicating complex, as i t occurs in vivo, may not have the tightly hydrogen-bonded, RNase-resistant structure observed after its isolation (118). An analogous structure, intermediate between native and denatured, has been considered for replicating DNA (119, 120).

Three other forms of RNA related to the virus can be detected by sedimentation analysis: (1) a double-stranded RNA sedimenting a t about 6 S, formed only under certain conditions and possibly an abnormal product; (2) a single-stranded RNA sedimenting a t 20 S, thought to be viral RNA of a special conformation; (3) viral RNA sedimenting a t 27 S.

B. Enzyme Studies 1. FORMATION OF NEW RNA-SYNTHESIZING ENZYMES IN PHAGE-I N FECTED BACTERIA Following infection with RNA phages, a new RNA-synthesizing activity is detectable in the host cell (41-43). Using as an assay the incorporation of CI4-GMP from CZ4-GTF’into acid-insoluble material in

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CHARLES WEISSMANN AND SEVER0 OCHOA

the presence of DNase, an MS2-induced enzyme system is detectable about 6 minutes after infection and reaches a maximum a t about 30 minutes (121). Similar kinetics were observed by August et al. (42) with phage f2. Lodish et al. (56) showed that chloramphenicol inhibits the appearance of phage-induced polymerase (s), when added within 10 minutes after infection, and halts further formation of enzyme whenever added. Several amber mutants of RNA phages (sus-3, sus-11, mu-9) cause the formation of 5 to 15 times the normal amount of both viral polymerase activity and double-stranded RNA, when grown on certain nonpermissive strains of E . coli (56, 57, 9 1 ) . I n these cases, the rate of enzyme formation is increased to a variable degree and, above all, synthesis continues beyond 60 minutes after infection, whereas i t normally ceases after 25-30 minutes. It is remarkable that the lesion in the three mutants just mentioned is localized in the coat protein cistron (52). Since the mutants appear to be single mutants, this implies that the increased enzymatic activity resulting in the host is due to disturbed control either of the synthesis of the enzyme or of its activity while the actual enzyme protein or proteins are normal. It was suggested (66),on the basis of experiments in which viral RNA synthesis was inhibited by precursor deprivation, that, in the case of wild-type phage, only the parental strand and perhaps some early progeny strands are utilized as messenger for the synthesis of the polymerase(s), while in the case of the hyperproducing mutants, late progeny RNA strands also serve this function, leading to an excessive production of enzyme. 2. PROPERTIES OF INDUCED ENZYMIM

Three laboratories have reported on the isolation and partial purification of phage-induced RNA polymerases. The properties of these preparations are different, so that each preparation is described separately. An attempt is made to correlate the findings. a. R N A Synthetase. Weissmann et al. (41, 70) isolated an RNAsynthesizing enzyme from E . coli infected with MS2 phage. After partial purification (about twentyfold), the preparation was free of DNAdependent RNA nucleotidyl transferase. All four nucleoside triphosphates were required for maximal rate of RNA synthesis. The preparation contained substantial amounts of endogenous RNA and in particular viral “minus” strands, most of which were presumed to occur in a doublestranded form. There was no stimulation by the addition of MS2 RNA or of any other RNA and treatment with RNase and subsequent removal of the nuclease led to irreversible inactivation of the enzyme. It was therefore concluded that the enzyme contained an endogenous template, presumably the viral “minus” strands.

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Radioactive RNA was synthesized with the partially purified enzyme, using radioactive nucleoside triphosphates as substrates. The base composition of the product was similar to that of MS2 RNA (1). Analysis by the double isotope specific dilution assay (Fig. 10) showed that the product was entirely virus-specific, consisting of 90% viral “plus” and about 7% “minus” strands (76). Only a small part of the product of RNA synthetase was resistant to RNase when tested prior to deproteinization ; however, following extraction with phenol, or treatment with sodium dodecylsulfate, over 50% of the product was RNaseresistant (118). This was particularly evident when crude preparations of RNA synthetase were used to prepare the product. The RNaseresistant fraction of the enzymatic product isolated after phenol extraction is indistinguishable from the MS2-specific double-stranded RNA found in vivo in its RNase resistance, T,, buoyant density, and annealing behavior. When subjected to the specific dilution assay, the radioactivity in the duplex is almost completely displaceable and is therefore present in “plus” strands ( 7 0 ) .It was therefore concluded that the nonlabeled “minus” strands in the duplex were present in the enzyme preparation prior t o the incubation with radioactive triphosphates. As already ,mentioned, the enzyme preparation in fact contained substantial amounts of viral “minus” strands. The above results suggest that RNA synthetase is associated with an endogenous template, namely viral “minus” strands either in a single- or double-stranded form, and that it catalyzes the synthesis of labeled viral rrplus”strands on incubation with radioactive triphosphates. The fact that, after phenol extraction, some of the newly formed radioactive single “plus” strands are found in a double-stranded state, whereas added radioactive viral RNA is not, suggests that the newly formed “plus” strands and the “minus” strands are held in proximity, possibly by the enzyme, so that annealing of the complementary strands is greatly facilitated (118). b. Viral R N A Polymerase. August et al. (122) utilized the nonpermissive host E. coli S26 infected with sus-11, an amber mutant of phage f2, as a source of enzyme. This host-phage system does not yield phage particles but it produces 5-8 times the normal amount of viral RNA polymerase (s) and virus-specific double-stranded RNA (66). The 100fold-purified preparation required RNA as a primer. A variety of RNA’s, such as f2, TMV, ribosomal, or soluble RNA, were about equally active but homopolynucleotides were not active. The nearest-neighbor frequency of the product was different for TMV or f2 RNA primer, suggesting that the added RNA determined the nucleotide sequence of the product. Its base composition, and the finding that, after deproteiniza-

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CHARLES WEISSMANN AND SEVER0 OCHOA

tion, more than half was resistant to RNase (123),indicated that it was a complementary or “minus” copy of the template. It was suggested that the enzyme functions in vivo to catalyze the synthesis of a (“minus”) strand complementary to the viral (“plus”) parental strand. Mug, an amber mutant of phage MS2, induces, like sus-11, the production of large amounts of polymerase and double-stranded RNA when grown on a nonpermissive host such as E. coli Hfr 3000 (91). Annealing assays showed that as much as 60% of the product formed in vitro by extracts of Mug-infected Hfr 3000 consisted of viral “minus” strands, in marked contrast to the results with extracts of cells infected with the

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wild-type phage (MS2) in which the corresponding value was 15%. These findings support the conclusions of Shapiro and August (123). c. R N A Replicme. Haruna e t al. (43, 124) isolated from E . wli infected with phage MS2 an enzyme that, under appropriate ionic conditions, specifically required the addition of MS2 RNA for activity. Ribosomal, soluble, and TMV RNA were ineffective but TYMV RNA caused some stimulation. Significantly, the RNA of phage Qp [which shows no

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FIG. 10. Identification of the radioactive product of RNA synthetase as viral RNA of the parental type. (a) When a mixture of Pa‘MS2 RNA is heated and reannealed with an excess of unlabeled double-stranded MS2 RNA, most of the P3’ RNA is converted to an RNase-resistant form (curve 1). Ci4-labeled product of RNA polymerase, primed with E . coli DNA, was also present in the mixture and remained RNase sensitive (curve 2). Ribosomal RNA, in place of double-stranded MS2 RNA, had no effect (inset), (b) When the experiment was carried out as above, but using the Ci4-1abeled product of RNA synthetase, both the Pa2-labeled (curve 1) and the C”-labeled (curve 2) RNA were converted to a double-stranded form to an equal extent (some radioactive RNA is RNase-resistant without addition of double-stranded RNA, due to unlabeled “minus” strands derived from the enzyme preparation). (c) A mixture of Ps2 MS2 RNA and C“-labeled product of RNA synthetase was converted into a double-stranded form as described under (b) and subjected t o the specific dilution assay: it was heated and reannealed in the presence of increasing amounts of MS2 RNA. P”-labeled (curve 1) and Ci4-labeled (curve 2) RNA were displaced from the duplex and converted to an RNase-sen& tive form, identifying them as MS2 “plus” strands. There was no effect of ribosomal RNA (curves 3 and 4). The combination of the steps of Figs. lob and 10c constitutes the double isotope specific dilution assay. When the fraction of RNaseresistant C“ radioactivity (fo) is plotted against the corresponding value for P” ( f ~ ) ,a straight line results. The fraction of radioactivity in “plus” st,rands is given by the slope, that in “minus” hy the intercept (cf. Appendix) (76).

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CHARLES WEISSMANN AND SEVER0 OCHOA

base sequence homology with MS2 RNA (It?)] did not stimulate the MS2 replicase. Moreover, a replicase isolated from E . coli infected with phage Qp was stimulated by intact Qp RNA but not by MS2 RNA. The activity of Qp replicase was not much enhanced by partially degraded Qp RNA. As in the case of the MS2-induced enzyme, i t was slightly stimulated by TYMV RNA. I n contrast to the two enzyme preparations already described, Q p replicase catalyzes the synthesis of RNA over several hours, producing several times the amount of input RNA, and is saturated a t relatively low ratios of RNA to protein ( 1 pg/40 pg protein). The template specificity of the Qp enzyme is modified by the addition of Mn2+to the Mgz+containing reaction mixture and, even more, by the substitution of Mn2+ for Mg2+.Under these conditions, the effectiveness of Qp RNA is markedly reduced while the effect of fragmented Qp RNA and of heterologous RNA’s, such as TYMV RNA, increases. It was suggested (125) that recognition of the template RNA by replicase depends on the over-all configuration of the RNA (hence the need for intact Qp RNA) and on the presence of a special nucleotide sequence a t one or both ends of the molecule. I n the presence of nonsaturating amounts of intact template RNA, the synthesis of RNA by replicase exhibits autocatalytic kinetics (126). This suggests that the reaction product is Qp RNA and functions as template. Indeed, after long incubation periods, when the amount of RNA synthesized greatly exceeds that originally added as template, the isolated product was ( a ) as effective a template of the Qp-induced replicase as Qp RNA itself, and ( b ) infective for E. coli spheroplasts (Fig. 11) ( 7 6 ) . The possibility that the latter results were due to an enhancement of the infectivity of the RNA used as template was ruled out by experiments involving repeated transfers of part of the product of an incubation to fresh incubation mixtures. Thus the original RNA template was diluted to less than one strand per tube in the fifteenth transfer, without decreasing the concentration of infectious units ( 7 6 ) . In contrast t o the reaction just described, an abnormal synthesis results when Qp replicase is primed with partially degraded Qp RNA. The rate of synthesis is lower, its extent is limited, and the product is largely double-stranded (125, 127). It is clear from the above results that the replicase system is capable of synthesizing viral RNA of the parental type. The question therefore arises whether this replication involves preliminary formation of viral “minus” strands as suggested by the in vivo experiments and by the results with other enzyme preparations, mentioned earlier. Annealing assays of the radioactive product isolated after various times of incubation of Qp replicase with intact Qp RNA (77) showed that “minus”

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REPLICATION OF PHAGE RNA

strands were predominantly, if not exclusively, synthesized during the first few minutes but that mainly ‘Lplus”strands were formed later on (Fig. 12).2Twenty per cent or more of the product was RNase-resistant lx RNA

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FIG. 11. Synthesis of RNA and formation of infective units by QB replicase. Qp replicase was incubated with Qa RNA as template and with radioactive nucleoside triphosphatcs. Aliquots were assayed for acid-insoluble radioactivity and for infectivity in a spheroplast assay system. From Spiegelman et al. ( 7 6 ) .

throughout incubation, but the bulk of the “minus” strands was RNasesensitive and demonstrable only through annealing with Qp RNA. *It has been suggested that the ends of the Qa RNA molecule could be complementary to each other (126, In). If this were the case, the simple appearance of radioactive RNA capable of annealing with “plus” strands would not necessarily signal the formation of “minus” strands; in fact, early in the reaction, when only short segments of RNA have been synthesized, a “plus” strand of this nature could not be distinguished from a “minus” strand by annealing techniques. However, after a long-term incubation, 30% of the newly synthesized RNA sedimenting with 27 S (i.e., full length strands) is nnnenlnblr with ‘‘plus” strands (77). If this were due to self-complcmentary, then authentic viral RNA stioulcl be self-annealable to the same extent. Haruna and Spiegelman (127) claim that up to 6% of QS RN.4 can be annealed with itself. Using carefully purified P”-labeled viral RNA, we have not found any self-annealing (less than 0.2ol), either in the case of MS2 RNA (70, 79) or Qa RNA ( 1 2 7 a ) . Other arguments (127) attempting to attribute the appearance of anncalable viral RNA during in vitro synthesis to self-complerncntary regions of “plus” strands remain speculative a t present.

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CHARLES WEISSMANN AND SEVER0 OCHOA

Sucrose gradient centrifugation of the product present after incubation for 40 minutes (Fig. 13) showed that about 30% of the radioactive strands sedimenting in the 27 S region (corresponding to full-length single-stranded viral RNA) were “minus” strands. These results suggest that the synthesis of two products is catalyzed by replicase preparations, although it is not known whether one or two enzymes are involved. It may be noted that, as is the case with the other enzymes of viral RNA replication, RNA replicase has thus far been only partially purified.

3. RNA-SYNTHESIZING ENZYMES IN ANIMALAND PLANT CELLS INFECTED WITH RNA VIRUSES Baltimore and Franklin (198) first demonstrated the induction of an RNA polymerase activity by RNA viruses using L cells infected with mengovirus. Analogous observations were later made (199) with HeLa cells infected with poliovirus. The enzymatic activity was associated with a particulate cytoplasmic fraction which, on incubation with

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REPLICATION O F P H A G E R N A

poliovirus RNA, along with some double-stranded RNA (130). The presence of a virus-induced polymerase in Krebs 11 ascites cells infected with encephalomyocarditis virus was suggested by the work of Cline et al. (102) and confirmed by Horton et al. (103). Similar findings were made with influenza virus (131) and vesicular stomatitis virus (132). As in the case of poliovirus-induced polymerase, both single- and doublestranded RNA are formed in witro by the encephalomyocarditis system (133). In no case has an enzyme been purified or an RNA dependence demonstrated directly. A virus-induced enzyme has also been reported in TYMV infection (134,136). A number of authors have reported the synthesis of infectious viral RNA or even of intact virus in crude enzyme systems from animal and plant tissue (136-143). These reports must be viewed with great caution for, as pointed out by Ralph and Matthews (144), viral RNA may be released from damaged virus particles during the incubation, thus in-

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creasing the infectivity of the extracts, to mention only one possible source of error.

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CHARLES WEISSMANN AND SEVER0 OCHOA

4. SUMMARY

Three enzyme preparations derived from E . coli infected with RNA phages have been described in the preceding sections: RNA synthetase, viral RNA polymerase, and RNA replicase. RNA synthetase is associated with an endogenous template (viral “minus” strands in single- or double-stranded form) and synthesizes viral “plus” strands in vitro, identified by annealing techniques. RNA polymerase is RNA-dependent. It utilizes a variety of exogenous RNA’s, including nonviral RNA’s, as primers and appears to synthesize predominantly “minus” strands, i.e., strands complementary to the priming RNA “plus” strands. RNA replicase is also RNA-dependent but it functions specifically with the RNA of the phage that induces the formation of the enzyme. The preparation catalyzes the replication of its specific viral RNA with an early production of viral “minus” strands followed by a later, much greater formation of viral “plus” strands. As mentioned in a previous section, two separate steps appear to be involved in the replication of viral RNA in vivo. Step I leads to the synthesis of “minus” strands, step I1 to the synthesis of “plus” strands. Lodish and Zinder (52) have shown that the two steps are genetically dissociable and may therefore be catalyzed by separate enzymes. It is tempting to assume that RNA synthetase is the enzyme catalyzing step 11, while the RNA polymerase, derived from an abnormal host-virus system that makes an excess of viral “minus” strands, is enriched in the enzyme catalyzing step I. RNA replicase preparations might contain both step I and I1 enzymes. It must be stressed, however, that these assignments are only tentative a t present.

VI. Conclusions and Summary The evidence discussed in the preceding sections is compatible with the following sequence of events. Following adsorption of the phage to the male-specific pili of the host, the viral RNA strand penetrates the cell and is used as a messenger to direct the synthesis of one or possibly two viral RNA polymerases. Within a few minutes after infection the parental RNA forms part of a replicating complex; synthesis of both viral “minus” and “plus” strands ensues and a large number of additional replicating complexes are formed. Whereas about equal amounts of “plus” and “minus” strands are formed a t first, this ratio soon becomes about 10: 1 in favor of the “plus” strands. The structure of the replicating complex is not known. It may consist of one or more viral RNA polymerase molecules, as well as of “minus” and “plus” strands and possibly also ribosomes attached to the latter. After deproteinization, the so-called replicative intermediate is isolated. It consists of both double- and single-stranded RNA and contains a

REPLICATION OF PHAGE RNA

387

full-length “minus” strand. It may have the structure depicted in Fig. 4. Precursor experiments suggest that synthesis of viral RNA involves a mechanism whereby newly formed “plus” strands displace pre-existing ones from the replicating complex, while the “minus” strands are conserved (semiconservative asymmetric replication). The newly formed “plus” strands may ( a ) undergo maturation and give rise to progeny virus, ( b ) function as messenger for the formation of virus-specific proteins, or (c) participate in a new replicating complex. The control mechanisms that regulate the relative amounts of viral proteins and viral RNA species formed are not completely understood. It has been suggested that under normal circumstances only the parental RNA strands (and possibly early progeny RNA strands) direct the formation of the viral polymerase responsible for the formation of “minus” strands, and that the coat polypeptide is involved in translation control. Phage mutants in which the relevant control mechanisms have been lost provide a powerful tool for the study of this problem. The results of in vitro studies of viral RNA synthesis are compatible with a mechanism involving a “minus” strand as a template for the synthesis of progeny “plus” strands, although an alternative interpretation of the relevant data has been proposed by Haruna and Spiegelman (127).Further work will be necessary to substantiate their suggestion that “minus” strands are neither formed nor required for viral RNA replication. Careful analysis of the further purified “RNA replicase” system, which forms infective progeny RNA when primed with parental RNA, will undoubtedly yield a clear answer to this question, a8 well as to the problem whether one or two separable enzymes are involved in viral RNA synthesis, The replication of most animal and plant viruses containing singlestranded RNA has many features in common with that of the RNA phages; however i t cannot be said a t present that the mechanism is similar in all cases. I n particular it is possible that replication of viruses such as Rous sarcoma virus involves hitherto unknown features (145).

VII. Appendix: Identification and Analysis of Viral RNA Since most experiments concerned with viral RNA replication involve the identification and quantitative determination of viral “plus” or “minus” strands or of double-stranded RNA, a brief discussion of the methods used for this purpose is presented here.

A. Identification of Virus-Specific Double-Stranded RNA Double-stranded RNA is most readily identified by its pronounced resistance to RNase digestion in 0.15 M NaCl (&jJ96). A purified sam-

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CHARLES WEISSMANN AND SEVER0 OCHOA

ple of radioactive RNA is subjected to digestion by pancreatic RNase and the reaction product is then analyaed by exclusion chromatography, chromatography on methylated albumin, sedimentation through a sucrose gradient, or, more simply, by determining the amount of acid-insoluble radioactivity. Whereas the more elaborate methods allow detection of discrete cleavage of an RNA molecule, the acid precipitation method has proved to be very reliable and is far more practical for routine quantitative determinations. The criterion of virus specificity is more difficult to establish. Many workers have limited themselves to the demonstration that the RNaseresistant RNA in question is found in virus-infected but not in normal cells. In the case of encephalomyocarditis and polio, the virus specificity of the RNase-resistant RNA was established by its capacity to infect appropriate host cells (94, 109). However, infectivity could not be demonstrated for the double-stranded RNA of bacterial viruses (98) and has not been reported for plant viruses. A more generally applicable method is based on the specific annealing properties of viral doublestranded RNA. Heat denaturation of the duplex leads to separation of the two strands and, under appropriate conditions, these can be annealed to reform double-stranded RNA in high yield. If, prior to annealing, PS2-labeled viral RNA is added to the reaction mixture, then a certain fraction of the radioactive RNA is also annealed and converted to an RNase-resistant state. If the experiment is carried out in the presence of increasing amounts of Psz-labeled viral RNA, an appropriate extrapolation allows the quantitative determination of “minus” and “plus” strands present in the sample (70).The high specificity of the reaction is demonstrated in Table 11.

B. Analysis of Double-Stranded RNA (Specific Dilution Assay) This assay is used to determine the distribution of radioactivity between the “plus” and the “minus” strands of purified labeled doublestranded RNA. I n principle, radioactive double-stranded RNA is heatdenatured and reannealed in the presence of an excess of unlabeled RNA. Only if the unlabeled RNA is identical with one of the strands of the duplex will it displace its radioactive counterpart from the double strand (70,74). Since, as pointed out above, double-stranded RNA is extremely resistant to the action of ribonucleases under conditions in which singlestranded RNA is quite sensitive, the displacement of radioactive strands from the duplex can be followed by determining the RNase-resistant radioactivity before and after heating and annealing. In order to determine the distribution of radioactivity between the two strands of virus-specific double-stranded RNA, the dilution assay is carried out by heating and reannealing a series of samples of the double-stranded RNA

389

REPLICATION OF P H A G E RNA

TABLE I1 SPECIFICITY OF ANNEALING ASSAY RNase-resistant radioactivity after annealing" Labeled RNA

Ref.

(I) P3' MS2 RNA (4pg, 11,630 cpm)

c

(2) P3z MS2 RNA (0.24 pg, 9867 cpm

d

(3) PazQe RNA (0.24 pg, 11,375 cpm)

d

TMV RNA (1.88 rg, 11,600 cpm)

e

E. coli RNA (1.12 pg, 3515 cpm) (6) P3z E. coli RNA (1.3 pg, 6600 cpm)

e

(4)

(5)

P3l

P32

c

Nonlabeled RNA

cpm over controlb

% of input

MS2 RNA (400 pg) 11 0.1 Ribosomal RNA (400 pg) 11 0.1 tRNA (400 pg) 11 0.1 TMV RNA (400 pg) 16 0.14 Normal E. coli RNA (400 0 0 rg) MS2 infected E. coli RNA 1095 9.4 (400 pg) Partially purified double- 3006 26 stranded MS2 RNA (88 PgJ) Purified double-stranded 8209 83.1 MS2 RNA (8.7 pg) Purified double-stranded 0 0 QB RNA (6.2 rg) Purified double-stranded 9050 79.5 Qa RNA (6.2 rg) 10 0.1 Purified double-stranded MS2 RNA (8.68 pg) Purified double-stranded 30 0.25 MS2 RNA (2.5 pg) Normal tobacco leaf RNA 54 0.46 (0.9 mg)f TMV infected tobacco leaf 2243 19.3 RNA (1.3 mg)f TMV-infectedtobaccoleaf 0 0 RNA (0.9 mg)! Partially purified double0 0 stranded MS2 RNA (400 pg) 120°C for 3 minutes and annealed at 80°C for 30-60

5 The mixtures were heated to minutes. b The control is the RNase-resistant radioactivity of the nonheated mixture. Control: Expt. 1, 36-100 cpm; Expt. 2, 45 cpm; Expt. 3, 110 cpm; Expt. 4, 153 cpm; Expt. 5, 96 cpm; Expt. 6, 300 cpm. c C. Weissmann, P. Borst, M. A. Billeter, R. H. Burdon, and S. Ochoa, Proc. Nut2 Acud. Sci. U.S. 61, 682 (1964). d L. Colthart and C. Weissmann, unpublished. e R. H. Burdon, M. A. Billeter, C. Weissmann, R. C. Warner, S. Ochoa, and C. A. Knight, Proc. Nut1 Acud. Sci. U.S. 62, 768 (1964). f The leaf RNA was first treated with RNase and the RNase was removed by phenol extraction.

390

CHARLES WEISSMANN AND SEVER0 OCHOA

wit,li iiicreasing amounts of unlabcled “plus” strands. Thereby labeled “plus” strands are diluted out while labeled “miIius” strands reaniieal completely ; the filial ratio of displaced to nondisplaced radioactivity corresponds to the ratio of label in “plus” to that in “minus” strands. The quantitative evaluation of the dilution assay is given elsewhere (19). I n order to demonstrate that nondisplaceable radioactivity is indeed due to “minus” strands, advantage is taken of the fact that the renaturation of denatured double-stranded RNA is concentration-dependent

47.0 pg/ml MS2

no additions or

0, RNA

TIME (min)

FIQ. 14. Demonstration that the radioactive RNA remaining RNase-resistant after heating and reannealing of labeled, double-stranded viral RNA in the presence of excess unlabeled “plus” strands is due to “minus” strands. Uniformly labeled double-stranded MS2 RNA was heated and reannealed in the presence of excess unlabeled MS2 RNA. The double-stranded RNA was reisolated after RNase digestion and denatured. It waa heated without additions or in the presence of varying amounts of nonlabeled MS2, Qp (100 pglml), or TMV RNA (500 pg/ml). The rate of reannealing of the radioactive RNA is proportional to the concentration of added MS2 RNA, as would be expected if the radioactivity were due to MS2 “minus” strands (19). (95), as is that of DNA (146). Since the rate-limiting step in annealing is presumably the collison of a “plus” and a “minus” strand, the rate of annealing should depend on the concentration of both “plus” and “minus” strands. Therefore, if one denatures double-stranded RNA

391

REPLICATION O F PHAGE RNA

labeled exclusively in the “minus” strand and determines the rate of reannealing of the radioactive RNA a t different concentrations of unlabeled “plus” strands, one should find it proportional to the “plus” strand concentration. Conversely, if the rate of reannealing of a labeled RNA is found to be specifically accelerated by a certain RNA preparation, and if annealing thereby goes to completion, this may be taken as evidence that the two have a complementary base sequence. I n the experiment of Fig. 14, labeled double-stranded MS2 RNA was subjected to heating and reannealing with a large excess of unlabeled MS2 RNA; the -<

TMV

RNA

RIB.

RNA

8

MS2

RNA 0 I

THEORETICAL FOUND

60

L

5 0 ~

4 VADDED

I ADDED

RNA

8

12 (pg-‘)

2 RNA ( p g )

3

FIG.15. Analysis of uniformly labeled double-stranded MS2 RNA by the specific dilution assay. The labeled double-stranded RNA was heated and reannealed in the presence of increasing amounts of MS2 RNA. As shown by the extrapolation (inset), half the radioactive RNA remains double-stranded at infinite concentrations of cold MS2 RNA and is, therefore, due to “minus” strands ( 1 9 ) .

product was treated with RNase and reisolated by Sephadex chromatography. This double-stranded RNA, putatively labeled in the “minus” strand only, was denatured and aliquots were heated a t low concentrations with varying amounts of added unlabeled MS2 RNA for different lengths of time. As shown in the figure, the initial rate of annealing was approximately proportional to the concentration of added MS2 RNA; Qp and TMV RNA had no effect whatsoever on the rate of reannealing. Figure 15 shows the specific dilution test as applied to uniformly labeled MS2-specific double-stranded RNA. It may be seen that half of the radioactivity of the double-stranded RNA is displaced in

392

CHARLES WEISSMANN AND SEVER0 OCHOA

the dilution test. The inset shows that the reaction goes to completion a t high MS2 RNA concentrations. The analysis of asymmetrically labeled duplex RNA is presented in Fig. 7.

C. Determination of “Minus” Strands One method of determining unlabeled “minus” strands is described in Section A of the Appendix. This method has the advantage of allowing a determination of “minus” strands on a weight basis. I n order to determine in a labeled RNA sample the fraction of the radioactivity due to “minus” strands, aliquot8 of the sample are heat-denatured and reannealed with increasing amounts of unlabeled viral RNA and the ribonuclease-resistant radioactivity is then determined. An appropriate extrapolation to infinite concentration of added viral RNA allows the determination of nondisplaceable radioactivity that, after correction for ribonuclease-resistant core (i.e., fragments of RNA nondigestible even in a single-stranded state), may be equated with “minus” strands. This method is of only limited accuracy when the level of radioactivity in “minus” strands is of the order of 1% or less.

D. Determination of “Plus” Strands 1. SEDIMENTATION ANALYSISOF RADIOACTIVE RNA

Certain treatments, e.g., UV irradiation or treatment of spheroplasts with actinomycin, lead to a reduction of host-specific RNA synthesis. Following exposure to labeled RNA precursors, the “plus” strands of RNA phages can be detected as radioactive RNA sedimenting with an szo,w of about 27 S (72, 36‘). However the presence of free “minus” strands from such a fraction can not be excluded a priori. Furthermore, these methods of inhibiting host RNA synthesis may give rise to quantitative and even qualitative alterations in viral RNA synthesis (72, 86). 2. DOUBLE ISOTOPE SPECIFICDILUTION ASSAY

This assay, based on annealing techniques, permits the simultaneous determination of radioactive “plus” and “minus” strands in the presence of radioactive nonviral RNA. I n principle, radioactive viral RNA (both “plus” and “minus” strands) is converted into a double-stranded form by annealing with an excess of denatured unlabeled double-stranded viral RNA (Fig. lob). Under appropriate conditions, 8590% of the labeled viral RNA thereby becomes RNase-resistant (76). The labeled double-stranded RNA thus obtained is then analyzed by the specific dilution assay to determine the distribution of radioactivity between

393

REPLICATION OF PHAGE RNA

“plus” and “minus” strands (Fig. 1Oc). Authentic viral RNA, labeled with an isotope different from the one used for the unknown sample, is added as an internal standard in order to permit quan,titation of the results. In practice, the unknown RNA sample, usually labeled with GI4, is iiiixed with P32-labeled viral RNA and an ainount of double-stranded viral RNA in about tenfold or greater excess over the viral RNA estimated to be present. Aliquots of the mixture are heated above the T, of the double-stranded RNA and then subjected to annealing conditions, both without further additions and with addition of varying amounts of

ADDED

MS2

RNA ( p ~ )

FIG. 16. Double isotope specific dilution assay of C“-labeled RNA from E . coli infected (a) with wild-type MS2 and (b) with a hyperproducing mutant, Mu9. The unlabeled C“-labeled RNA (-C-) was mixed with P” MS2 RNA (--I, double-stranded MS2 RNA, and varying amounts of unlabeled MS2 RNA. After heating and annealing, the RNase-resistant radioactivities were determined. Inset : a plot of the fraction of C“ radioactivity converted to a double-stranded form (fa) against the corresponding P” value ( f ~ ) .The slope of the straight line gives the fraction of C’* radioactivity in “plus” strands, the intercept, the C“ radioactivity in “minus” strands. (a) slope 28%, intercept 3.570; (h) slope 26%, intercept 17.7% (19).

unlabeled viral RNA (ranging from about 0.1 to 10 times the amount of the double-stranded RNA) . The RNase-resistant P32and C’” radioactivities are determined and expressed as fractions of the corresponding total input radioactivities (fP and fc, respectively). When fc is plotted as a function of f P a straight line results, the slope of which corresponds to the C14 radioactivity present in “plus” strands, while the intercept gives the CI4 radioactivity in “minus” strands (Fig. 1Oc) (19). The

394

CHARLES WEISSMANN AND SEVER0 OCHOA

method has been tested by reconstruction experiments in which varying amounts of C1*-labeled viral “plus” strands, “minus” strands, and E. coli RNA were mixed and then analyzed. Two examples demonstrate the application of the method. Figure 16a shows the results of the double isotope specific dilution assay carried out on the RNA of MS2-infected cells labeled with C14-uracil. It is seen that the virus-specific RNA constitutes 32% of the total labeled RNA, and that the ratio of “plus” to

Pa

80

La

0

I

I

I

I

10

20

30

40

ADDED

50

60

RNA ( p g )

FIQ. 17. Base sequence homology between MS2, Qp, and f2. A mixture of P3’labeled MS2 RNA, double-stranded MS2 RNA, and varying amounts of an unlabeled RNA waa heated and annealed as indicated. Unlabeled MS2 RNA completely dilutes out the P” MS2 RNA, f2 RNA does so to a lesser extent, and Qp RNA not at all, indicating partial homology between MS2 and f2 RNA, but none between MS2 and Qs RNA (19).

“minus” strands is about 8. I n contrast (Fig. 16b), the labeled RNA of E. coli infected with an amber mutant of MS2 (Mug) shows a ratio of “plus” to “minus” strands of 1.5. 3. OTHERMETHODS Synthesis of “plus” strands can be followed by determining infectious RNA (29,SO, 69, 147-160) or by allowing infected cells to incorporate labeled RNA precursors for a limited period of time and then measuring the fraction of radioactive RNA recovered in virus particles a t the end of infection (31,60,161).

395

REPLICATION OF PHAGE RNA

E. Determination of the Base Sequence Homology between Viral RNA’s In order to determine whether or not two RNA’s have-entirely or in part-the same base sequence, one of them must have radioactive label and be in a double-stranded form. I n the case of RNA phages, double-stranded RNA labeled exclusively in the “plus” strand is readily prepared by annealing labeled viral RNA with the denatured homologous viral double-stranded RNA. With the specific dilution assay, one then determines how much of the labeled strand is displaced by the other unlabeled RNA. For complete homology, the displacement should approach completeness asymptotically with increasing additions of the unlabeled species. The assay is conveniently carried out by heating and reannealing a mixture of P3?-labeled viral RNA and homologous doublestranded RNA in the presence of increasing amounts of the second (unlabeled) RNA and determining the resulting RNase-resistant radioactivity. Figure 17 shows that there is no base sequence homology between the RNA’s of phages MS2 and Qp, whereas there is a considerable overlap between those of MS2 and f2. It may be seen that unlabeled MS2 RNA displaced all of the labeled “plus” strands from double-stranded MS2 RNA, f2 RNA displaced a considerable fraction, and Qp none a t all.

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CHARLES WEISSMANN AND SEVER0 OCHOA

96. R. Langridge, M. A. Billeter, P. Borst, R. H. Burdon, and C. Weissmann,

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106. D. Baltimore, Y. Becker, and J. E. Darnell, Science 143, 1034 (1964). 106. Y. Watanabe, K. Watanabe, N. Ishida and Y . Hinuma, J. Biochem. ( T o k y o ) 58, 322 (1965). 107. J. M. Bishop, D. F. Summers, and L. Levintow, Proc. Natl. Acad. Sci. U.S. 54, 1273 (1965). 108. F. Brown and B. Cartwright, Nature 204,855 (1964). 109. M. Pons, Virology 24, 467 (1964). 110. P.Hausen, Virology 25,523 (1965). 111. J. Sonnabend, L. Dalgarno, R. M. Friedman, and E. M. Martin, Biochem. Biophys. Res. Commun. 17, 455 (1964). 112. H. G . Mandel, R. E. F. Matthews, A. Matus, and R. K. Ralph, Biochem. Biophys. Res. Commun. 16, 604 (1964). 113. W. Shipp and R. Haselkorn, Proc. Natl. Acad. Sci. US. 52, 401 (1964). 114. R. H. Burdon, M. A. Billeter, C. Weisamann, R. C. Warner, S. Ochoa, and C. A. Knight, Proc. Natl. Acad. Sci. US. 52,768 (1964). 116. T . Sreevalsan and R. Z. Lockart, Jr., Proc. Natl. Acad. Sci. US. 55, 974 (1966). 116. R. L. Sinsheimer and M. Lawrence, J . Mol. B i d . 8, 289 (1964). 117. M. Chamberlin and P. Berg, J. Mol. Biol. 8,297 (1964). 118. P. Borst and C. Weissmann, Proc. Natl. Acad. Sci. US. 54, 982 (1965). 119. R.Rolfe, Proc. Natl. Acad. Sci. U.S.49,386 (1963). 120. B. H. Rosenberg and L. F. Cavalieri, Proc. Natl. Acad. Sci. US. 51, 826 (1964). 1Zl. C. Weiasmann, L. Simon, P. Borst, and S. Ochoa, Cold Spring Harbor Symp. Quant. Biol. 28,99 (1963). 1g2, J. T. August, L. Shapiro, and L. Eoyang, J. Mol. Biol. 11, 257 (1985). 123, L. Shapiro and J. T. August, J. Mol. B i d . 11, 272 (1965). 124. I. Haruna and S. Spiegelman, Proc. Natl. Acad. Sci. US. 54, 579 (1965). 1%. I. Haruna and S. Spiegelman, Proc. Natl. Amd. Sn'. U.S. 54, 1189 (1965). 126. I. Haruna and 5.Spiegelman, Science 150,884 (1965). 127. I. Haruna and S. Spiegelman, Proc. Natl. Acad. Sci. U S . 55, 1256 (1966). 1.87~.L. Colthart and C. Weissmann, unpublished results. 198. D. Baltimore and R. M. Franklin, Biochem. Biophys. Res. Commun. 9, 388 (1962). 129. D. Baltimore, H. J. Eggers, R. M. Franklin, and I. Tamm, Proc. Natl. Acad. Sci. U.S. 49, 843 (1963). 130. D. Baltimore, Proc. Natl. Acad. Sci. US. 51, 450 (1964). 131. P.P.K. Ho and C. P. Walters, Biochemistry 5,231 (1966). 1%. R. G.Wilson and J. P. Bader, Biochim. Biophys. Acta 103, 549 (1965). 133. L. Dalgarno, E. M. Martin, S.-L. Liu, and T. S. Work, J. Mo2. Biol. 15, 77 (1966).

REPLICATION OF PHAGE RNA

399

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Author Index Numbers in parentheses are reference numbers and indicate that an author's work is referred to although his name is not cited in the text. Numbers in italic show the page on which the complete reference is listed. A

Apgar, J., 40(20), 46(20), 56(108), 65(20), 70(20), v , 7 9 Applebaum, S., 235(115), 255 dpplequist, J., 131, 141 Aranda, L. C., 214,218 Arc&,M., 62, SO Ardrey, R., 189(5), 217 Arilrawa, S., 19(138), 37 Arion, V. J., 275(84), 277(84), 282(84), 283 (84), 284 (84), 3 17(84), 338 (84), 342(304), 346, 361 Armstrong, A., 46, 49(61), 51(61), 53(61), 78 Aronson, A. I., 44, 78 Asano, K., 275(85), 545 Askonas, B. A., 316(217), 317, 348 Asrijan, I. S., 262(35), 294(35), 295(35), 343 Astier-Manifacier, S., 385(134, 142), 399 Attardi, B., 341(301), 351 Attardi, G., 7(76), 35, 226(17, 21), 263, 287(110), 341(301), 345 Atwood, K. C., 221(13), 222(13), 223031, 224(13), 226(13), 229(13), 263 Auber, J., 173, 186 August, J. T., 305(165), 306(165), 310(165), 314(165), 347, 355(42), 377(42), 378, 379, 380(123), 381, 396, 39s Augusti-Tocco, G., 60, 80 Aurisiccio, A., 307(187), 347 Avers, C. J., 166, 186 Axelrod, V. D., 49(77), 51(77), 53(77), 79

Abhott, M. T., 306(181), 309(181), 347 ACS,G., 53(86), 79 Adanis, 8.,92(53), 136 Adams, H., 228(64), 264, 291(127), 34s Adams, J. M., 57(112), 80, 358(64), 397 Adiga, P. R., 7(78), 36, 261(11), 342 Adelberg, E. A,, 30(178), 31(186), 38 Adler, J., 85(13), 135 Admann, R., 85(9), 136 Afzelius, B. A., 163 (64), 166(83), 185 Agranoff, B. W., 205, 218 Aitchozin, M. A., 316(219), 317(227), 348, 349 Akinrimisi, E. O., 102(167), 103(167), 139 Albers, M., 305(166), 347 Alberts, B. M., 62(134), 80 Alexander, H. E., 3(31), 17(126), 21(31) 34, 37 Alexander, M., 53(90), 79 Allen, F. W., 40(15), 77 Allfrey, V. G., 225(31), 237, 263, 256, 262(21, 31), 268(60, 61), 288(114), 292(21, 31, 114, 137, 138), 293(31), 294(31), 301(60), 303 (1141, 326 ( 1141, 330(265), 332(275), 333(282, 2881, 337(283), 340(299, 300),343,344,34s, 350, 361 Amano, M., 323(235), 349 Ammann, J., 368, 370, 388(98), 398 Anagnostopoulos, C., 18(131), ST Ananieva, L. N., 269(68), 273(68), 275(68), 276(68), 277(68), 278(68), 6 279(68), 280(68), 281(68), 308(68), 317(68), 338(68), 344 Bach, M. K., 249(160), 267 Angell, C. L., 95, 137 Bacon, D. F., 12(106), 36 Angdov, E. Z., 276(89), 345 Bader, J. P., 385(132), 398 Aoki, I., 68(163), 81 Apgar, A,, 47(65), 49(65), 51(65), 53(65), Bak, I. J., 170, 171, 186 Bakks, J., 63(145), 80 59(65), 62(65), 78 40 1

402 Balbinder, E., 5(50), 35 Baldwin, R. L., 19(133), 37, 85(20), 100(199), 113(195, 196, 197, 198), 114(195, 199, 200), 115(200), 20(199, 200), 122(198), 129(198), 136, 139, 140, 390(146), 399 Ballard, P., 312(208), 345 Baltimore, D., 371, 384(139), 385(130), 398 Baltus, E., 144(12), 148, 153, 291(128), 301(128), 346 Bang, I., 98(134), 135 Bank, A., 43(39), 77 Barlow, G. H., 354(20), 36 Barnes, J. E., 12(111),36 Barnett, L., 10(92), 13(92), 21(92), 25(92), 30 Rarondes, S. H., 278(93), 316(93), 345 Barr, G. C., 332(277), 360 Barrell, B. G., 44(51), 45(51), 78 Barrnett, R. J., 166, 155 Bass, I. A., 339(297), 340(297), 361 Bass, R., 164(71), 171(71), 185 Bassel, A,, 150(33), 161(33), 154 Bausum, H. T., 4(34), 34 Bautz, E., 19(134), 21(151), 37 Bautz, E. K. F., 7(79), 19(139, 1401, 36, 37 Bauta, F. A., 19(139), 31 Bautz Freese, E., 21(151), 37 Baxill, G. W., 333(287), 360 Bayev, A. A., 44, 51(77), 53(77), 79, 91, 136 Beaudreau, G., 359(76), 382(76), 383(76), 397 Beaven, G. H., 91,136 BeCareviE, A., 247(155), 249(155), 266 Becker, Y., 239(149), 240(149), 245(149), 247(156), 266, 298(148), 316(218), 317(218, 2241, 319(148), 346, 345, 349, 371(1051, 395 Beer, M., 97, 138 Beermann, W., 206(24), 217, 222(23), 223(22), 263 Berrs, R. F., 84(1), 101(154), 107(182, 183), 126(219), 133(1), 134,139, 140 Befort, N., 63(145), 64(152), 74(152), 50, 51, 85(6), 136 Belitsina, N. V., 316(219), 317(227, 2281, 348, 349

AUTHOR INDEX

Bell, D., 52, 53(81), 72, 79 Bellamy, A., 288(111), 345 Bellamy, L. J., 95, 137 Belozersky, A. N., 269(67), 344 Belaer, N. O., 10(98), 36 Bender, M. A., 227(49), 237(49), 264, 329(262), 349 Bendich, A., 314(214a), 345 Bendigkeit, H. E., 5(44, 45), 34 Bennett, J. P., 40(5), 77 Ben-Shaul, Y., 151(35), 154 Ben-Zeev, N., 28(169), 38 Benzer, S., 7(70, 72), lO(89, 90, 91, 92, 93), 13(72, 91), 15(93), 18(91), 19(93), 21(91, 921, 23(93), 25(72, 90, 92, 93, 154), 27(72), 35, 36, 37, 39(2, 31, 40(3), 76 Berends, W., 21(148), 37 Berg, P., 40, 43(38), 53(89), 54, 71(180), 77, 79, 52,98(131), 99(142), 114(200), 115(200), 120(200), 121(131), 138, 140, 305(171, 172), 310(171), 313 (212), 314(212), 347, 348, 376(117), 398 Bergman, F. H., 71(180), 52 Bergquist, 40(25), 42(29), 43(25), 44(25), 46, 49(25), 55, 77,78, 79 Bergstrand, A., 262(17), 343 Berlowita, L., 226(35), 253 Bernfield, M., 40(7), 74(7), 77 Bernhard, W., 239(145), 240(145), 256 Bertini, F., 170(94,) 185 Beukers, R., 21(148), 37 Beurling, K., 92(53), 136 Biemann, K., 52(77b), 79 Biezunski, N., 286(108), 317(108), 346 Billen, D., 333(286), 360 Billeter, M. A., 354(1, la), 359(70, 73, 74), 361(70), 362(1), 366(95), 367(95), 368(96), 369, 370(96, 1011, 371(73, 95, 114), 372(74, 75), 373(73), 374(73), 375(73), 378(70), 379(1, 70, 75), 381(75), 383(70), 387(95), 388(70, 741, 389, 390(95), 392(75), 396, 397, 395 Biot, J. B., 91, 1% Birnboim, C., 341(303), 361 Birnstiel, M. L., 220(8), 221, 222(18), 223, 225, 226(12, 18), 228(66), 229(71), 236, 237(127, 1281, 239(146),

AUTHOR INDEX

403

S.,62(1401,SO, 225(27), 259 Boyve, R. P., 6(64), 31(64), 35 Brachet, J., 144(12), 148, 183, 262(15), 304(15), 343 Bradley, D. E., 354(15), 396 Bradley, D. F., 131(245, 245a), 141 Brayg, J. K., 130(235), 140 Brahms, J., 62(128), 80, 94(77, 78, 79, 80, 81, 82, 83, 84), lOl(74, 78, 80), 103(78, 83), 104(74), 106(81, 84), 126(83), 133(82), 137 136 Blake, R.D.,91(41),107(41,186), 108(41), Brahms, K., 106(176), 139 Brandes, D., 170,185 1109(41),115(41), 128(41), 136, 130 Brattgbrd, S.-O., 193(17,191,196(19), 217 Blakeslee, A. F., 28(170), SS Braunitzer, G., 2(19), 25(19), 31(19), 34 Bloch, D. P., 332(279), 350 Brawerman, G., 144(15), 149, 158(15), Blumenfeld, L.A., 97,138 16'3, 202(22), 206(25), 217, 269(69), Boedtker, H., 62(133), SO, 99(146), 275(69), 278, 286(108), 316(94, 221), 116(146), 138, 369, 354(7), 395 317(108), 338,344, 345, 348, 358(66), Boer, A., 308(196), 348 397 Bogdanov, A. A., 314(214), 348 Bogorad, L.,150(32), 151(32), 158(49), Brcckenridge, B., 341(302),351 Bremer, H., 305(170), 306(180), 310(180), 164(32), l S 4 314(170), 315(170), 347 Bolle, A,, 26(159), 37, 356(48), 396 Rrenner, S., 10(92), 13(92), 21(92), Bollum, F. J., 85(14), 120(208), 135, 140 25(92), 26(159, 160, 161), 36, 37, Bolron, M., 234(112), 258 249(162), 257, 356(47, 48), 396 Bolton, E. T., 202(23), 217, 260(8), Breslow, E., 106(181), 110(181),113(181), 277(91), 279(96), 283(91), 308(96), 139 316(91), 328,342, 345 Breugnon, M. M., 385(139), 399 Bond, H. E., 235(114), 255 Bridges, B. A., 21(146),37 Bond, V. P., 227(48),254 Briggs, A,, 329(258),349 Bonner, D. M., 26(158), ST, 309,348 Brimacombe, R.,40(7), 74(7), 77 Bonner, J., 225(30, 331, 245(33), 253, Brinkley, B. R., 239(147), 256 262(22), 264(52), 276(go), 291(52), Brinton, C . C., Jr., 354(22), 396 299(90, 1561, 309, 310, 326(52), Britten, R.J., 231(94), 255 330(204), 331,332,333,334(289), 337, Brodsky, V.J., 293(141), 346 339(90), 343,344,3..$5,847,348,350 Brookes, P., 19(135,136),37 Bonner, W. D., Jr., 144(10), 150(10), Brown, D. D., 220,227(6,40,41,44,46), 162(10), 1SS 228(6), 229(6), 231(6), 251(170), 253, Bopp-Hassenkamp, G., 290(117), 346 254, 257, 275, 280(81), 309(81, 1971, Borek, E., 228(66), 254, 327(253), 349 316(197), 323(81, 1971,340(298), 345, Boriosova, 0. F., 62(130), 90 34S, 351 Brown, D. M., 85(9), 135 Borkowska, I., 232(105), 255 371(108), 398 Borst, P.,354(1), 358(118), 359(69, 70, Brown, F., 74), 360(69), 361(70), 362(1), 368(96), Brown, G . L., 40,42(4), 47(67), 59(67), 60,63(141),77, 78, 80, 92(52), 136 369, 370(96), 372(74, 75), 377(118), 324(243), 349 378(70, 121), 379(1, 70, 75, 1181, Brown, R., 1(2), 33 381(75), 383(70), 388(70, 741, 389, Browne, T., ' Brownlee, G. G., 44(51), 45(51), 7 s 392(75), 305, 397, 398 Bruscov, V. I., 342(306),351 Bourgeois, A,, 314(213), 348 253, 254, 256, 264(52), 291(52, 126, Rourgcois,

129, 298(126), 301, 302(129, 162), 303(126), 324,326(52, 126,161,162), 327(249, 251), 344, 346, 347, 349 Bishop, D. H. L., 354(15), 395 Bishop, J. M., 371(107),398 Bishop, J. O., 252(175), 257 Bjurstam, K.,193(20),917 Bladen, H.A., 310(205), 320(205), 34s 109(41), 114(41), 115(41), 128(41),

404

AUTHOR INDEX

Cartledge, J. L., 28(170), 3s Bruskov, V. I., 295(145), S.$G Cartwright, B., 371 (1081, 398 Brgson, V., 2(9), 10(105), 25(9), 33, 36 Budowsky, E. I., 56(104), 68(104), 19 Caspersson, T., 262(14), 304(14), 343 Cassassa, E. F., 86(25a), 135 Buetow, D. E., 170(94), 185 Buettner-Januscbh, J., 2(18), 25(18), 34 Caughlin, C. A., 31(186), 38 Burdick, C. J., 333(283), 337(283), 350 Cavalieri, L. F., 121(213), 140, 377(120), 398 Burdon, R. H., 354(1), 359(70, 74), 361(70), 362(1), 368(96), 369, 370(96), Cernd, J., 66(160), 67(160), 68, 75(160), 81 371(114), 372(74, 75), 378(70), 379(1, 70, 75), 383(70), 388(70, 741, 389, Chalkley, G. R., 333(281), 350 Chamberlin, M. J., 98(131), 99(142), 392(75), 395, 397, 398 lOO(205, 2591, 114(200), 115(200,255), Bursztyn, H., 28(165), 38 116(205), 118(205), 120(200, 2051, Busch, H., 219, 228(64), 229(821, 231(96), 121(131, 2051, 122(205,214),124(205), 232, 233(96), 234(96), 253, 254, 255, 128(255), 138, 140, 141, 305(171), 275(86), 290, 291(127), 299(152), 310(171), 347, 376(117), 398 301(152, 158, 159), 323(239), 325(86, 158), 331(274), 333(274, 2801, 345, Chambers, R. W., 69, 81 Chambon, P., 312(209, 2101, 348 346, 347, 349, 350 Champe, S. P., 25(154), 37 Butler, G. C., 52(79), 79 Butler, J. A. V., 307(216), 315(216), Chan, S. I., 97(115), IS8 Chandler, B. L., 290(115), 329(115), 346 332(277), 348, 350 Chandra, P., 69, 81 Byrd, C., 85(15), 135 Chandrasekhar, B. K., 385(138), 399 Byrne, R., 310(205), 320(205), 34s Chang, A. Y., 8(82), 35 Byvoet, P., 219(5), 253 Changeux, J.-P., 328(255), 349 C Chapeville, F., 39, 76 Chargaff, E., 3(30), 34, 40(23), 42(31), Caillet, J., 132(248), 1.41 46, 52(31), 77, 262(23), 263(23), Cairns, J., 144(18), l S 4 290(125), 343, 34G Callan, H. G., 329(263, 264), 350 Chein Ching Chen, G., 30(178), 38 Calvet, F., 262(18), 343 Cheng, T., 56(109), 79 Calvori, C., 62(139), 80 Chentsov, J. S., 262(29, 341, 290(29, 341, Camahan, J., 354(22), 390 300(29, 34), 343 Cameron, I. L., 164(67), 170(67), 185 Chevremont, M., 163, 164(61), 185 Canellakis, E. S., 53(88), 79 Chiang, K. S., 159, 161(50), 184 Cantoni, G. Id., 41(57), 43(47, 49), 45, Chiasson, L. P., 21(147), 37 46(57), 47, 48(54), 49(47), 59(120), Chidester, J. L., 385(138), 399 78, 80, 88(33), 13G Chipchase, M. I. H., 220(8), 221(8), Cantor, C. R., 94(70), 13300, 2511, 136, 236 ( 126, 128), 237 ( 128), 253, 256, 264(52), 291(52, 126, 1291, 298(126), 141 301, 302(129, 161), 303(126), 324, Capecchi, M. R., 57(112), SO, 358(64, 326(52, 126, 161), 327(249), 344, 346, 68), 397 Carbon, J. A., 42(34), 63, 74(1491, 77, 347, 349 Chun, E. H. L., 147,184 81 Carlton, B. C., 7(73), 10(73), 21(73), Clark, A. J., 354(8), 395 Clark, A. M., 251(169), 257 26(73), 27(73), 35 Clark, B. F. C., 358(65), 397 Carreni, N., 308(196), 348 Carrier, W. L., 6(63), 21(148a), 31(63), Clark, J. M., Jr., 8(82), 35, 71(176), 81 Clausen, J., 182, 186 35, 37

405

AUTHOR INDEX

Claverie, B., 132(248), 141 Cleveland, L. R., 174(103), 176(103), 186 Clever, U., 225(29), 253 Cline, M. J., 371(102), 385,398 Cochram, G. W., 385(138, 143), 399 Cohn, M., 225(27), 663 Cohn, W. E., 40, 77, 127(225), 140 Colon, E., 209(34), ,918 Colthart, L., 354(19), 383(127a), 389, 390(19), 391(19), 393(19), 394(19), 396, 398 Comb, D. G., 229(73, 74), 234(74), 254, 324 (243), 349 Connelly, C. M., 46(60, 631, 47(63), 78 Contois, D. E., 6(56), 35 Contreras, G., 355(38), 396 Coons, A. H., 193,217 Cooper, P. D., 355(39), 396 Cooper, S., 355(31, 33, 42, 45), 357(56), 359(31), 377(42), 379(56), 394(31), 396, 397 Cooper, W. D., 144(14), 183 Corneo, G., 144(9), 162(9), 163, 168, 183 Cornuet, P., 385(134, 142), 399 Counts, W. B., 235(114), 255 Cowan, C. A., 157(46), 184 Coward, S. J., 334(291), 350 Cox, E. C., 12(107), 36, 235(117), 255 Cox, R. A,, 91(45), 110(189), 122(189), 136, 139 Gamer, F., 59, SO Crampton, C. F., 290(125), 346 Crane, H . C., 130(228), 140 Crawford, E. M., 354(23), 396 Crawford, I. P., 18(131), 37 Crick, F. H. C., 2(23), 10(23), 18(23), 31(23), 34, 39, 76, 95, 101(157), ,122(157,215), 130, 137, 139, 140 Crocker, T. T., 262(20), 343 Crothers, D. M., 130(239), 131, 141 Cubiles, R., 43(41), 78 Curtis, R., 12(116), 36 D Dais, D., 40(6), 77 D’Alesandro, P. A., 163(63), 185 Dalgarno, I,., 371(111), 39,s D’Amato, F., 28(174), 38

Damle, V., 131, 141 Daneholt, B., 193(19), 196(19), 217 Daniel, V., 54(94), 79 Darmstadt, R. A,, 6(58), 35 Darnell, J. E., 229(76, 79), 230(79, 88), 233(79, 88, 1101, 234, 235(116), 239(88, 1101, 240(88, 110), 654, 665, 264(43), 267(43), 269 (43), 271(43), 275(43), 285(103), 286( 103), 287(103), 316(218), 317(218, 224), 318(248), 319(229), 326(229, 248), 341(303), 343, 345, 348, 349, 351, 371(1051, 398 Das, N. K., 228(51), 254 Davern, C. I., 249(161, 1621, 257, 354(13), 356(49), 364(91), 378(91), 380(91), 396, 396, 397 Davie, E. W., 71(181), 82 Davies, D. R., 91(39), 98(152), 101(157), 106(39, 178), 113(198), 116(202, 203), 122(157, 198, 216), 124(202), 129(198), 136, 138, 139, 140 Davies, J., 63(143), 76(143), 80 Davis, F. F., 40, 77 Davis, J. E., 355(29), 357(59), 360, 394(59), 396, 397 Davison, P. F., 222(16), 253 Dawson, G. W. P., 12(115), 36 De, D. N., 290(119), 346 Dehov, S. S., 288(113), 345 De Giovanni, R., 5(48), 6(53, 54), 10(100, 102), 12(100), 31(100, 102, 1851, 32(102), 34,35,36,38 De Giovanni-Donnelly, R., 2 (41, 10(4), 13(4), 17(4), 18(4), 33 Dekker, C. A., 40(14), 41(14), 77 DeKloet, S. R., 268(60), 301(60), 344 Delbriick, M., 2(22), 31(22), 34 Delius, H., 368(98), 370(98), 388(98), 398 Delong, J. C., 234(112), 655 De Margerie-Hottinguer, H., 167(79), 1S6 Demerec, M., 2(3), 3(29), 4(33, 35), 7(71), 8(71), lO(3, 94, 95), 11(3), 12(35), 17(129, 1301, 33, 34, 36, 36, 37

De Moss, J. A., 26(158), 37 Dc Robcrtis, E. D. P., 174(100), 1SO De Salle, L., 162(60), 186

406 Desjardins, R., 231(96), 232(96), 233(96), 234(96), 266 Dessev, G. N., 276(88), 288(112), 346 Deutscher, M., 72, 82 de Vries, H., 3(27), 34 deWaard, A,, 12(110), 36 Dhaluval, A. S., 385(138), 399 Diacumakoa, E. G., 167, 186 Dibben, H. E., 85(10), 135 Dickson, R. C., 162(158), 164(58), 169(58), 185 Dicckmann, M., 53(89), 54(98), 71(180), 79, 82, 313(212), 314(212), 348 DiGirolamo, A., 286(107), 317(107), 3-45 Dimarzio, E. A,, 130(234), 140 Dingman, C. W., 278(93), 316(93), 334 (290), 546, 360 Disbrey, C., 144(16), 172(16), 176,184 Djerassi, C., 93, 136 Dohzhansky, Th., 1(1), 7(1), 25(1)~ 29(1), 3.9 Doctor, B. P., 46(63), 47(63), 78 Doerfler, W., 305(166), 347 Doi, R. H., 354(20), 355(35), 360(81), 396, 397 Dolapchiev, L., 251 (172), 267 Doly, J., 312(209,210), 348 Dondon, J., 63(142), 76(142), SO, 84(3), 127(223), 136, 140 Dondon, L., 127(225), 140 Donnellan, J. E., 21(149), 37 Doty, P., 62(133, 1341, 63(147), 75(147), 80, 81, 91(45), 98, 99(146), 101(151), 110(151), 115(129), 116(146), 121(128), 131(244a), 136, 138, 141, 147(27), 149(26), 184, 331, 337(271), 550 Dounce, A. L., 264, 288(50), 291(50), 343 Douthit, H. A., 146,183 Douzou, P., 134(252, 2531, 141 Drake, J. W., 28(168a), 58 Drude, P., 91,136 Drynin, J. F., 314(214), 348 Duhnau, D., 2(14), 25(14), 34, 222(15), 263 Dubuy, H. G., 162(56), 182(112), 186 Diitting, D., 48(68), 49, 50, 51(68), 52(77b), 53(68), 55(77a), 78, 79 Duhn, W., 130(230), 1.40

AUTHOR INDEX

Dunn, D. B., 5(47)9 77 Dure, L., 229(84) !266 Dutton, G., 62(136), 80

41(30)j 42(30)9

Eason, R., 371 (102), 385(102), 398 Ebel, J. P., 63(145), 64(152), 74(152), 80: 81, 85(6), 94(80), 101(80) 136, 137 ~ h ~ ~W.~T,, ~ 179, l d186, Ebstein, R., 235(115), 266 ~ d ~M., l157,~184 ~ ~ , Edgell, M . H., 354(26), 596' E ~ ~ A.,~ 215(50), ~ G $18~ , EdstrGm, J.-E., 191(14), 192(14), 193(14, 16), 206(16, 24), 215, 217, 218, 223(24), 227(45), 228(60), 253, 264, 292, 299, 300, 346, 347 Egami, F., 42, 43(43), 78 Eggers, H. J., 384(129), 398 Egyhazi, E., 190(9, lo), 194, 195, 196(38), 199(9, lo), 200(10), 201(9), 202(9), 203, 205(9), 207(9), 209(9, lo), 211(9, lo), 217, 218 Eichner, D., 215(50), 218 Eidlic, L., 43(37), 77 Eigner, E. A., 74,82 Eisenberg, H., 86(25a), 136 Elsenstadt, J. M., 144(15), 149, 158115)t 18.9, 202(22), 217, 278(94), 286(108), 316(94), 317(108), 346, 358(66), 397 Eisenstnrk, A., 31(188), 38 Ekholm, R., 196, 218 Elias, H. G., 85(24), 136 Eliasson, N. A., 262(17), 343 Ellem, K. A. O., 251(166, leg), 267, 269(70), 275(70), %9(70), 344 Elliott, A. M., 170, 171,186 Ellis, D. B., 355(40), 359(78), 396, 397 Elson, D., 262(23), 263(23), 343 Emerson, T. R., 133(251a), 141 Engelhardt, D. I,., 57(116), 80, 355(30), 357(52a, 53, 54, 58), 358(53, 54), 394 (30), 396, 397 Enger, M. D., 354(6, 171,596 Englander, J . J., 59,80 Englander, S. W., 59, 80 Eoyang, L., 379(122), 598 Ephrati-Elizur, E., 17(123),18(123), J R

407

AUTHOR INDEX

Ephrussi, R., 165(76), 167, 15’6 Ephrussi-Taylor, H., 4(37), 9(85), 3 4 , S G Epstein, H.T., 149(28), 150(31), 151(35), 155,156,157(44,46),184 Erickson, R. I,., 359(71,72),360,361(71), 364, 365(72), 366(72), 371(85), 372(72), 392(72), 397 Ernster, L., 166(83), 185 Errera, M., 228(57, 58), 232(106), 237(131,135), 254, 256, 323(237),349 Estable, C., 290(124), 346 Evans, E. A., 267(57), 344 Everett, G. A., 7(74), 35, 47(65), 49(65), 51(65), 53(65), 59(65), 62(65), 78 Eyring, H., 113(194), 139 Eyzaguire, J. P., 71(176), 81 F

Fabre, J. H., 188(4), 217 Falkow, S.,2(13),25(13), 34 Falzone, J. $., 268(64), 344 Fancher, H., 313(212), 314(212), 348 Fasman, G. D., 62(127), 80, 103(169), 104(169), 139 Faulkner, R., 340(299), 351 Fawcett, D. W., 172(86c), 174, 178(104), 186

Feinendgen, L.E., 227(48), 254 Feix, G., 359(77), 364(91), 378(91), 380(91,382(77), 384(77), 385(77), 397 Feldmann, H.,52(77a, 77b), 79 Felsenfeld, G., 59,80, 88(31, 33), 91(40, 42), 101(42), 103(42, 1591, 106(39, 40), 107(40, 42), 109(42), 112(42), 113(159), 114(42), 115(42), 128(42), 133(159), 135, 136, 139 Fenster, J. H., 237(136),256 Fenwick, M. L., 359(71, 72), 360(71, 851, 361(71), 364, 366(72), 371(85), 372, 392(72), 397 Ficq, A., 227(43), 237(135), 254, 256 Firket, M., 164(65), 186 Fischberg, M.,221(12), 225(12), 226(12),

Flciwnrr, E., 228(66),254, 327(253), 349 Flessel, C. P., 101(148), 138 Fletcher, M.J., 171,IS6 Fletcher, W.E,85(10),136 Flexnrr, J. I3., 205(48),21s Flexner, 1,. B., 205,21s Flickinger, R. A., 334(291), 350 Flory, P.J., 85(16), 135 Fouace, J., 394(148), 399 Fowler, A,, 332(278), 350 Fox, C. F., 105(173), 139, 306(173), 313(211), 339(211,296),347, 348, 350 Fox, J. J., 91,136 Fox, M.S., 5(42),34 Fradkin, L.I., 160(52),IS4 Franklin, N.C., 7(75), 35 Franklin, R.M., 306(175), 308(175), 34?, 360(71, 85), 361(71), 364(71, 72), 366(72), 371(85), 372(72), 384(129), 392(72), 397, 398 Frazicr, J., 95(99, 1001,107(99), 109(100, 137

Freese, E.,2(8, 12, 211, 9(88), 10(91), 13(91, 120, 1211, 16(121), 18(91), 19(134), 20(21, 881, 21(91, 120, 121, 151),25(8,121,28(8, 88, 164),31(21), 33, 34, 36, 37, 130(233), 140 Frenster, J. H., 225(32), 237(136), 253, 256, 262(31), 292(31, 137), 293, 294(31), 330,333(284,293), 343, 346, 360

Fresco, J. R., 62(133, 134), 80, 84(1), 86(25), 88(23, 30), 89(28), 91(41), 92(53),94(65), 98(128), 99(138, 146), lOl(148, 151, 152, 156), 103(152), 104(170), 105(174), 107(41, 186), 108(28, 411, 109(41), 110(151), 114(41), 115(28, 41), 116(146), 121(128), 128(41), 133(1), 135, 136, 138, 139

Friedman, R. M., 371(111), 398 Friedman, S. M., 63(143), 76(143), SO Frolova, 1,. Y., 62(130), 66(159), 68, 75(159), 80, 81 253 Fromageot, P., 130(227), 140 Fischer, S.,355(38), 396 Frontali, L.,62(139), 80 Fitts, D. D., 93(61), 136 Fuchs, E., 305(166), 315(167), 347 Fixman, M., 130(229), 140 Fujimoto, M., 160(53), 184 Flaks, J. G., 235(117), 255 Flamm, W. G., 235(114), 236, 237(127, Fujimura. R..236(123). 256 Fujimura, Y.; 57(110),’58(110), 70 1281,255, 256

AUTHOR INDEX

408 Fukada, T., 251(165, 168), 267, 275(79, 80), 280(79, 801, 317(79, So), 346 Fukutome, H., 68(163, 164, 165), 74(164), 81

Fuller, W., 47(67), 59(67), 78 Furth, J. J., 53(90), 79, 305(1@), 307(185), 312(185, 2071, 3-47, 348

G Gaines, K., 306(180), 310(180), 34'7 Galibert, F., 234(112), 266 Gall, J. G., 229(80), 230(80), 233(80), 239(80), 240(80), 266, 292(130)3 299(130), 300(130), 329(264), 346, 360 Garen, A., 355(44), 356(46), 396 Garen, S., 355(44), 396 Garnjobst, L., 167(78), 1% Cause, G. G., 322(233), 338(233), 349 Gavrilova, L. P., 236(121), 266, 269(67), 344

Gagarjan, K. G., 275(83), 280(83), 285(83), 346 Geiduschek, E. P., 95, 97, 121(212), 137, 140, 305 (169), 307( 187), 308 (NO), 347, 348, 360, 387(84), 397 Gelboin, H. U., 41(57), 46(57), '78 Gellert, M., 98, 138 Gemski, P., Jr., 3!54(22),396 Genchev, D., 251(172), 267 Georgiev, G. P., 228(61), 229(83), 230(83), 232, 264, 264 259(2, 61, 261(12), 262(27, 28, 29, 32, 34, 351, 263(46, 74), 264(40, 42, 46, 47, 481, 265(39), 266(74), 267(40, 42, 59), 268(46, 47, 48), 269(68), 271(46, 73, 74, 75, 76), 273(42, 68, 76, 78), 274(78), 275(68, 73, 74, 75, 76, 841, 276(68), 277(46, 68, 841, 278(68), 279(68), 280(68, 75), 281(12, 68,102), 282(84), 283(84), 284(84), BS(76, log), 287(78), 288(28,76, 781, 290(27, 28, 29,34), 292(32, 131, 133), 293(32), 294(35, 133), 295(35, 145), 296(146), 298(146), 299(6), 300(29, 34, 741, 301(74), 304(2, 6), 308-08, 73, 78), 309(47, 48), 310(206), 311(206), 313(206), 317(68, 84, 226), 319(75, 230), 320(6, 206, 230), 321(76, 1461,

323(48, 68, 73, 74, 75, 78, 1337 325(76), 326(74, 75), 327(133), 331(269), 335(269), 336(269), 337 (269), 338(48, 68, 84, 131), 342(304, 306), 346, 343, 344, 346, 346, 348, 349, 361 Gerassimova, H., 28(172), 38 Gesteland, R. F., 354(7,23), 369, 396, 396 Giannoni, G., 116(206), 117, 122(206), 140 Gibbons, J. R., 174(101), 186 Gibbs, J. H., 130(234), 131(242, 243), 140, 141 Gibor, A,, 144(11), 147, 148, 149, l5O(24), 153(42), 157, 158(45), 159, 183, 184 Gierer, A., 281(101), 346 Gilbert, W., 43(38), 63(143), 76(143), 77, 80, 231(93), 266 Gilden, R. V., 333(289), 334(289), 350 Gilmour, R. S., 342(307, 308), 361 Ginoza, W., 354 (26), 396 Girard, M., 230(88), 233(88, 1101, 239(88, 110), 240(88, 110), 266, 318(248), 319(229), 326(229, 2481, 341 (303), 349, 361 Gladstone, L., 305(164), 347 Glasky, A. J., 385(140), 399 Glasstone, S., 113(194), 139 Glitz, D. G., 40(14), 41(14), 77 Godman, G. C., 332(279), 360 Godson, G. N., 360(82), 397 Gold, L., 202(22), 217, 278(94), 316(94), 346 Goldberg, A. L., 3(28a), 8(28a), 34 Goldberg, I. H., 20(141), 37, 306(174), 310(203), 331(203), 347, 348 Goldmann, M., 305(168), 347 Goldstein, A., 10(99), 12(99), 31(99), 32(99), 36 Goldstein, J., 40(5), 77 Goldstein, L., 262(20), 308( 193), 343, 348 Gomatos, P. J., 370(99), 398 Gomirato, G., 210(35), 212(35), 218 Gonatas, N. K., 166(84), 166 Gonzalez, P., 228(54), 264 Goodgal, S. H., 18(132), 37 Goodman, M. H., 306(178), 347 Gorini, L., 25(155), 37, 63(143), 76(143), SO, 357(61), 397 Gorlenko, Z. M., 339(297), 340(297), 361

AUTHOR INDEX

Goto, K., 160(53), I S 4 Could, J. L., 363 (86), 364(86), 365(86), 366(86), 392(86), 397 Grabnar, M., 25(157), 37 Grachev, M. A., 56,68,?9 Grado, C., 355(38), 396 Graffe, M., 84(5), 135 Graham, A. F., 229(78), 230(78), 233(78), 271(71), 240(78), 254, 259(1), 275(71), 286, 304(1), 342, 344, 354(9), 355(9), 363(89), 371, 394(151), 395, 397, 399 Grampp, W., 193(16), 206(16), 217 299(153), 300(153), 347 Granboulan, N., 239(141), 256, 309(198), 34s Granboulan, P., 309(198), 348 Grandchaml), S., 167, 165 Granick, S.,146(21), 151(21), 153(42), 157, 158(45), 184 Gratzer, W. B., 91(45), 136 Graziosi, F., 307(187), 347 Green, D. M., 17(125), 37 Green, D. W., 84(1), 133(1), 134 Green, M., 144(17), 184 Green, M. H., 231(95), 255, 308(191), 348 Green, P. B., 152, 184 Greenberg, H., 229(85), 230(85), 231(85), 233(85), 234(85), 235(85), 240(85), 251(171), 255, 267 Greenberg, J., 30( 177), 31 (181, 182, 183, 184), 36 Greenspan, C. M , 307(188), 3.48 Greer, S., 2(26), 5(38, 48), 10(1m), 12(100), 18(26), 19(26), 21(26), 28(26), 31(100), 34, 36 Griboff, G., 3(32), 5(46), 31(46), 34 Griffin, B. E., 126(220), 140 Grimstone, A. V., 174(101), 186 Groeniger, E., 85(14), 135 Grogan, D., 231(96), 232(96), 233(96), 234(96), 265 Gros, F., 7(76), 35, 62(140), SO, 231(93), 955, 300(155), 314(213), 341(302), 347, 348, 351 Grosjean, M , 94, 137 Grossman, L., 69(170), 61, 85(8, g), 103(167), 104(169), 135, 139 Grossman, 13. I., 144(9), 162(9), 163(9), 168(9), 183

Grunherg-Manago, M., 63(142), 76(142, 80, 84(2, 3, 5 ) , 122(2), 127(223, 224, 225), 135, 140 Guerrier, C., 151(37), 159(37), 184 Guest, J. R., 26(162), 37 Guinier, A., 97, 138 Gulland, J. M., 85(10), 135 Gumport, R. I., 313(211), 339(211), 348 Gunther, J. K., 81 Gupta, S. L., 300(155), 347 Gurdon, J. B., 220, 227(6, 46), 228(6), 229(6), 231(6), 253, 254, 275, 280(81), 309(81), 323(81), 340(298), 345,351 Guschlbauer, W., 62(135), 80, 88(28, 34, 35), 89(28), 90(35, 36), 92, 107(28), 115(28), 135, 136 Gussin, G. N., 358(68), 397 Gutlman, B., 7(77), 35 Guttes, E., 164(70), 165 Guttes, S., 164(70), 185 Gvozdev, V. A,, 285(104), 286(105), 293(142), 322(104, 105), 338(104), 345, 346

H Hadjiolov, A. A,, 249(164), 251, 257 Hadjivassiliou, A., 316(221), 338, 346 Haemmerling, J., 148(25), 184 Hagemann, R., 152, 157(41), 184 Hagopian, H., 46(61), 49(61), 51(61), 53(61), 78 Hahn, E., 17(126), 37 Hall, B. D., 220(7), 253 Hall, C. E., 97, 138, 317(224), 349 Hall, J. B., 7(78), 35, 261(11), 342 Hall, R. H., 42, 52(28, 32), 77 Halvorson, H. O., 146, 183 Hamherger, A,, 208(28, 291, 218 Hamilton, M. G., 237(137), 256, 292(138), 546 Hammarsten, E., 262( 171, 343 Hanawalt, P. C., 6(65), 35, 157(47), 184 Harbers, E., 306 (176), 347 Hardman, J. K., 26(163), 37 Harel, J., 308(196), 548 Harel, L., 308(196), 348 Harkness, D. R., 70(175), 71(175), 81 Harris, H., 230(89), 255, 259(3), 267(58), 285(3), 329(261), 340(261), 342, 344, 349

410 Hartman, P. E., 25(157), 37 Hartman, Z., 25(157), 37 Hartmann, K. A., 102(258), 141 Haruna, I., 355(43), 359(76), 377(43), 381, '382(76, 125, 126, 127), 383(76, 125, 127), 387, 39Y, 397, 398 Haschemeyer, A. E. V., 111(191), 139 Haselkorn, R., 62(133), 80, 99(146), 1105(173), 116(146, 2041, 121(210), 135, 159, 140, 150(30), 159(30), 162(60), 184, 186, 371(113), 398 Hashizume, H., 68(164), 74(164), 81 Haslam, W. J., 126(220), 140 Hausen, P., 371(110), 376(110), 398 Hayashi, H., 71(179), 72(179, 73(184), 75, 82 Hayashi, M., 7(75), 35, 231(95), 266 307(186, 189), 308, 347, 348 Hayashi, M. N., 307(186, 1891, 308(189), 34s Hayashi, Y., 52(78), 49(72), 69(171), 70(174), 75(171), 78, 79, 81 Hayatsu, H., 40(8), 64(153, 154), 65(154), 66(156), 75(154), 77, 81 Haynes, R. H., 6(65), 36 Haywood, A. M., 355(36, 371, 359(37), 392(36), 396 Hecht, L., 43(41), 78 Hecht, L. I., 53(87), 79 Heldenmuth, L. H., 2(4), 10(4), 13(4), 17(4), 18(4), 29(176), 30(176), 33, 38 Helene, C., 134(252, 253), 141 Helge, H., 164(71), 169(90), 171(71), 186 Helinski, D. R., 4(36), 8(36), 21(36), 26(36), 27(36), 34 Hell, A., 228(57), 237(131), 264, 866, 323(237), 349 Helmkamp, G. K., 97(115), 101(153), 103, 104(166), 138, 139 Henley, D. D., 86(25), 136 Hennix, U., 162(55), 184 Henshaw, E. G., 239(150), 240(150), 247(150), 866, 286(107), 298(149), 317(107), 319(149), 321, 3-45, 346 Heppel, L. A., 54, 79, 131(244, 245, 245a), 141 Herbert, E., 53(88), 54, 55, 56(102, 1031, 79

AUTHOR INDEX

Herriott, R. M., 9(87), 20(87), 28(87), 36 Hess, A., 208(26), 217 Hiatt, H. H., 229(81), 231(93), 233(81), 252 ( 176), 2&5, 866, 267, 264 (44), 267 (44), 269 (441, 271 (44), 275 (44), 286(106, 107), 298(149), 317(107), 319(149), 321(149), 322(106), 325 (246), 346, 346, 349 Hidley, J., 331(268), 360 Higashi, K., 231(96), 232(96), 233(96), 234 (96), 266, 275 (861, 325 (86), 346 Highton, P. J., 97( 1221,138 Higuchi, M., 160,184 Higucki, S., 60(124), 80 Hill, R. F., 30, 38 Hill, R. L., 2(18), 25(18), 34 Hill, T., 130, I40 Hilmoe, R. J., 70(175), 71(175), 81 Hindley, J., 332, 360 Hinds, H. A., 6(58), 36 Hinuma, Y., 371(106), 395 Hirsch, C., 252(176), 267 Hirschbein, L., 105(174), 139 Hirschman, S. Z., 88(31), 136 Hnilica, L. S., 225(34), 253, 323(240), 324(240), 333(280, 286), 349, 360 Ho, P. P. K., 385(131), 398 Hoagland, M. B., 40, 77, 316(217), 317, 348 Hoffman, E. J., 177,186 Hoffmann-Berling, H., 354(11, 121, 355 (32), 368,396,396,398 Hoffman-Ostenhof, O., 28(174), 38 Hofschnaider, P. H., 305(167), 315(167), 347, 354(10), 355(34), 368(98), 370(98), 388(98), 396, 396, 398 Hohlhage, H., 66(158), 81 Holcomb, D. N., 84(1), 94(69), 101(69), 103(69), 133(69, I), 136, 136 Holiday, E. R. 91,136 Holland, I. B., 359(76), 382(76), 383(76), 397 Holland, J. J., 233(109), 235(109), 266 Holler, B., 231(100), 235(100), 266 Holley, R. W., 7(74), 36, 39(3), 40(3, 201, 45(56), 46,47,49, 51(65), 53(65), 56(108), 59, 62(65), 65, 70, 76, 77, 78, 79

411

AUTHOR INDEX

Holm, R. E., 280(97), 346 Holmes, A., 31(188), 33 Holper, J. C., 385(140), 399 Holtzer, A,, 95, 97,137 Holtzman, E., 229(85), 230(85), 231 (85), 233(85), 234(85), 235 (85), 240 (85), 265 Holzwarth, G., 94, 137 3' Homma, M., 363(89), 371, % Honig, G. R., 322(232), 338(232), 349 Hopkins, J . M., 179(110), 186 Hopkins, J. W.,220(9), 221(9), 226(9), 247(157), 263,257, 262(301, 293(30), 298(150) 319(150), 324(244), 326(150), 343,346, 349 Horiuchi, K., 356(51), 359(50), 394(50) 396 Horn, E. E., 9(87), 20(87), 28(87), 36' Horn, V., 12(107), 36 Horowitz, N. H., 144(20), 184 Horton, E., 371(103), 385,398 Hosokawa, K., 236(123), 166 Hotta, Y., 150(33), 161, 184, 262(25, 261, 292(25, 261, 294(25), 301(25, 26), 343 Howard, B. D.9 13(119), 21(119)9 36 Howard-Flanders, P., 6(64), 31(64), 35 Howell, R. R., 249(163), 267 Hoyer, B. H., 202(23), $17, 277, 283, 316(91), 329(260), 346, 3-49 IIsu, W.-T., 306(173), 347 Huang, P., 226(17, 211, 263 Huang, R. C., 225(30, 331, 245(33), $53, 276(90), 287(110), 299(90), 309(199), 310, 330(204), 331(270, fl2) 332(270, 272), 333(270, 2891, 334(289), 3377 339(90), 346, 347, 348, 360 Huang S. L., 121(213),1/0 Hudson, W. R., 385(141), 399 Humm, D. G., 170,186 Humm, J . H., 170, 186 Humphrey, G. B., 299(151), 346 Hung, L., 42(34), 77 Huppert, J., 385(139), 394(148), 399 ~ ~ ~ lR.b B,, ~ 225(34), ~ t , 228(65), 253, 264, 323(240), 324(240), 349 Hurwitr, J., 53(90), 19, 305(165, 1681, 306(165, 17g), 310(165), 314(165, 179), 332(278), 347, 960 IIwang, M. I . H., 341(301), 361

Hyde, B. B., 236(126), 239(146), $56, 302(161, 162), 326(161, 162), 347 Hyden, H., 190(9, 10, 11), 191(15), 193(17, 20), 194, 195, 196, 197(40, 42, 43), 199(9, lo, I],), 200(10), 201 (9), 202 (9), 203, 204(27), 205(9), 207(9), 208(27, 28, 291, 209(9, 10, 27), 210(35), 211 (9, lo), 212(35), 214, 216(40), 217,218

I Ibn]l, J., 98(133), 138 Ibuki, F., 40(16), 17 Iitaka, Y., 60(124), 80 Ikeda, Y., 364(90), 397 I & ~ D., ~ , 166(83), 185 Imahori, K., 68(164), 74(164), 81 Imamoto, F., 315(215), 348 Imbenotte, J., 308(196), 3-48 Infante, A. A., 280(98), 318(98), 3-46 Ingle, J., 280(97), 345 Ingram, V. M., 25(156), 37, 40(24), 41(53), 46(61), 49(61), 51(61), 52, 78, 79 Inman, R. B., 99(145), 100(145, 199, 207), 102(145), 103(145), 106(145), 113(195, 196), 114(195, 1991, 118(145, 207), 119(207), 120(199), 123(145, 1207), 124(145), 129, 138, 139, 140, 390(146), 399 Ipprn, K. A,, 354(24), $96 Ishida, N., 371(106), 398 Ishida, T., 55, 56, 57(101), 70 Ishikura, H., 48(69), 49(69), 78 D. A., 236(121), 266 IX~anov, Iwamura, T., 150, 158, 160, 184 Izawa, M., 144(1I), 147,183,333(282), $60

J Jacob, F , 7(76), 36, 225(26), 253, 328 (2551, 349 J:lcob, H., 166(85)~168(85), Jacob, J., 229(71), ,754, 323(241), 327(2519 252), 349 Jacob, M.7 195, 318, 354(20), 9' ' Jacob, S. T.,231(96), 232(96)~233(96)1 234(96), 265 ,J:lcot,son, A . B., 158(49), 184

412

AUTHOR INDEX

Katolr, A,, 161(54), 184 Katz, S., 229(73), 264, 324(243), 349 Kaudewitz, F., 394(149), 399 Kaufmann, B. P., 28(168), 38, 290(119), 346 Kauzmann, W., 91, 95, 133(249), 136, 141 Kawade, T., 251(168), 287 Kawade, P., 49(75), 62(137), 68(163, 165), 74(164), ?8, 80, 81, 251(165), 257, 275(79, 80), 280(79, 80), 317(79, go), 346 Kawata, M., 56, 79 Kedrowsky, B. V., 262(16), 343 Kcller, E. B., 62, SO Kelley, D. E., 228(68), 229(68), 231(68), 243(151), 245(151, 152), 247(151), 249(151), 254, 256, 271(77), 273(77), 344 Kelly, R. B., 360(83), 361(83), 363, 364(86), 365, 366(86), 392(86), 397 Kelner, A., 6(61), 36 Kempf, J., 229(87), 232(87), 255 Kendrew, J. C., 95, 137 Kern, M., 71(182), 76, S2 K Kerr, S. E., 91, 136 Key, J. L., 280(97), 345 Kabat, S., 226(17, 211, 253, 287(110), 345 Khessin, R. B., 328(256), 339(297), Kadoya, M., 310(201), 348 340(297), 349 Kagi, J., 62(126), 80 Khorana, H. B., 40, 52(80, 821, 77, 79, Kaerner, H. C., 368,398 85(15), 136 Kaesberg, P., 354(6, 171, 396 Khrushthev, N. G., 323(242), 349 Kaffiani, K. A., 275(82), 280(82), 346 Kidson, C., 267 (551, 344 Kaiser, A. D., 19(133), 37 Kieras, F. J., 150(30), 159(30), 184 Kaji, H., 48(62), 78 Kikuchi, G., 160(53), I S 4 Kalf, G. F., 169(89), 186 Kikugawa, K., 66(156), 81 Kallenbach, N. R., 84(1), 133(1), 136 Kilgore, W. W., 31 (182), 38 Kanazir, D., 232(106), 266 Kim, Y. T., 385(136, 141), 399 Kaplan, N. O., 2(20), 25(20), 31(20), 34 Kimura, I., 60(123), 60(123), SO Kaplan, S., 26(161), 37, 356(47), 396 Kimura, K., 262(36, 37), 264(36), 265(36, Karam, J. D., 12(112a), 36 37), 267(36, 37, 59a), 343, 344 Karasaki, S., 239(142), 266 King, T. J., 329(258), 349 Karasek, M., 385 (137), 399 Kirby, K. S., 259(4), 262, 265(41, 541, Karav, W., 48(68), 49(68), 50(68), 51(68), 267(55), 349, 343, 344 53(68), 78 Kirchner, C. E., 10(104), 12(104), 36 Kassanis, B., 8(80), 36 Kirk, C., 394(151), 399 Kasten, F. H., 228(55), 264 Kirk, J. T. O., 144(13), 148, 183 Kataja, E., 25(155), 37, 357(61), 397 Kirkwood, J. G., 93(61), 136 Katchalsky, A., 214, 218 Kislev, N., 150, 151(32), 164(32), 184 Kates, J. R., 159(50), 161(50), 18.4 Kisselev, L. L., 62, 66(159), 66(159), 68, 75(159), SO, 81 Kato, K., 323(241), 327(250), 349

Jacobson, K. B., 268(63), 344 Jardetaky, C. D., 97,138 Jardetzky, O., 97,138 Jaskunas, S. R., 133(251), l4l Jayarama, J., 144(7), 162(7), 169(7), 183 Jesensky, C., 43(39), 77 Jinks, J. L., 146, 174(2), 182(2), 183 Job, P., 90, 136 Johnson, E. A,, 91,136 Johnson, H. G., 249(160), 267 Johnston, F. B., 264(51), 291(51), 344 Joklik, W. K., 239(149), 240(149), 245(149), 247(156), 256, 298(148), 319(148), 346 Jones, D. S., 40(8), 42(34), 77 Jones, K. W., 6(57), 21(57), 36, 327(250), 349 Jordan, D. O., 85(10), 136 Josse, J., 99(143), 106(175), 120(175),138, 139 Judes, D., 195(37), 218 *Tulien,J., 234(111), 255

413

AUTHOR INDEX

Kit, S., 262(33), 267(56), 292(33), 343, 344 Kitzinger, C., 113(193), 139 Kleinfeld, R. G., 233(108), 255 Kleinschmidt, A. K., 370(101), 39s Klempercr, E., 101(152), 103(152), 198 Kline, B., 88(32), 136 Klinger, P. D., 205(47), 218 Klotz, L. C., 62(132), SO, SS(28, 30), 89(28), 91(41), 107(28, 411, 108(41), 109(41), 114(41), 115(28, 411, 135, 136 Knight, C. A., 43, 48, 78, 371(114), 359, 39s

Knollc, P., 394(149),, 399 Knorre D. G.,91(51), 13G Korh, A. L., 20,SY Koelle, G. B., 191(13), 217 Koenig, E., 191(13), 193(17), 217 Kohn, K. W., 6(66), 35 Komzolova, S. G.,91(51), 136 Konigsberg, W., 354(3), 357(52a), 395,

Kuhn, T. S., 189(7), 217 Kukhanova, M. K., 62(130), SO Kumar, S., 13(118), 21(118), 24(118), 36 Kurland, C. A., 231 (931, 256 Kuwano, M., 69, 75(171), S1 Kuykov, V., 69 (173a), 81 Kyogoku, Y., 60(124), 80, 95(88, go), 137

1 Imakey, M. D., 182(112), 186' Lacks, S., 13(122), 18(122), 36 Lafontaine, J. G., 228(52), 254 Lagerkiiist, U., 54, Y9 Laidler, K. G.,113(194), 1% Lajtha, A., 197, 218 Lamborg, M. R., 62(126), SO, 94(73), 13Y Landauer, T. K., 209(30), 218 Landridge, R., 103(168), 122(168), 137, 139

Lane, B. G., 40(15), 52(79), YY, 79 Lang, N.,278(95, 95a), 316(95), 345 Langan, T., 338,550 396 Lange, P. W., 204(27), 208(27), 209(27), Konrad, N. W., 305(170), 306(180, 182), 214, 2lY 309(182), 310(180), 314(170), Langridge, R., 368,369, 370(96, 99), 398 315(170), 34Y Lanni, F., 2(11), 25(11), 33 Koppelman, R., 267 (57), 344 Kornberg, A., 5(51, 52), 8(51), 20(51, Larsen, C. J., 234(112), 265 52), 35, 85(12, 13), 99(143), 106(175), Larsson, S., 21s Latham, A. B., 31(187), 38 120(175), 136, 138, 139 Kornberg, R. D., 313(212), 314(212), 348 Latham, H., 229(79), 230(79), 233(79, 110), 234, 235(116), 239(110), Korner, A,, 317(223), 34s 240(110), 254, 865, 285(103), Kozlov, J. V., 310(206), 311(206), 286(103), 287(103), 318(248), 313(206), 320(206), 348 326(248), 345, 349 Krakow, J. S., 306(181), 309(181), Laursen, R. A., 84(4), 129(4), 136 339(295), 34Y, 350 Lawley, P. D., 19(135, 136), 3Y Kratky, O.,95, 97, 197 Lawrence, M., 98( 130), 121(130), 1 3 , Krechevsky, I., 189(6), 217 376(116), 399 Kreig, D. R., 19(137), 37 Krichevskaya, A. A,, 295(145), 296(146), Leader, P., 40(7), 74(7), '77 298(146), 321(146), 342(306), 346, 351 Leblond, C. P., 323(235), 349 Leboy, P., 235(117), 255 Krone, W., 63(148), 81 Ledbetter, M. C.,173,186 Krsmanovic, V., 232(106), 955 Lee, H. H., 5(41), 34 Krug, R., 53 (901, Y9 Krutilina, A. I., 49(77), 51(77), 53(77), Lee, M. H.,385(138), 399 Lee, S., 40(11), YY, 92, I36 56(104), 68(104), YO Kuhitschek, H. E, 5(44, 45), 21(14.5), Legrand, M., 94, 13Y Lehman, I. R., 12(110),36 34, 37 Lehmnnn, I., 85(13), 135 Kuff, E. L., 162(59), 170, 1S5

AUTHOR INDEX

414 Leidy, G., 3(31), 17(126), 21(31), 34,37 Leng, M., 103(159), 113(159), 133(159),

Litt, M., 62(133), SO, 99(146), 116(146), 138

Littau, V. C., 237(137), 866, 292(138), 330(267), 333(283), 337, 346, 360 Lengyel, P., 74 (71), 78 Littauer, U. Z., 43(38), 54(94), 77, 79 Leonard, N. J., 84(4), 127(4), 136 Le Pecq, J. B., 107(185), 124(205a), 139 Littna, E., 227(41, 44), 264, 309(197), 316(197), 323(197), 34s 140 Lerman, M. I., 229(83), 230(83), 233(83), Liu, S.-L., 371(103), 385(103), 398 266, 263(75), 266(74), 269(68), Lockart, R. Z., Jr., 371(115), 398 271(73, 74, 75, 76), 273(68, 76), Locke, J., 188, 217 275(68, 73, 74, 75, 76), 276(68), Lodish, H. F., 356(50, 51, 521, 357(56, 57), 359, 362(57, 85a), 364(57), 365, 277(68), 278(68), 279(68), 280(68, 376,378(52,57), 379(56), 386,394601, 75), 281(68, 102), 286(76, 1091, 396, 397 288(76), 300(74), 301(74, 157), 308(68), 317(68), 319(75, 230), Loeb, J. N., 249 (163), 267 320(230), 321(76), 323(73, 74, 751, Loeh, T., 354(4), 355(4), 396 325(76), 326(74, 751, 338(68), 344, Loftfield, R. B., 74, 82 Logan, R., 262(24), 292(24), 294(24), 343 346, 347,349 Levin, J. G., 310(205), 320(205), 348 Loh, P., 307(185), 312(185, 2071, 347, 34s Levine, J., 17(128), 37 Lohrmrtnn, R., 40(8), 77 Levinthal, C., 130(228), 140 Longuet-Higgins, H. C., 130(232), 140 Levintow, L., 317(107), 398 Longworth, J. W., 97(119a), 138 Li, L., 49(77), 51(77), 53(77), 79 Loper, J. C., 25(157), 37 Li, T., 62(126), 80 Lord, Fr. A., 228(52), 264 Liau, M. C., 225(34), 228(65), 263, 254, Loring, H. S.,144(14), 183 291 (127), 323(240), 324(240), 346,349 Luborsky, S. W., 41(57), 46(57), 78 Libonati, M., 354(1a, 19), 358(79), Luck, D. J. L., 144(8), 162, 164, 165, 359(73, 791, 371(73), 372031, 166, 168, 169(8), 170(71a), 171(74), 373(73), 374(73), 375(73), 383(79), 183. 156 390(19), 391(19), 393(19), 394(19), Luco, J. V., 214, 218 199

396, 396, 397

Luft, R., 166(83), 186 Lieberman, I., 323 (238), 349 Luria, S. E., 7(75), 36, 308(192), 348 Lifson, S., 131(247), 141 Luazati, V., 95, 97(112), 101(158), Lin, F. H., 166(81), 186 103(158), 137, 138, 139 Lindahl, T., 86(25), 136 Lindblow, C., 62(127), SO, 103(169), Lwoff, A., 177, 1S6 Lyman, H., 157, 1S4 104(169), 139 Lyon, M. F., 226(37), 264 Lindegren, C. C., 144(6, 19),1S3 Lyttleton, J. W., 158(48),184 Lindegren, G., 144(6, 19), 183, 184 Lindenmayer, G., 228(64), 254 M Lindsley, D. L., 221(13), 222(13), 223(13), 224(13), 226(13), 229(13), 263 Lipmann, F., 39(2), 40(5), 53(50, 8 6 ) , McCarthy, B. J., 202(23), 817, 231 (94), 256, 260(8), 277(91), 279(96), 283(91), 70, 77, 78, 79 308(96), 316(91), 328, 329(260), 342, Lipsett, M. N., 42(33), 77, 98(132, 1371, 346, 949 101(147), 104(171, 172), 120(147), Macleod, H., 144(20), 184 131(245, 245a), 138, 139 l 4 l McClintock, B., 10(97), 36 Iipshitz, R., 290(125), 346 McConkey, E. H., 220(9), 221(9), 226(9), Lipshitz-Wiesner, R. 40(23), 46, 77 231 (92), 232 (92), 247 ( 1571, 251 (92), Iitman, R. M., 5(49), 9(85, 86), 34,36

415

AUTHOR INDEX

257, 298(150), 319(150), 324(2441,326( 150), 346, 349 McCully, K. S., 43(47, 491, 47, 49(47), 78 McDonald, C. C., 97,103(117), 121(117), 263,

138

McElroy, W.D., 6(62), 35 McEwen, B. S., 193(20), 197(42, 43), 198(42, 431, 217, 818, 333(288), 350 McGregor, D.,190(11), 197(12), 217 Mach, B.,268(66a), 344 McLaughlin, C. S.,52,79 McMullen, D., 92(52), 136 McVittie, A., 179(110), 186 Madison, J. T., 7(74), 35, 42, 47(65), 49(65), 51(65), 53(65), 59(65), 62(65), 77, 78 Magee, W. S., 131,141 Maggio, R., 237(139), 256, 264(53), 291(53), 293(53), 298(53), 301(53), 344 Maheshwari, N., 309(199), 347 Mahler, H.R.,62(136), 80, 88(32), 136, 144(7), 162(7), 169(7), 183 Maitra, V., 306(179), 314(179), 347 Maling, B. D., 4(36), 8(36), 21(36), 26(36), 27(36), 34 Maling, M., 115(255), 128(255), 141 Malkoff, D. B.,170(94), 185 Mandel, H.G., 371(112), 376, 398 Mandel, M., 2(13), 25(13), 30(178), 34, 38, 150(31), 184 Mandel, P.,195(37), 218, 229(87), 232(87, 105), 256, 312(309), 348 Mandell, J. D., 30(177, 1791, 31(181), 38

Mantieva, V. L., 262, 263(46), 264(10, 46,47,48), 265,266(74), 267(40, 591, '268(46,47, 48), 271(73, 74), 275(73, 74), 277(46), 300(74), 301(74), 306(183), 308(73), 309(47, 48, 183), 312(183), 323(48, 73, 74), 326(74), 338(48), ~ 3 0 4 343,344,347,551 ) ~ Manton, I., 165(73), 185 Marcaud, I,., 341(302),351 Marciello, R., 62,80 Marcker, K. A., 57, 59(111), 79, 80, 358(65),397 Margoliash, E., 2(15, 16), 25(15, 16), 26(15, 16),27(16), 34

Marinozzi, V., 239(143), 256 Markov, G. G., 276(88), 288(112), 345 Marmur, J., 2(13, 141, 17(128), 25(13, 14), 34, 37, 98(126, 127, 128, 129), 115(129), 121(128), 138, 144(9), 149(26, 27), 162(9), 163(9), 168(9), 185,184,222(15), 253,307(188),34s Marquisee, M., 45(56), 47(65), 49(65), 51(65), 53(65), 59(65), 62(65), 7 8 Marshak, A., 262(18), 343 Marshall, R.,63(144), 80 Martin, E. M., 371(103, 1111, 385(103), 398

Marullo, N., 3(32), 34 Marvin, D. A., 354(11, 12), 395 Mason, S.F., 94(62), 136 Masson, F.,101(58), 103(58), 139 Massoulik, J., 88(28), 89, 90, 91(41), 94(85), 99(138), 101( 1551, 103(161), 105(155), 107(28, 41, 184, 1871, 108(41, 187), 109(41, 187), llO(187, 188), 111(187), 112(187), 114(41, 85), 115(28, 41, 1871, 120(187), 123(187, 188), 125(85, 217), 127(85), 128(41, 85), 129(85). 133(16), 135, 136, 137, 138, 139, 140 Mathius, A., 101(158), 103(158), 139 Matsukage, A., 68(163), 81 Matsuo, K., 95(90), 137 Matsushita, S.,40(16), 77 Matsuzaki, K., 40(17), 41(17), 44, 77 Mattaei, J. H., 74(70), 78, 278(92), 345 Mattern, C. F. T., 162(56), 180' Matthews, R.E. F., 42(29), 77, 371(112), 376(112), 385,398, 399 Mat,us, A., 371(112), 376(112), 398 Maurer, H.R., 333(281), 360 Mayer, A . M., 28(175), 38 Mayr, E., 2(7), 33 MazC, R., 355(32), 396 Mrhler, A. H., 77 Mehrotra, B. D., 62(136), SO, 88(32), 136 Melchers, F., 48(681, 49(68), 50(68), 51(68), 53(68), rs Merker, H. J., 169(90), 185 Mcrrill, S. H.,40(20), 46(20), 47(65), 49(65), 51(65), 53(25), 59(65), 62(65), 70(20), 77, rs Meselson, M. N.,98, 138, 162, 185, 236(125), ~ ~ ( 1 6i1 62, ), 256, 257

416

AUTHOR INDEX

Michaelidis, P., 195(37), I18 Michelson, A, M., 84(1, 3,4), 85(7, Il), 88(29, 34), 89, 93, 94(83, 84, 851, 98(136), 99(29, 139, 140), 101(150), 103(83, 1641, 105(29, 139), 106(84, 140), 114(85), 120(29), 123(29), 124(221), 125(7, 85,217,217a), 126(7, 83, 221), 127(7, 85, 223, 224, 225), 128(29,85,2261,129(4,85), 131(246), 133(1, 161,248a,251a), 134(252, 253, 254), 134, 135, 136, 137, 138, 139, 140, 141

Miyaeaki, M., 49, 50(73), 51(74), 56(106), 78, 79 Miaoguchi, T., 69(167), 81 Moffitt, W., 93,136 Molnar, J., 295(145), 342(306), 346, 361 Mommaerts, W.F. H. M., 62(128), SO, 94(79, 80,81), 101(80), 106(81), 137 Monier, R., 234(111), 255 Monny, C., 84(4), 99(140), 101(1M), 124(221), 126(221), 106( 140), 127(226), 128(226), 129(4), 130,135, 138, 140

Monod, J. J., 225(26), 253, 328(255), 349 Montagnier, L.,288(111), 346, 367(94), 368(94), 371(94), 388(94), 397 Montgomery, T.H., 219(1), 253 256 Mikulska-Macheta, A., 385(143), 399 Moohr, J. W., 121(212), 140, 360(84), 387(84),397 Miles, H.T.,95, 107(98, 99), 109(100), Moon, M. W., 85(15), 136 111, 116(98,202), 124(202), 137, 140 Moore, B.C., 177,I S 6 Miller, C., 20,37 Moore, B. W., 190(11), 197, 217 Miller, 0. L., Jr., 164(68), 168(68), 1% Mills, D., 359(76), 382(76), 383(76), 397 Moore, C., 144(9), 162(9), 163(9), Mills, S.E.,26(158), 37 168(9), 183 Mil'man, L.S., 267(59),344 Morell, P., 2(14), 25(14), 34 Minckler, S.,144(6, 19), 183, 184 Morgan, C . H., 98(133), 138 Mintz, B., 227(391,264 Morikawa, N.,315(215),34s Mirsky, A. E., 225(31), 237(136, 137)~ Morimoto, T., 268(62), 344 253, 256, 262(21, 31), 268(60, 61)~Morisawa, S.,42(31), 52(31), 77 288(114), 292(21, ii4, 132, 137, Morita, T.,68(165), 81 1381,293(31, 294(31), 301(60)9 Maser, c.,334(291), 351) 303(114), 326(114), 330(265, 26717 Maser, H., 5(43), 34 332(275), 333(282, 283, 288), Mothe', K'' 151(38)'184 338(292), 340(299), 343, 344, 346, 350, Moundrianakis, E.N., 97(121, 123,124),

Michiko, D., 394(150), $99 Micou, J., 262(20), 303(193), 343, 34s Midgley, J. E. M., 40(19), 77, 235(113),

351

Mirzabekov, A. D., 49(77), 51(77), 53(77), 56(104), 68(104), 79 Mitra, S., 354(6), 395 Mitsui, H., 310(201), 348 Mitsui, Y., 60(124), SO Miura, K.I., 40(17, 18), 41(17), 42(18), 43(44, 48), 44(48), 45, 49(72, 74), 51(74), 52(78, 83), 55, 56, 57(101, 110), 58(110), W(123, 124), 64(154, i55), 65(48, 154, m), ~(155, 156, i61), 67, 68(155), 69(171), 70(83, 174), 71(179), 72(179, 184), 73(184), 75(154, 155,171,186),77, 78, 79, 80, 181, 82, 371(104), 398 Miyagi, M., 334(291), 350 Miyake, T., 10(103), SG

Mounolou, 138

185 J. c,, 166, Mueller, G. C., 229(77), 230(91), 232, 233(91), 235(91), 254, 265, 271(72), 275(72), 301,302(160), 325,344,347 306(176)7347 Muench* K . v 43(38)3 '?

w.! H.

J.j

3)761(82

3s

M1lnkresa Ii. D . ~ 1677 i86 Munro, A. J., 317(223), 348 Munson, R. J., 21(146), 37 Muramatsu, M., 231(96), %2(96), 233(96), 234(96), 255, 275(86), 301(152), 291(127), 299(152), 325(86), 345,346, 347 Muriay, K.,331(272), 332(272), 350

AUTHOR INDEX

Murray, M. J., 28(170), SS Muto, A., 64, 65(154, 155), 66(155, 156), 70(174), 75(154), 81, 371(104), $98

Novick, A., 5(39, 40), 7(77), 34, 36, 328(257), 349 Nowinski, W. W., 174(100), 186 NVZU, K., 355(43), 377(43), 381(43), 396

N:igaj, H., IS5 ?;agai, S., 166(77), 165 Nakada, D., 238(140), 252(173), 256, 257, 326(247), 849 Nakamoto, T., 305(169), 347 Namiki, O., 160(53), 184 Nanney, D. I,., 178(109), 186 Naono, S., 7(76), 35 Naora, H., 237( 138), 238 (138), 256 Nardone, It. M., 228(54), 254 Nass, M. M. K., 162(55), 163,184,155 Nass, S., 162, 163(64), 184, 1,s Nathans, D., 261(10), 342, 357(62, 63), 397 Nawashin, M. S., 28(172), 38 Neidhardt, F. C., 43(37), 77 Nemer, M., 227(42), 229(42), 231 (42), 247(159), 254, 257, 280(98), 316 (220), 318(98), 545, 346 NeSkoeiE, B. A., 228(56), 254 Ner, H. C., 54, 79 Neubert, D., 164(7l), 169(90), 171, 185 New, K., 17(130), 37 Newell, G. F., 131(243), 141 Nichols, C., 28(173), 38 Nickel, L., 18(132), 37 Nihei, T., 45, 48(54), 59, 78 h'iklowitz, M'., 290(120), 346 Nirenberg, M. W., 40, 63(144), 74(7, 701, 77, 78, SO, 278(92), 310(205), 320, 345, 348 Rishimura, S., 40(8), 59, 71, 75(177), 77, 80, 81 Noll, F., 317(225), 349 Xoll, H., 231(99), 255, 322(231), 349 Nomura, M., 236(123), 249(162), 966, 267 Nomura, N., 7(70), 36 Nonoyama, M., 364(90), 397 Nordberg, E., 262(17), 345 Notani, G. W., 354(3), 357(52a, 621, 396, 306, 397 Novelli, C . D., 59, 71, 75(177), 79, 81

Ochoa, S., 40, 74(9, 71), 77, 78, 354(1, 2), 355(41), 359(70, 74), 361(70), 362(1), 371(114), 372(74, 75), 377(41), 378(41, 70, 1211, 379(1, 70, 75), 381(75), 383(70), 388(70, 741, 38,9, 392(75), 396, 396, 397, 398 Ofengand, J., 69, 70, 71(180), 81, 82 Ogawa, K., 166,185 Ogur, M., 144(6, 19), 183, 184 Ohta, N., 17(129), 37 Ohtaka, Y., 355(43), 358(67), 377(43), 381 (43), 396, 397 1% Ohtsuka, E., 40(8), 77, Oishl, M., 222(14), 226(14), 229(75), 263, 254 Okada, Y ., 69 (167, 168), 81 Okagaki, H., 262(36), 264(36), 265(36), 267(36), 543 Okamoto, T., 49, 52(84). 62(138), 78, 79, $0, 247(154), 256, 298(147), 346 Okamura, N., 229 (82), 266, 275(86), 301 (158), 325(86, 1581, 346, 547 O'Neal, C., 40(7), 74(7), 77 Oplatka, A., 214, 218 Orgel, L. E., 225(27), 2'53 Osawa, S., 231(101), 25.5, 262(25, 261, 288(114), 292(25, 26, 114, 132), 294(25), 301(25, 261, 303(114), 310(201), 316(222), 326(114), 343, 346, 348 Otaka, E., 310(201), 316(222), 346 Oura, H., 231(99), 255, 317(225), 349 Ovcrhy, L. It., 354(20), 390

Painter, R. B., 227(48), 264 Palade, G. E., 264(53), 291 (53), 293(53), 298(53), 301 (53), 344 Pnlmieri, G., 166(831, 185 Paoletti, C., 107(185), 124(205a), 139,140 Paranchych, W., 354(9, 271, 355(9, 401, 359(78), 394(147), 595,396, 397,399

AUTHOR INDEX

418 Pardee, A. B., 5(49), 34 Parnas, H., 341(301), 361 Parsons, J. A., 162(58), 164(58, 66), 168(58), 185 Patterson, D. L., 100(205), 116(205), 118(205), 120(205), 121(205), 122(205, 214), 124(205), 140 Paul, A. V., 12(110), 36 Paul, J., 342(307, 308), 361 Pauling, L., 2(17), 25071, 26(17), 34 Pavlovec, A., 236(120), 256 Pease, D. C., 173, 174(98), 186 Pedersen, K. O., 85(17), 135 Pelling, C., 222(23), 263 Pelling, G., 308(194), 34s Penman, S., 97(117), 103(117), 121(117), 138, 229(85), 230(85, 88), 231(85), 232(102, 1031, 233(85, 88, 110), 234, 235(85), 239(88, 110), 240, 251(171), 266, 267, 316, 317(218, 224), 318(248), 319(229), 326(229, 2481, 342,348, 349, 351 Penniston, J. T., 63, 75,81 Penswick, J., 97(118), 138 Penswick, J. R., 47(65), 49(65), 51(65), 53(65), 59(65), 62(65), '78 Perkins, H. J., 160, 184 Perry, R. P., 221(10), 228(10, 57, 58, 59, 62, 63,67, 68), 229(63, 67,68), 230(63, go), 231(63, 67, 681, 233(10, 671, 235(67), 237(131), 239(10), 240(10, go), 241, 242, 243(90, 151), 245(10, 151, 152), 247(151), 249, 263, 264, 266, 256, 264(45), 271(77), 273(77) 286(45), 288, 308(45), 319, 323(236, 2371, 326(228a), 343, 344, 346, 349 Pestka, S., 63(144), 80 Peterkofsky, A., 43(39,40), 77 Peterlin, A., 85(22, 23), 86(22), 97, 136, 138

Peterman, M. L., 236(119, 120), 256 PetroviE, J., 247(155), 249(155), 266 Petrovib, S., 247(155), 249(155), 266 Pfeiffer, D., 357(59), 394(59), 397 Philips, J. H., 85(9), 136 Phillips, W. D., 97(117, 118), 103(117), 121(117), 138 Philpot, J. S. L., 333(287), 360 Pierce, B. L. S., 12(113), 36

Pierrr, J. G., 41(53), YS Rgon, A., 190(11), 199(11), 217 Plato, 188, 217 Platova, T. P., 293(143), 346 Plaut, W., 150(34), 151(34), 184 Porhon, F., 85(7), 88(29), 89, 94(85), 99(29), 105(29), 114(85), 120(29), 123(29), 125(7, 85), 126(7), 127(7, 85, 225), 128(29, 85), 129(85), 135, 137, 140 Poddar, R. K., 13(118), 21(118), 24(118), 36 Pogo, A. O., 237(137), 266, 292(138), 346 Pogo, B. G. T., 237(137), 266, 292(138), 346 Polanyi, M., 188, 189, 217 Poljakoff-Mayber, A., 28(175), 38 Pollard, C. J., 146,184 Pons, M., 371(109), 388(109), 398 Poole, F., 80 Porod, G., 95, 137 Porter, K. R., 172(96a), 173, 186 Pratt, A. W., 91, 92, 136 Preiss, J., 53(89), 71(180), 79, 82 Prescott, D. M., 227(49, SO), 237(49), 254, 259(5), 262(19), 304(5), 329(262), 342, 343, 349 Preuss, A., 305(167), 315(167), 347 Price, T. D., 6(58), 35 Prokof'ev, M. A., 314(214), 348 Prokoschkin, B. D., 275(83), 280(83), 285(83), 345 Prosorov, A. A., 339(297), 340(297), 351 Pullman, A., 2(24), 34 Pullman, B., 2(24), 34, 132(248), l4l R Rabinowitz, M., 20(141), 37, 162(60), 186, 306(174), 322(232), 338(232), 347,349 Radding, C. M., 85(12, 131, 99(143), 106(175), 120(175), 135, 138, 139 Rahn, R. O., 97,138 Rajewsky, B. N., 28(166), 38 Rake, A. V., 229(78), 230(78), 233(78), 240(78), 264, 259(1), 271(71), 275(71), 286, 304(1), 342, 344 Ralph, R. K., 52, 79, 267(55), 344, 371(112), 376(112), 385(135), 398,399

AUTHOR INDEX

Ramenskaya, G. P., 267(59), 344 Rammler, D. H., 53(50), 75, 79 Ramuz, M., 312(209, 210), 345 Rancourt, M. W., 166(81), 155 Randall, J., 144(16), 172(16), 176, 179, 154, 156 Ravin, A. W., 2(10), 25(10), 33 Rawitscher, M. A,, 103(163), 111(163), 112(163), 131, 139 Ray, D. S., 157, l S 4 Ray, U '. J., Jr., 39(2), 76 Reddy, T. K. R., 21(150), 28(150, 1651, 37, 35 Reese, C. B., 126(220), 140 Reich, E., 121(211), 130(227), l 4 V , 144(8), 162, 166, 168, 169(8), 170(71a), 153, 186, 306(174, 175), 308(175), 347 Reichard, P., 262(17), 343 Reichmann, M. E., 8(81, 82), 35 Reid, J. C., 91, 92, 136 Reilly, E., 7(79), 35 Reiner, B., 6(53), 20(143), 35, 37 Rcther, B., 64(152), 74(152), 51, 85(6), 135 Revel, M., 239(150), 240(150), 247(150), 256, 2%(106), 298(149), 319(149), 321(149), 322(106), 325(246), 345, 346, 349 Rho, J. H., 262(22), 291(126), 298(126), 303(126), 326(126), 343, 34s Rhoades, M., 10(96), 36 Rice, S. E., 130(235), 140 Rich, A., 84(1), 91(39, 40), 99(144), 101(1571, 102(258), 103(168), 106(39, 40, 178, 179, 1801, 107(40), 116(201, 203, 2061, 117, 122(157, 168, 206), 133(1), 134, 136, 13S, 139, 140, 141, 147(23), l S 4 , 306(178), 317(224), 347, 349, 370(100), 398 Rich, K., 6(53, 54), 31(185), 35, 35 Richards, E. G., 62(132), SO, 88(30), ~ ( 5 3 1 ,101(48), 135, 136, 13s Richards, H. H., 41(57), 46(57), 48(69), 49(69), 78 Riley, F., 162(56), 155 Riley, M., 115(255), 128(255), 141 Ris, H., 150(34), 151(34), 154, 290(115, 116), 329(115), 338(292), 346, 350 Risebrough, R. W., 231(93), 256

419 Ritossa, F. M., 221, 222(13), 223, 224, 226(13), 229(13), 253, 324, 349 Ro, T. S., 228(64), 231(96), 232(96), 233(96), 234(96), 254, 655, 323(239), 349 Roberts, D. W. A., 160, 154 Roberts, R. B., 209(31), 218, 231(94), 238( 139a), 255, 256 Robertson, J . M., 46,78 Robinson, W. S., 306(173), 347 Rolfe, R., 377(119), 39s Rollins, E., 334(291), 3550 Roman, H., 167(79), 155 Rosenberg, B. H., 377(120), 39s Rosenkrana, H. S., 314(214a), 345 Ross, P. D., 103(162, 163), 111(162, 163, 192), 112(163), 131(163), 139 Rosset, R., 234(111), 255 Rothsrhild, J. A., 245( 153), 256 Rottman, F., 40(7), 74(7), 77 Rouvicke, J., 7(76), 35 Roaencwajg, R., 385(139), 399 Rubin, A. I,., 198, ,215 Rudner, R., 5(50), 35 Rudainska, M. A,, 163, 178(109), 155, 156 Rupert, C. S., 6(67), 35 Rushizky, G. W., 43, 46, 47(63), 75, 92 (54), 131(244), 13G, 141 Ryan, F., 6(59, 60), 35 Rychlik, I., 66(160), 67(160), 68(160), 75(160), 51 S Sabatini, Y., 325(246), 349 Sadler, J. R., 328(257), 349 Sadron, C., 94(79), 106(176), 137, 139 Saez, F. A., 174(100), 156 Sagan, L., 151(35), 184 Sagik, B. P., 231 (95), 265 Samarina, 0. P., 228(61), 229(83), 230 (83), 233 (83), 254, 255, 262 (32, 35), 263(75), 269(68), 271(75), 273(68), 275(68, 75), 276(68), 277, 278(68), 279(68), 280(68, 75, 99), 281 (68), 292(32, 133), 293(32), 294(133, 1441, 295(35, 145), 296(146), 298(146), 303(163), 308(68, 1951, 316(99, 195), 317(68, 99, 2261, 313(99,

AUTHOR INDEX

420 195), 319(75), 321(146), 323(75,133), 326(75, 163), 327(133), 338(@), 342(306), 343, 344, 345, 346, 3477 351 Samejima, T., 62(125), 80, 9 4 ( f % 67, 68), l-3G Samis, H.F., 268(64), 344 Sampson, M., 161,154 Samuels, H.H., 306(181), 309(181), 347 Sanadi, D. R., 144(9), 162(9), 163(9), 168(9), 171,lS3,lSG Sandakhchiev, L. S., 56(104), 68(104), 7'9 Sandeen, G., 59(119), SO, 68, 135 Sander, C . , lOl(l53), 102(167), 103,139 Sanders, F.,367(94), 368(94), 371(94), 388(94), 39Y Sanger, F., 44,45,57,75, SO San Karanarayanan, K., 239(146), 256 Sarabhai, A. S., 26(159), SY,356(48), 39G Sarin, P.S., 63(146), 76,81, 94(71, 721,

Schulte, J., 219(2), 225(28), 226(28), 227(2, 28,36),253,254 Schuppe, N. G., 275(83), 280(83), 285(83),345 Schuster, F. L., 185 Schuster, H., 9(84), 21(84), 36, 66(157), 81

Schwarte, J. H., 357(62), 358(66), 397 Schwarte, N. M., 12(117), 36 Schweigcr, M., 46(59), 75 Schweitzer, M.P., 97,138 Scott, D. W., 354(16),395 Scott, J. F., 40(25), 43(25), 44(25), 46, 49(25, 76),53(87),7'7, 79 Scruggs, R. L., 103(162), 111(162), 112(162), 139 Seaman, E., 17(128), 3Y, 62(127), 80 Sedat, J. W., 7(78), 35, 261(11), 342 Seed, J., 228(53), 654 Seidel, H., 59,SO 137 Sarkar, P. K., 98(135), 99(141), 101(149), Sekeris, C. E., 278(95), 315(95), 345 Seraidmian, K., 43(41), YS, 91, 1% 105(149), 138 Setlow, R. B., 6(63), 21(148a, 1491, Sarnat, M., 307(187, NO), 347, 348 '31(63),35, 3Y Sasisekharan, V., 111(190), 122(190), 1% Setterfield, G., 264(51), 291(51), 344 Sato, K., 42,YB, 315(215), 348 Severtzov, A . N., 229(83), 230(83), Sato, T., 60(124), SO 233(83), 855 Saunders, M., 111(192), 139 Seymour, W.F.K., 6(56),35 Schachman, H. K., 85(13, 18,20,136 Shapiro, L.,355(42), 377(42), 378(42), Schachtschabel, D., 63(148), 81 379(122), 380(123), 381, 396, 398 Schadk, J. P., 209,218 Shapiro, R., 69(173a),81 Schaeffer, T., 17(127), 3Y Schemer, K.,264(43), 267(43), 269(43), Shatkin, A. J., 306(175), 308(175), 34Y 271(43), 275(43), 285(103), 286, Shemjakin, M. F., 339(297), 340(297), 651 287(103), 316(218), 317(218), Sheridan, J. W., 251(166), $5257, 269(70), 341(302), 343, 345, 348, 351 275(70), 279(70), 944 Schiff, J. A,, 149(28), 150(31), 151(35), Sheridan, W. F., 151(36),184 155,157(44, 461,184 229(76,79), 230(79), 233(79), Schildkraut, C . L., 98(127,128), 121(128), Sherrer, K., 234,254 131(247), 138, 141, 149(26), I S / , Shibuya, H., 394(150), 399 Schlessinger, D., 249(162),25Y Shimanouchi, T., 95(88, 89, go), 13Y Schmidt, G., 43(41), YS Shipp, W. S., 150(30), 159,184, 371(113), Schmitt, F.O., 209(32), 215 398 Schneider, W.C . , 162(59), 170, 155 Shlyk, A. A., 160,184 Scholl, D. A., 209,218 Schor, N., 193(16), 206(16), 217, Shreeve, W.W., 227(48), 264 Shugar, D., 91, lOO(222, 257), 115(218, 299(153), 300(153),347 222, 256, 257), 116(222), 120(209), Schramm, G., 385(137), 340 125(218), 127(218, 2221, 128(218), Schull, W.J., 2(5), 3(5), 33 136, 140, 141 Schulman, H.M., 309,348

42 1

AUTHOR INDEX

Shulmann, R. G., 97(119, 119:1), 1% Shy, G. M., 166(84),1S5 Sibatani, A., 262, 264(36), 265, 267(37, 59a), 268(60, 62), 301(60), 316(222), 343, 344, 34s

Sicurella, N. A., 21(144), 37 Siebert, G., 228(64), 254, 299(151), 346 Siegel, E. C . , 10 (low, 36 Sickevits, P., 264(53), 291(53), 293(53), 298(53), 301(53), 325(246), 344, 349 Sigler, P. B., 111(190), 116(202), 122(190), 124(202), 139, 140 Silver, D. G., 229(74), 234(74), 254 Simon, L., 355(41), 377(41), 378(41, 121), 396, 398

Simon, M. I., 69, 81 Simpson, L., 162(57), 164(57), 185 Sinclair, J., 162(60), 185 Singer, M. F., 41(57), 46(57), 7S, 131(244), 141 Singh, V. N., 267(57), 344 Sinha, N. K., 354(17), 395 Sinsheimer, R. L., 21(152), 37, 98(130), 121(130), 138, 345(5), 355(29, 36, 371, 357(59), 359(37), 360(82,83), 361(83), 363(86), 364(86), 365(86), 366(86), 309, 370(5), 376(116), 392(36, 861, 394(29, 59), 395, 396, 397, 398 Sirlin, J. L., 219, 221(12), 225(12), Z6(12), 229, 238(4), 253, 254, 323(241), 327(251), 349 SjGquist, I., 46(61), 49(61), 51(61), 53(61), 78 Sjoquist, J. A., 40(24), 77 Skaar, P. D., 10(101), 3G Skalka, A., 332(278), 350 Slonimski, P. P., 166(85), l68(85), 185 Smellie, R. M. S., 371(102), 385(102), 398

Smetana, K., 219(5), 253, 290(122), 291(127), 301(159), 346, 347 Smirnov, M. N., 229(83), 230(83), 233(83), 255, 263(75), 271(75), 275(75), 280(75), 319(75), 323(75), 326(75), 344 Smith, E. L., 2(15, 16), 8(83a), 25(15, 16), 26(15, 16), 27(16), 34, 36 Smith, I., 2(14), 25(14), 34, 222(15), 229(85), 230(85), 231(85), 233(85),

234(85), 235(85), 240(85), 253, 255, 342, 351 Smith, J. D., 41(30), 42(30), 55(102, 103), 56(102, 103), ?7, 79 Smith, J. S., 5(47), 34 Smilh, L., 338, 350 Smith, L. D., 329(259), 349 Smith, R. A., 385(141), 399 Smith-Keary, P. F., 12(115), 36 Smooth, J. S., 10(99), 12(99), 31(99), 32 (99), 36 So, A. G., 71(181), S2 Sobell, H., 111(191), 139 Sober, H. A., 46(60, 63), 47(63), 78, 92(54), 131(244), 136, 141 Soeiro, R., 341(303), 351 6611, D., 40(8), 77 Sonnahend, J., 371(111), 398 Sonnenberg, B. P., 331(273), 350 Sonnenbichler, J., 52(77b), 79 Sonnenborn, T. M., 3(28), 4(28), 8(28), 34, 146, 178, 1S3, 186 Sorm, F., 66(160), 67(160), 68(160), 75(160), 81 Sotelo, J. R., 174(102), 175(102), 186, 290(124), 346 Spach, G., 94(77), 137 Spahr, P. F., 41(30), 42(30), 43(38), 77 Spate, H. C., 113(197), 140 Spears, C. L., 6(66), 35 Spencer, M., 47, 59, 78, 80 Spcyer, J. F., 12(112, 112a), 36, 74(71), 7s

Spirgelman, S., 7(75, 82), 35, 220(7), 221, 222(13, 19), 223, 224(13), 226(13, 19, 20), 229(13), 231(95), 253, 255, 260(7), 306(177), 307(186, 1891, 308(189), 324, 342, 347, 348, 349, 354(20), 355(35, 43), 358(67), 359(76), 360(81), 367(93), 377(43), 381(43), 382(76, 125, 126, 127), 383, 387, 396, 397, 39s Spinelli, V., l0(98), 36 Spirin, A. S., 231(98), 236( 121, 122, 124), 247(158), 249(158), 255, 256, 257, 269(67), 281(100), 316(219, 220), 317, 344,345,348,349

Sporn, M. B., 278(93), 316(93), 334(290), 345, 550

AUTHOR INDEX

422 Sreevalsan, T., 371(115), 398 Srinivasan, P. R., 17(123), 18(123), 36, '228(68), 229(68), 231(68), 254, 271(77), 273(77), 344 Staehelin, M., 45, 46, 78 Staehelin, T., 236(125), 236(125), 255, 256, 317(225), 322(231), 349 Stahl, F. W., 98, 138, 164(72), 185 Stahl, R. C., 25(157), 37 Stanchev, B. D., 268(65), 278, 344 Stanley, W. M., Jr., 369 Starr, J. L., 43(36), 77 Stedman, E., 330,350 Steele, W. J., 231(96), 232(96), 233(96), 234(96), 255, 275(86), 290(122), 291(127), 301(158), 325(86), 1581, 333(280), 346, 346, 347, 350 Steffensen, D. M., 151(36), 184 Steigbigel, N. H., 6(66), 35 Steiner, R. F., 84(1), 91, 101(154), 107(182, 183), 113(193), 126(219), 130(238), 133(1), 134(254), 134, 136, 139, 140, 141 Steinert, G., 164(65), 184 Stenram, U., 237(133), 256 Stent, G., 299(154), 306(154, 180, 182), 309(182), 310(154, 180), 312(154), 313(154), 314(154), 320(154), 326(154), 347 Stenzel, K. H., 198, 218 Stephenson, M. L., 49(76), 53(87), 79 Stern, H., 150(33), 161(33, 541, 163, 164(65), 184, 185, 264(51), 291(51), 293(140), 344, 34G Stevenin, J., 195(37), 218 Stevens, B. J., 239(144, 148), 256 Stevens, C. L., 91(42), 101(42), 103(42), 107(42), lOS(421, 109(42), 112(42), 114(42), 115(42), 128(42), 136 Steward, M., 63(147), 75(147), 81 Stickler, N., 43(41), 78 Stirlin, J. L., 229, 254 Stodolsky, M., 307(187), 347 Stokes, A. R., 92(52), 95, 136, 137 Stone, G. E., 164(68), 168(68), 185 Store, J., 385(143), 399 Strack, H. B., 9(88), 20(88), 28(88), 36 Strauss, D. B., 84(1), 133(1), 135 Straum, J. H., Jr., 354(5), 355(29), 369, 370(5), 394(29), 3.95, 396

Strausser, F. F., 228(55), 254 Strelzoff, E., 307 (184), 347 Stretton, A. 0. W., 26(159, 160, 161), 37, 356(47,48), $96 Stubbe, H., 28(171), 38 Stubbe, W., 156, 184 Studier, F. W., 365(92), 370, 397 Sturtevant, J. M., 103(163), 111(163), 112(163), 131(163), 139 Subbaiah, V., 14(124), 17(124), 18(124), 36 Sueoka, N., 56(109), 79, 98(126), 138, 159(50), 160(50), 184, 222(14), 226(14), 229(75), 253, 264, 261(13), 343 Summers, D. F., 371(107), 398 Sundarlingham, M., 137 Suskind, R. G., 237(132), 256 Suyama, Y., 144(10), 150(10), 162(10), 163 Suzuki, H., 310(201), 348 Suzuki, N., 60(123), 60(123), 80 Svedberg, T., 85(17), 136 Swan, R. J., 99(139), 105(139), 133(251a), 138, 141 Swanson, C. P., 6(62), 35 Swartz, M. N., 5(51), 8(51), 20(51), 35 Swierkowski, M., 100(257), 115(257), 120(209), 140, 141 Swift, H. H., 150(32), 151(32), 158(49), 162(66), 164(32), 184, 186, 239(144), 266, 290(123), 333(285), 34G, 350 Sypherd, P. S., 252(174), 257 Szer, W., 100(222,257), 102(260), 115(218, 222, 256, 2571, 116(222), 120(209), 125(218), 127(218, 2221, 128(218), 140, 141 Szilard, L., 5(39, 40), 34 Szybalski, W., 121(2611, 141 T Tada, M., 40(21), 77 Takagi, Y., 310(201), 3.48 Takahagi, T., 262(37), 265(37), 267(37), 343 Takahashi, K., 43(43), 78 Takanami, M., 59(114), 62(138), 66(114, 161), 67, 80, 81, 247(154), 256, 298 (147), 346

423 423

AUTHOR INDEX

Takata, K., 262(25), 292(25), 294(25), 301(251, 343 Takemura, S., 49(73), 50(73), 56, 78, 79 Taketo, A., 394( 150), 399 Talwar, G. P., 300(155), 347 Tamaoki, T., 229(77, 861, 230(86, 91), 232, 233(86, 911, 235(85), 264, 256, 271(7.2), 275(72), 301, 302(160), 325, 344, 347 Tamaoki, T., 271(72), 275(72), 301, 302(160), 325,344, 347 Tamm, I., 384(129), 398 Tanaka, K., 48(69), 49(69), 78 Tandler, C. J., 327(252), 349 Tanford, C., 85(19), 86(19), 95, 97, 135, 231(97), 266 Tata, J. R., 189(8a), 217 Tatum, E. I,., 22(153), 37, 167(78), 185, 306(175), 308(175), 347 Taylor, A. L., 12(114), 36 Taylor, C., 333(280), 360 Taylor, J. H., 227(47), 264 Tecce, G., 62(139), 80 Teeter, E., 268(63), 344 Temin, H. M., 387(145), 399 Tener, G. M., 43, 52(81), 53(81), 72, 78, 79 Terskich, V. V., 271(76), 273(76), 275(76), 286(76), 288(76), 319(230), ‘320(230), 321(76), 325(76), 344,349 Tessman, I., 13(118, 1191, 21(118, 119),

Toccini-Valentini, G. p., 307 (187), 308(190), 34r, 34s Tomita, K,-I., 370(100), 398 Tomkins, G. M., 249(163), 267 Tomlinson, R. U.,43, 52(81), 53(81), 78, 7s

Tomoda, J., 267(59a), 344 Torres-Gallardo, J., 71 (1821, 76, 82 'I'rager, W., 163(63), 186 Trautner, T. A., 5(51), 8(51), 20(51), 36

Treffers, H. P., 10(98), 12(106), 36 Trujillo-Cen6z, o., 174(102), 175(102), 1SG

Truman, D. E. S., 6(57), 21(57), 36 Trnpin, J., 40(7), 74(7), 77 Tsanev, R., 276(87, 88), 288(112), 345 Ts'O, P. 0. P., 97(115), 101(153), 102(167), 103, 104(166), 138, 189 Tsuboi, M., 60(124), SO, 95, 137 Tsugita, A., 69, 81 Tsukada, K., 323(238), 349 Tsunakawa, S., 52(77b), 79 Tucker, M. D., 83(147), 75(147) 81 Tumanjan, V. G., 269(68), 273(68), 275(68), 276(68), 277(68), 278(68), '279(68), 280(68), 281(68), 308(68), 317(68), 338(88), 344 Tuppy, H., 169(87,88), 186 Turba, F., 261(91, 268(9), 278, 342 Tyndall, R. L., 268(63), 344

SG Tewari, K. K., 144(7), 162(7), 169(7), 183 Thang, M. N., 84(5), 136 Thannhauser, S. J., 43(41), 78 Theil, E. C., 6(55), 36 Thiebe, R., 56, 79 Thomas, R., 87,156 Tichonov, V. H., 285(104), 322(104), 338(104), 346 Timofeef-Ressovsky, N. W., 2(22), 28(166), 31(22), 34, 38 Timofceva, M. J., 275(82), 280(82), 345 Tinoco, I., 84(1), 94, 101(69), 103(69, 160), 133, 135, 136, 139, 141 Tissicres, A , , 62, 80, 314(213), 348 Toal, J. N., 92(54), 130 Tobey, R. .4., 363037, 88), 371, S.(I?

U Uchida, T., 43(43), 7s Uehara, K., 69, SI Uemura, I., 7(78), 36, 261(11), 348 likits, T., 64(153, 154), 65(154), 66(156), 69, 70(174), 75(154), 8f Ulbricht, T. L. V., 99(139), 105(139), 133(251a), 138, 1.41

V Valrntine, Valmtine, R. R. C., C., 354(24, 354(24, 25), 25), 357(58, 357(58, 60), 60), 395 395 Vdeva, Valeva, 1,. 1,. I., I.,276(89), 276(89), 346 345 J'dlee, l’nllee, B. B. L., L., 62(126), 62(126),SO 80 viln v i m Backer, Backer, H., H., 209(34), 209(34), 21s 218

AUTHOR INDDX

424 Van der Dehn, 169, 186 Van Holde, K. E., 85(20), 94(83, 841, 103(83), 106(84), 126(83), 136, 137 van Vunakis, H., 69,81 Van Wisselingh, C., 152(40), 184 Vassalli, P., 268(66a), 344 Vasilenko, S. I<.,91, 136 Vaughan, M. H., Jr., 147(23), 184 Veelmetter, W., 9(84), 21(84), 36 Venkov, P. V., 251(172), 26?, 276(89), 346 Venkstern, T. V., 49(77), 51(77), 53(77), 79, 91(47), 136 Verwoerd, D. W., 66(158), 81 Villalobos, J., 228(64), 264 Vincent, W. S., 219, 229(72), 236(129), 237(72), 263, 264, 266, 264, 291(49), 291 (128), 301 (128), 3.43, 346 Vinograd, J., 98, 138 Viiiuela, E., 354(1a), 357(55), 359(73), 371(73), 372(73), 373(73), 374(73), 375(73), 397 Vladimirzeva, E. A., 271(76), 273(76), !Z75(76), 286(76), 288(76), 321(76), 325(76), 344 Vlasenok, L. I., 160(52), 184 Voet, D., 91, 136 Vogel, H. J., 2(9), 25(9), 33 von Ehrenstein, G., 39(2), 40(6), ?6,77 von Vbich, H., 262(17), 343

W Wacker, A., 69, 81 Wada, A., 68, 81 Wagner, R. P., 4 (34), 34 Wallace, H., 221(12), 222(18), 223, 225(12), 226(12, 181, 228(69), 263, 264 Walters, C. P., 385(131), 398 Waltschewa, L. W., 268(66), 344 Wang, T.-Y., 292(134, 135, 136), 346 Ward, D. C., 130(227), 140 Warner, H. R., 12(111), 36 Warner, J. R., 235(118), 266, 341(303), 361

Warner, R. C., 106(177, 1811, 110(181), 113, 139, 306(181), 309(181), 347,

366(95), 367(95), 369(95), 371(% 114), 387(95), 389, 390(95), 397,398 Warr, J. R., 179(110), 186 Warshaw, M. M., 103(160), 139, 1.41 Watanabe, I., 62(138), 89, 95(88), 137, 354(14), 396 Watanabe, K., 371(106),398 Watanabe, Y., 371(106), 398 Waters, L., 229(84), 266 Watson, J. D., 2(23), 10(23), 18(23), 31(23), 34, 101(157), 122(157, 2 1 9 , 130,139, 140,231(93), 266 Weber, K., 354(3), 396 Webster, R. E., 57(116), 80, 357(53, 54), 358(53, 54), 396 Wedel, H., 354(24, 25), 396 Weigert, M. G., 356(46), 396 Weill, J. H., 63(145), 64, 74(152), 80, 81, 94(80), 101(80), 137 Weiler, E., 227 (38), 264 Weill, J., 85(6), 136 Weinberg, R., 31(1871, 38 Weinstein, I. B., 63(143), 76(143), 80 Weisblum, B., 39(2), 40(3), 76 Weiss, S. B., 121(212), 140, 305(164, 169), 306(173, 1871, 310, 313(211), 339(211, 296), 347, 348, 360, 360 (84), 387 (84), 397 Weissmann, C., 354(1, la, 18, 19), 355(41), 357(55), 358(79, 1181, 359(69, 70, 73, 74, 75, 77, 79), 360(69), ,361(70), 362(1), 364(91), 366(95), 367(95), 368(96), 369,369(95), 370(96, 101), 371(73, 95, 114), 372(73, 74, 751, ,373(73), 374(73), 375(73), 377(41, 118), 378(91, 121), 379(1, 70, 75, 118), 380(91), 381(75), 382(18, 771, 383(70, 77, 127a), 384(77), 385(77), 387(95), 388(70, 74), 389, 390(95), 392(75), 394(19), 396, 396, 397, 398 Welkie, G. W., 385(138), 399 Went, H. A,, 172(96b), 174(99a), 179(96b), 186 Wettstein, F. O., 231 (99), 266, 317(225), 322(231), 349 Wikler, M., 354(3), 396 Wildman, S. G., 385(136, 141), 399 Wilhelm, R. C., 355(44), 357(54), 358(54), 396

425

AUTHOR INDEX

Wilkie, D., 146, 183 Wilkins, M. H. F., 47(67), 59(67), Y8 Williams-Ashman, H. G., 189(8), 217, 312(208), 348 Wilson, C. W., 54, 55(102, 1031, 56(102, 1031, Y9 Wilson, H. R., 98(133), 13s Wilson, R. G., 385(132), 39s Winnick, T., 7(78), 35, 261(11), 342 Wintersberger, E., 169(87, 881, 185 Witkin, E. M., 6(68, 69), 21(144), 36, 37

Wittes, R. E., 3(28a), 8(28a), 34 Wittrnann-liebold, B., 354(21), 396 Witz, J., 95, 97, 101(158), 103(158), 137, 138, 139

Wojcik, S. J., 385(135), 399 Wolfenden, R., 53(50), Y8,79 Wolkenstein, M. V., 130(231), l 4 O Wollgiehn, R., 151(38), 184 Wood, W. B., 305(172), 34Y Woods, M. W., 182(112), 186 Woods, P. S., 229(70), 254, 323(234), 349 Woodward, D. O., 167, 185 Woody, P. I,., 31(181), 38 Woody-Karrer, P., 31(183, 184), 38 Work, T. S., 371(103), 385(103), 398 Wulff, V. J., 268(64), 344 Wyatt, G., 235(115), 266

Yasuzumi, G., 290(118), 346 Yermolaeva, L. P., 292 (131), 323(242), 338(131), 346, 349 Yoneda, H., 85(14), 135 Yoshida, A,, 2(8), 25(8), 28(8), 33 Yoshida. M., 69, 70(174), 81 Yoshikawa, M., 251(165), 257, 275, 280(79), 317(79), 345 Yoshikawa-Fukada, M., 251 (167, 168), 257, 275(80), 280(80), 317(80), 345 Yoshimasa, K., 95(89), 13Y Yotsuyanagi, Y., 151(37), 159, 184 Young, R. J., 52(80, 821, 79 Yu, C., 64, 64(151), 74,SI Z

Zacshau, H. G., 46(59), 48(68), 49(68), 50(68), 51(68), 52(77a, 77b), 53(68, 86), 56, 66(162), 68, 75, Y8,79, S1 Zajdela, F., 341(302), 351 Zalokar, M., 237(130), 956 Zalta, J. P., 385(139), 399 Zamccnik, P. C., 40(22), 49, 53(87), 62(126), 63(146), 64, 74, 76, 77, 79, 80, S l , 94,13Y Zamenhof, P. J., 12(108, log), 28(165), 29(176), 30(176), 36, 38 Zamenhof, S., 2(4, 6, 25, 261, 3(6, 30, 31, 32), 5(6, 25, 38, 46, 48), 6(25, Y 53, 54, 55, 58), 10(4, 100, 102), 12(100), 13(4), 14(124), 17(4, 25, 123, Yagi, K., 40(21), YY 124), 18(4, 25, 26, 123, 124), 19(25, Yamana, K., 262(36, 37, 38), 264(36), 26, 1381, 20(143), 21(25, 26, 31, 147, 265(36, 37, 38), 267(36, 37), 325(246), 150), 28(25, 26, 150, 165, 169), 343, 349 29(176), 30(176), 31(46, 100, 102, Yamamoto, O., 268(62), 344 185), 32(102), 33,34,36, 36,37,38 Yamamoto, S., 268(62), 344 Zamir, A., 7(74), 35, 45, 47(65), 49(65), Yamamoto, Y., 68(163), 81 51(65), 53(65), 59(65), 62(65), 78 Ynmane, T., 56(109), 79 Zbarsky, I. B., 262(28), 267(59), 288(28, Yanagishima, N., 166(77), I S 5 113), 290(28), 292(131), 294(144), Yang, J. T., 62(125), SO, 93(60), 94, 322(233), 323, 338(131, 2331, 343, 34.4, 98( 135), 99(141), 101(149), 105(149), 34.5, 346, 349 136, 138 Yankofsky, S . A., 222(19), 226(19, 20), Zillig, W., 63 66(158), 81, 305(166, 167), 315(167), S47 253, 306(177), 34Y Yanofsky, C., 4(36), 7(73), 8(36, 83), Zimm, B. H., 130(232, 235, 2391, 131, 136, 140,141 lO(73, 831, 12(107), 21(36, 73, 83), 26(36, 73, 83, 162, 163), 27(36, 73, Zimmer, K. G., 2(22), 31(22), 34 Zirnmerman, E., 261(91, 268(9), 278,342 83), 34, 35, 36, 3Y

426

AUTHOR INDEX

Ziinmerman, E. F., 231(100), 235(100),

Zollner, N., 43(41), 78 Zubay, G., 59(114), 62, 63(141), 66(114), 80, 229(70), 264, 331(273), 337(271), Zinder, N. D., 57(116), 80, 354(2a, 41, 360 3!55(4, 28, 30, 31, 33, 42, 45), 356(50, 52)7 5 4 ~56i 58, Zubkoff, P. L., 40(20), 46(20), 65(20), 60, 6 3 , 358(53, 54, 661, 359(31, 60), 70(20), 77 362(57’ 85a)’ 364(567)’ 365’ 376’ Zuckerh~ndl, E., 2(17), 25(17), 26(17), 377(42), 378(42, 52, 57), 379(56), 34 386, 394(30, 31, 501, 379(56). 386, 389, 394(30,31, 50), 395, 396,397 265

519

357(52a9

53j

57j

Subject Index A Anticodon sequence, transfer ribonucleic acid and, 48-52 B Brain cells, acidic proteins specific for, 197-199 biosynthesis of rapidly labeled ribonucleic acid in, 194-197

hereditary factors and, 151-157 labile or “metabolic,” 160-161 extranuclear, general considerations, 143-146 mi t ochondrial, evidence for, 161-165 genetic significance, 168-172 hereditary factors and, 165-168 Diffusion, molecular weight and, 85-87

E C Centriole, deoxyribonucleic acid of, 172-179 Chloroplast, deoxyribonucleic acid, evidence for, 146-151 genetic significance, 157-160 hereditary factors and, 151-157 labile or “metabolic,” 160-161 Chromosomes, messenger ribonucleic acid synthesis in, 30%315 Circular dichroism, polynucleotide, 92-94 Cistrons, properties, ribosomal nucleic acid and, 221-224 Cytoplasm, appearance of ribosomes in, 239-249 Cytosine, associations with hypoxanthine, 116-119 D

Deoxynurleoprotein, ribonucleic acid of, 299-300 Deoxyribonucleic acid, centriole, differentiation of procentriole, 174-176 evidence for, 176177 function of proccntriole, 172-174 genetic aspects of, 177-179 chloroplast, evidence for, 146-151 genetic significance, 157-160

Electron mirroscopy, polynucleotides, 97 Electron spin resonance, polynucleotides, 97 Equilibrium density gradient centrifugation, polynucleotides, 98

G Genes, nuclear and cytoplasmic, interactions of, 181-182 ribosomal nucleic acid and, 221-228 Glia cells, methods of analysis, 191-193 ribonucleic acid, possible transfer of, 20s210 H Helix-coil transitions, theory and practice of, 130-131 Homopolynucleotides, 98-104 Hypoxanthine, associations, with cytosine, 116-119 I Infrared spectroscopy, polynucleotides, 94-95 1

I,earning, adenine-uracil rich ribonucleic acid and, 202-208 Light scattering, polynucleotides, 97 427

428

SUBJECT INDEX

M

Memory, working hypothesis, 213-217 Messenger ribonucleic acid, biosynthesis in chromosomes, 308-315 transport of, 315-322 Mitochondria, deoxyribonucleic acid, evidence for, 161-165 genetic significance, 168-172 hereditary factors and, 165-168 Mutability, definitions, 2-6 development and, 28 inheritance, 10-32 modes of, 10-27 regulation of changes in, 29-32 Mutation ( s ) , definition of, 3-6 detection of, 6-10 protection against, 28 spontaneous and induced, 2 N Nerve cells, methods of analysis, 191-193 ribonucleic acid, possible transfer of, 208-210 Nuclear magnetic resonance, polynucleotides, 97 Nuclear ribonucleic acid, biosynthesis and transport of, 304-327 fractionation, main approaches,261-264 localization and mechanism of biosynthesis, 304-308 phenol fractionation and characteristics of, chromosomal, 277-286 methods, 264-269 molecular classes of newly formed, 269-276 nonchromosomal, 286-288 terminology and, 260 Nuclear sap, ribonucleoproteins of, 291-299 Nucleoid, constitution of, 180 cytoplasmic, role in inheritance, 179-182 redundancy and genetic analysis, 179-181

Nucleolar organizer, ribosomal nucleic acid and, 220-221 Nucleolus, ribonucleic acid of, 300-303 ribosomal nucleic acid synthesis and, 228-229 ultrastructure, ribosomal nucleic acid and, 238-239 Nucleonemata, ribonucleic acid of, 300-303 Nucleotides, arrangement in transfer ribonucleic acid in, 43-59 terminal sequence in transfer ribonucleic acid, 52-59

0 Oligonucleotides, transfer ribonucleic acid aminoacylation and, 71-76 Optical rotatory dispersion, polynucleotide, 92-94 Organelle(s), nucleoid redundancy and, 180-181 P

Parkinson’s disease, ribonucleic acid in, 210-213 Phages, see Ribonucleic acid phages Phosphate residues, polynucleotide, 119-120 Polyadenylic acid, association with polyinosinic acid, 116 complexes with polyuridylic acid, 106-108 thermodynamics of interaction, 111-113 properties of, 101-103 Polyadenylic. Polyuridylic acid, base pairing in, 111 stability, at acid pH, 110-111 at alkaline pH, 110 thermal dissociation of, 108-110 Polycytidylic acid, complexes with polyguanylic acid, 104-106 properties of, 103-104

429

SUBJECT INDEX

Polyguanylic. acid, complexes with polycytidylic acid, 104-106 properties of, 98-99 Polyinosinic acid, associations with polyadenylic acid, 116 properties of, 99-100 Polynucleotide (s), alternating copolymers of, 113-115 analog-containing, 125-130 circular dichroism, 92-94 complexes, 104-109 displacement reactions of, 123-125 electron microscopy of, 97 equilibrium density gradient, centrifugation, 98 helices, parallelism or antiparallelism of, 122 helix-coil transitions, theory and practice of, 130-131 infrared spectroscopy of, 94-95 molecular weight, 85-87 nuclear magnetic resonance and electron spin resonance of, 97 optical rotatory dispersion, 92-94 physical chemistry, techniques for investigation of, 85-98 preparation of, 84-85 reversibility of, 122-123 role of sugar phosphate backbone, 119-122 structure, factors governing, 131-134 thermal stability, 87 . titrimetry of, 94 ultraviolet absorption spectra, 87-92 X-ray diffraction and light scattering of, 95-97 Polyribosomes, ribosomal nucleic acid and, 245-249 Polyuridylic acid, complexes with polyadenylic acid, 106-108 thermodynamics of interaction, 111-1 13 properties of, 101 Procentriole, deoxyribonucleic acid, evidence for, 176-177 differentiation of, 174-176

function of, 172-174 genetic aspects of, 177-179 Protein(s), acidic, specific for brain, 197-199 ribosomal, synthesis of, 235-238 R Ribonucleic acid, adenine-uracil rich, learning and, 202-208 base ratios, during chemical induction of synthesis, 199-202 during physiological stimulation, 199 biosynthesis, regulation of, 32S340 cellular, main classes of, 260-261, 269-276 droxynucleoprotein fraction, 299-300 determination of “minus” strand, 392 determination of “plus” strand, 392-394 dou ble-stranded, analysis of, 388-392 identification of virus-specific, 387-388 nature, subnuclear structures and, 303-304 neural function and, historical, 187-190 problem discussed, 190-191 nuclear, see Nuclear ribonucleic acid nucleoli and nucleonemata, 300-303 Parkinson’s disease and, 210-213 possible transfer between glia and neuron, 208-210 rapidly labeled, biosynthesis in brain cells, 194-197 transfer, see Transfer ribonucleic acid viral, 35S359 base sequence homology between, 395 enzymatic studies of replication, 377-386 identification and analysis of, 387-395 messenger function of, 357-358 replication in vivo, 359-377 Ribonucleic acid phages, general properties and biology of, 354-355 mutants of, 355-357

430

SUBJECT INDEX

Ribonucleoproteins, nurlear sap, 29-299 Ribosomal ribonucleic acid, genes coding for, properties of cistrons, 221-224 regulation of transcription, 224-228 relation to nucleolar organizer, 22&22l subsequent events in formation, cleavage of precursor, 233-235 relation to nucleolar ultrastructure, 238-239 synthesis of ribosomal protein, 235-238 synthesis and transport of, 322-326 synthesis of precursor, association with other cell components, 232-233 evidence that it is 45s component, 229-231 nucleolus as site of, 228-229 properties of 45s component, 231-232 Ribosomes, appearance in cytoplasm, distinctive properties, 243-245 polyribosomes and, 245-249 subunits, 239-243 turnover of, 249 synthesis of, addendum, 251-252 synopsis, 249-251

T

Thermal stability, polynucleotides, 87 Titrimetry, polynucleotides, 94 Transcription, regulation, ribosomal nucleic acid and, 224-228 Transfer ribonucleic acid, aminoacylation, oligonucleotides and, 71-76 anticodon sequence of, 48-52 base composition of, 40-43 nucleotide arrangement in, 43-59 nucleotide distribution in, 43-48 specific sites, modification of, 63-71 studies on functional sites, 63-76 synthesis and transport of, 327 terminal nucleotide sequence of, 52-59 three-dimensional structure of , 59-63 U

Ultracentrifugation, molecular weight determination by, 85-87 Ultraviolet absorption spectra, polynucleotides, 87-92 V

S Subnuclear structures, fractionation, 288-291 ribonucleic acid of 288-304 Sugar, polynucleotide, 120-122

Viscosimetry, molecular weight and, 85-87 X

X-ray diffraction, polynucleotides, 95-96